Biological Control of Insect Pests: Southeast Asian Prospects - EcoPort
Biological Control of Insect Pests: Southeast Asian Prospects - EcoPort
Biological Control of Insect Pests: Southeast Asian Prospects - EcoPort
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<strong>Biological</strong> <strong>Control</strong><br />
<strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>:<br />
<strong>Southeast</strong> <strong>Asian</strong><br />
<strong>Prospects</strong><br />
D.F. Waterhouse
<strong>Biological</strong> <strong>Control</strong> <strong>of</strong><br />
<strong>Insect</strong> <strong>Pests</strong>:<br />
<strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
D.F. Waterhouse<br />
(ACIAR Consultant in Plant Protection)<br />
Australian Centre for International Agricultural Research<br />
Canberra<br />
1998
ii<br />
The Australian Centre for International Agricultural Research (ACIAR) was<br />
established in June 1982 by an Act <strong>of</strong> the Australian Parliament. Its primary<br />
mandate is to help identify agricultural problems in developing countries and to<br />
commission collaborative research between Australian and developing country<br />
researchers in fields where Australia has special competence.<br />
Where trade names are used this constitutes neither endorsement <strong>of</strong> nor<br />
discrimination against any product by the Centre.<br />
ACIAR MONOGRAPH SERIES<br />
This peer-reviewed series contains the results <strong>of</strong> original research supported by<br />
ACIAR, or deemed relevant to ACIARÕs research objectives. The series is<br />
distributed internationally, with an emphasis on the Third World<br />
©Australian Centre for International Agricultural Research<br />
GPO Box 1571, Canberra, ACT 2601.<br />
Waterhouse, D.F. 1998, <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong>.<br />
ACIAR Monograph No. 51, 548 pp + viii, 1 fig. 16 maps.<br />
ISBN 1 86320 221 8<br />
Design and layout by Arawang Communication Group, Canberra<br />
Cover: Nezara viridula adult, egg rafts and hatching nymphs.<br />
Printed by Brown Prior Anderson, Melbourne
Contents<br />
Foreword vii<br />
1 Abstract 1<br />
2 Estimation <strong>of</strong> biological control prospects 2<br />
3 Introduction 3<br />
4 Target insect pests 9<br />
4.1 Agrius convolvuli<br />
9<br />
4.2<br />
4.3<br />
4.4<br />
Rating 10<br />
Origin 10<br />
Distribution 10<br />
Biology 10<br />
Host plants 11<br />
Damage 11<br />
Natural enemies 11<br />
Attempts at classical biological control 14<br />
Discussion 16<br />
Anomis flava<br />
17<br />
Rating 18<br />
Origin 18<br />
Distribution 18<br />
Biology 18<br />
Host plants 19<br />
Damage 19<br />
Natural enemies 21<br />
Attempts at classical biological control 27<br />
Major natural enemies 31<br />
Discussion 31<br />
Aphis craccivora<br />
33<br />
Rating 34<br />
Origin 34<br />
Distribution 34<br />
Biology 34<br />
Host plants 35<br />
Damage 35<br />
Natural enemies 36<br />
Comments 44<br />
Aphis gossypii<br />
45<br />
Rating 46<br />
Origin 46<br />
Distribution 46<br />
Biology 46<br />
Host plants 47<br />
Damage 47<br />
iii
iv<br />
4.5<br />
4.6<br />
4.7<br />
4.8<br />
4.9<br />
Natural enemies 48<br />
Attempts at biological control 60<br />
The major parasitoid species 70<br />
An aphid-specific predator 79<br />
Comments 80<br />
Cosmopolites sordidus<br />
85<br />
Rating 86<br />
Origin 86<br />
Distribution 86<br />
Biology 87<br />
Damage 88<br />
Host plants 89<br />
Natural enemies 89<br />
Attempts at biological control 95<br />
Biology <strong>of</strong> main natural enemies 103<br />
Comments 104<br />
Deanolis sublimbalis<br />
105<br />
Synonyms 106<br />
Rating 106<br />
Origin 106<br />
Distribution 106<br />
Biology 107<br />
Host plants 108<br />
Damage 109<br />
Natural enemies 111<br />
Comment 112<br />
Diaphorina citri<br />
113<br />
Rating 114<br />
Origin 114<br />
Distribution 114<br />
Biology 115<br />
Host plants 116<br />
Damage 116<br />
Natural enemies 120<br />
Attempts at biological control 127<br />
Major natural enemies 131<br />
Comments 133<br />
Dysdercus cingulatus<br />
135<br />
Rating 136<br />
Origin 136<br />
Distribution 136<br />
Biology 136<br />
Host plants 137<br />
Damage 137<br />
Natural enemies 137<br />
Comment 138<br />
Dysmicoccus brevipes<br />
141<br />
Rating 142<br />
Origin 142<br />
Distribution 142<br />
Taxonomy 142
4.10<br />
4.11<br />
4.12<br />
4.13<br />
Biology 143<br />
Hosts 144<br />
Damage 144<br />
Natural enemies 145<br />
Attempts at biological control 145<br />
Major natural enemies 155<br />
Comments 156<br />
Hypothenemus hampei<br />
157<br />
Rating 158<br />
Origin 158<br />
Distribution 158<br />
Biology 159<br />
Host plants 163<br />
Damage 164<br />
Natural enemies 166<br />
Attempts at biological control 170<br />
Major parasite species 176<br />
Comments 180<br />
Leucinodes orbonalis<br />
185<br />
Rating 186<br />
Origin 186<br />
Distribution 186<br />
Biology 187<br />
Host plants 187<br />
Damage 189<br />
Natural enemies 189<br />
Attempts at biological control 192<br />
Major natural enemies 192<br />
Comments 194<br />
Nezara viridula<br />
197<br />
Rating 198<br />
Origin 198<br />
Distribution 198<br />
Biology 199<br />
Damage 199<br />
Natural enemies 200<br />
The role <strong>of</strong> pheromones and other chemical secretions 208<br />
Attempts at biological control 209<br />
Biology <strong>of</strong> the major species 228<br />
Comments 233<br />
Ophiomyia phaseoli<br />
235<br />
Rating 236<br />
Origin 236<br />
Distribution 236<br />
Biology 237<br />
Host plants 237<br />
Damage 238<br />
Natural enemies 239<br />
Attempts at biological control 249<br />
The more important parasitoids 251<br />
Comment 255<br />
v
vi<br />
4.14<br />
4.15<br />
4.16<br />
Phyllocnistis citrella<br />
257<br />
Rating 258<br />
Origin 258<br />
Distribution 258<br />
Biology 259<br />
Host plants 262<br />
Damage 263<br />
Natural enemies 264<br />
Attempts at biological control 279<br />
Major natural enemies 282<br />
Comment 285<br />
Planococcus citri<br />
287<br />
Rating 288<br />
Origin 288<br />
Distribution 288<br />
Biology 289<br />
Host plants 289<br />
Damage 290<br />
Natural enemies 290<br />
Attempts at biological control 296<br />
Biology <strong>of</strong> important natural enemies 311<br />
Comments 315<br />
Trichoplusia ni<br />
317<br />
Rating 318<br />
Origin 318<br />
Distribution 318<br />
Biology 318<br />
Host plants 319<br />
Damage 320<br />
Natural enemies 320<br />
Introductions for biological control <strong>of</strong> T. ni<br />
334<br />
Major parasitoid species 340<br />
Comment 345<br />
5 References 349<br />
6 Index <strong>of</strong> scientific names <strong>of</strong> insects 477<br />
7 General index 531
Foreword<br />
Since its inception in 1982, ACIAR has been a very strong supporter<br />
<strong>of</strong> classical biological control as a key element in the management <strong>of</strong><br />
exotic arthropod and weed pests. When practiced with appropriate<br />
safeguards, it <strong>of</strong>ten provides a sustainable and environmentally<br />
friendly alternative to the growing use <strong>of</strong> pesticides, particularly<br />
when integrated, if necessary, with the use <strong>of</strong> resistant plant varieties<br />
and cultural controls.<br />
Classical biological control has been very successful in regions <strong>of</strong><br />
the world (e.g. Australia, California, New Zealand, Oceania) where a<br />
large number <strong>of</strong> the major insect pests and weeds are exotic. This<br />
situation applies to a far lesser extent to <strong>Southeast</strong> Asia but, in a<br />
recent survey commissioned by ACIAR, Waterhouse (1993b)<br />
identified 40 major arthropod pests that merited evaluation as<br />
possible targets for biological control. Not all <strong>of</strong> these (e.g. the<br />
indigenous fruit flies) are attractive targets, but some at least are.<br />
The present volume is a companion to <strong>Biological</strong> <strong>Control</strong> <strong>of</strong><br />
Weeds: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong> (Waterhouse 1994). It<br />
summarises what is known about the natural enemies (principally the<br />
parasitoids) <strong>of</strong> the major exotic insect pests and indicates prospects<br />
for their biological control. The aim has been to facilitate, for<br />
countries <strong>of</strong> the region, the selection <strong>of</strong> promising individual, or<br />
collaborative, priority insect pest targets. This should also provide<br />
donor agencies with a readily accessible overview <strong>of</strong> the regionÕs<br />
major exotic insect pest problems and with an evaluation, where<br />
possible, <strong>of</strong> prospects for their amelioration by introduction <strong>of</strong><br />
natural enemies. This should assist in the selection, for support, <strong>of</strong><br />
projects that are best suited to their individual terms <strong>of</strong> reference.<br />
R. Clements<br />
Director<br />
Australian Centre for International<br />
Agricultural Research<br />
vii
1 Abstract<br />
Introduction 1<br />
<strong>Biological</strong> control programs have been mounted in some region(s) <strong>of</strong> the<br />
world against 13 <strong>of</strong> the 16 dossier pests and substantial or partial success has<br />
been achieved in one or more countries for 8. On the basis <strong>of</strong> available<br />
information there are good to excellent prospects for reducing, in at least<br />
some parts <strong>of</strong> the region, the damage caused by the following: Leucinodes<br />
orbonalis,<br />
Nezara viridula,<br />
Ophiomyia phaseoli and Planococcus citri.<br />
There are also good reasons for believing that there will prove to be valuable<br />
natural enemies for the following: Agrius convolvuli, Anomis flava,<br />
Aphis<br />
craccivora,<br />
Aphis gossypii,<br />
Diaphorina citri,<br />
Dysmicoccus brevipes,<br />
Hypothenemus hampei,<br />
Phyllocnistis citrella and Trichoplusia ni.<br />
There<br />
seems to be little prospect for classical biological control <strong>of</strong> Dysdercus<br />
cingulatus, too little is known about Deanolis sublimbalis and the prospects<br />
for control <strong>of</strong> Cosmopolites sordidus are unclear, although its lack <strong>of</strong> pest<br />
status in Myanmar is puzzling.
2 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
2 Estimation <strong>of</strong> biological control<br />
prospects<br />
<strong>Insect</strong> Rating Family Any Attractiveness<br />
biological as a target in<br />
control<br />
successes<br />
SE Asia<br />
Agrius convolvuli<br />
7 Sphingidae yes medium<br />
Anomis flava<br />
10 Noctuidae yes low to medium<br />
Aphis craccivora<br />
15 Aphididae ? medium<br />
Aphis gossypii<br />
19 Aphididae yes medium<br />
Cosmopolites sordidus 13 Curculionidae ? uncertain<br />
Deanolis sublimbalis 3 Pyralidae no uncertain<br />
Diaphorina citri<br />
8 Psyllidae yes medium<br />
Dysdercus cingulatus 11 Pyrrhocoridae no v. low<br />
Dysmicoccus brevipes 10 Pseudococcidae yes medium<br />
Hypothenemus hampeii 12 Scolytidae yes medium<br />
Leucinodes orbonalis 15 Pyralidae no medium to high<br />
Nezara viridula<br />
10 Pentatomidae yes high<br />
Ophiomyia phaseoli 14 Agromyzidae yes high<br />
Phyllocnistis citrella 16 Phyllocnistidae yes medium<br />
Planococcus citri<br />
7 Pseudococcidae yes high<br />
Trichoplusia ni<br />
7 Noctuidae yes medium
3 Introduction<br />
Introduction 3<br />
Waterhouse (1993b) published information, collected from agricultural and<br />
weed experts in the 10 countries <strong>of</strong> <strong>Southeast</strong> Asia, on the distribution and<br />
importance <strong>of</strong> their major arthropod pests in agriculture. Ratings were<br />
supplied on the basis <strong>of</strong> a very simple system<br />
+++ very widespread and very important<br />
++ widespread and important<br />
+ important only locally<br />
P present, but not an important pest<br />
The advantages and limitations <strong>of</strong> this system were discussed by<br />
Waterhouse (1993b). Of 160 insect and mite pests nominated as important in<br />
<strong>Southeast</strong> Asia, a subset <strong>of</strong> 47 was rated as particularly so.<br />
The aim <strong>of</strong> the present work has been to summarise information relevant<br />
to the prospects for classical biological control <strong>of</strong> the most important <strong>of</strong><br />
those <strong>of</strong> this subset <strong>of</strong> 47 that are thought to have evolved outside <strong>Southeast</strong><br />
Asia. The assumption is that many <strong>of</strong> these have been introduced without<br />
some (sometimes without any) <strong>of</strong> the natural enemies that help to control<br />
them where they evolved. The chances are very much lower for arthropod<br />
pests that evolved in <strong>Southeast</strong> Asia <strong>of</strong> introducing effective, sufficiently<br />
host-specific, organisms from outside the region. On the other hand, there is<br />
reason to believe that some parasitoids <strong>of</strong> pests that are thought to have<br />
arisen in, or adjacent to, the Indian subcontinent may not yet occur<br />
throughout the eastern region <strong>of</strong> <strong>Southeast</strong> Asia and several such pests are<br />
dealt with.<br />
In regional considerations <strong>of</strong> this sort, it is to be expected that not all <strong>of</strong><br />
the top 20, or even the top 10, <strong>of</strong> any one countryÕs arthropod pests will<br />
necessarily be included. Indeed, at least some <strong>of</strong> those omitted might well<br />
merit the production <strong>of</strong> additional dossiers if they are <strong>of</strong> such importance<br />
locally that a biological control program might be justified. ACIAR would<br />
be interested to learn <strong>of</strong> pests that might be considered in this category.<br />
The summary accounts presented are designed to enable a rapid review<br />
to be made <strong>of</strong> (i) the main characteristics <strong>of</strong> the principal insect pests <strong>of</strong><br />
agriculture that are believed to be exotic to part or all <strong>of</strong> <strong>Southeast</strong> Asia, (ii)<br />
what is known <strong>of</strong> their enemies, particularly those that have high or<br />
moderate levels <strong>of</strong> host specificity and (iii) what the prospects appear to be<br />
for reducing their pest status by classical biological control.
4 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
In most instances four databases (and particularly CABI) were searched<br />
for relevant information:<br />
AGRICOLA (Bibliography <strong>of</strong> Agriculture) 1970+<br />
BIOSIS (<strong>Biological</strong> Abstracts) 1989+<br />
CABI (CAB International) 1972+<br />
DIALOG (<strong>Biological</strong> Abstracts) 1969+<br />
In addition, in many instances abstracting journals and other published<br />
sources prior to the above commencement dates were also searched.<br />
Furthermore, useful information was also obtained from other references<br />
and from unpublished records. Nevertheless, in many cases the search<br />
cannot be described as exhaustive. Even more relevant than attempting an<br />
exhaustive search would be a fresh, detailed, field survey targeted on the pest<br />
in the region where it is causing problems. This is in order to determine what<br />
natural enemies are already present and, in particular, whether any <strong>of</strong> the<br />
organisms that might be considered for introduction are already present.<br />
The species dealt with are drawn from tables 4 and 5 <strong>of</strong> ÔThe Major<br />
Arthropod <strong>Pests</strong> and Weeds <strong>of</strong> Agriculture in <strong>Southeast</strong> Asia: Distribution,<br />
Importance and OriginÕ (Waterhouse 1993b). It is quite possible that<br />
additional arthropod pests rating highly in these tables will prove to be exotic<br />
to <strong>Southeast</strong> Asia (or significant parts <strong>of</strong> it) and, alternatively, that some<br />
considered to be exotic will, on further evidence, be shown to have evolved<br />
in the region. The ratings <strong>of</strong> the pests in the Pacific and Southern China<br />
included at the beginning <strong>of</strong> each dossier are based on information in<br />
Waterhouse (1997) and Li et al. (1997).<br />
The natural enemies most commonly selected against insect pests in<br />
modern classical biological practice are specific or relatively specific<br />
parasitoids. Although predators also clearly play an important role in<br />
reducing pest numbers (and have achieved considerable successes against<br />
scale insects and mealybugs) the majority <strong>of</strong> predators attack a wide<br />
spectrum <strong>of</strong> hosts. National authorities responsible for approving the<br />
introduction <strong>of</strong> biological control agents are becoming increasingly<br />
reluctant to do so for natural enemies that may possibly have adverse effects<br />
on non-target species <strong>of</strong> environmental significance. For this reason far more<br />
emphasis has been placed in the dossiers on parasitoids than on predators.<br />
There appears to be a widespread view that, when biological control<br />
alone results in a spectacular reduction in pest populations (as it <strong>of</strong>ten does)<br />
it is very worthwhile, but a lesser reduction is <strong>of</strong> little or no value. Nothing<br />
can be further from the truth, since far lower levels can have a major impact<br />
when integrated with other means <strong>of</strong> pest control. This applies particularly to
Figure 1.<br />
Introduction 5<br />
integration with the use <strong>of</strong> plant varieties that are partially resistant to the<br />
pest (Waterhouse 1993a).<br />
Plant resistance serves to decrease numbers, in particular by lowering<br />
reproductive rate and slowing growth rate. Resistance can be brought about<br />
inter alia by alteration <strong>of</strong> the physical characteristics (e.g. hairiness, cuticle<br />
thickness) <strong>of</strong> the plant and/or its chemical composition. If, as usually occurs,<br />
parasitoids and predators are not affected to an equal extent, an improved<br />
ratio <strong>of</strong> natural enemy to the pest will result and the impact <strong>of</strong> biological<br />
control will be increased. This was pointed out many years ago (van Emden<br />
1966; van Emden and Wearing 1965) and is well illustrated by glasshouse<br />
tests with the aphis Schizaphis graminum on susceptible and resistant barley<br />
and sorghum varieties and the parasitoid Lysiphlebus testaceipes (Starks et<br />
al. 1972). If it is assumed, as in the illustrative example in Figure 1, that the<br />
economic injury level is 100 aphids per plant, then neither the resistant<br />
variety alone, nor the parasitoid alone will prevent the injury level being<br />
exceeded, whereas the combination <strong>of</strong> resistance and parasitoids achieves<br />
this by a wide margin. As another example, biological control <strong>of</strong> Myzus<br />
persicae with Aphidius matricariae was only effective on chrysanthemums<br />
if the variety involved was partly aphid resistant (Wyatt 1970).<br />
300<br />
200<br />
100<br />
0<br />
0<br />
1 2<br />
Weeks<br />
3 4<br />
Susceptible<br />
No parasitoid<br />
Resistant<br />
Susceptible<br />
Parasitoid<br />
present<br />
Resistant<br />
Population growth <strong>of</strong> Schizaphis graminum on susceptible<br />
and partly resistant barley in the presence and absence <strong>of</strong> the<br />
parasitoid Lysiphlebus testiceipes.
6 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Efforts to achieve pest control by high levels <strong>of</strong> plant resistance alone<br />
may prove counterproductive if significant energy or other resources are<br />
diverted by the plant, since they cannot then be used for growth or<br />
reproduction. Thus, van Emden (1991) quotes data on 31 pigeon pea<br />
varieties screened at the International Crops Research Institute for the Semi<br />
Arid Tropics for insect pod damage. These data predicted a 31% yield loss<br />
for 90% resistance to insects. To accept a loss <strong>of</strong> this order is surely an<br />
unacceptable ÔsolutionÕ to the problem, particularly when even a low level <strong>of</strong><br />
natural enemy attack combined with moderate plant resistance is likely to<br />
achieve a far better yield. However, the interaction <strong>of</strong> resistance and natural<br />
enemies may not be a simple one, as pointed out by Wellings and Ward<br />
(1994) and such interactions urgently deserve further study. Nevertheless,<br />
the fact remains that, when integrated appropriately with plant resistance and<br />
other measures, even comparatively low levels <strong>of</strong> attack by natural enemies<br />
can lead to disproportionately large improvements in pest control.<br />
Although the major focus <strong>of</strong> the dossiers has been on the applicability <strong>of</strong><br />
the information to biological control in <strong>Southeast</strong> Asia, much has far wider<br />
applicability. In particular, a great deal is relevant to classical biological<br />
control in the oceanic Pacific which, until the past few decades, has received<br />
almost all its important insect pests from <strong>Southeast</strong> Asia. A brief tabulation<br />
<strong>of</strong> the distribution and importance <strong>of</strong> each pest in the Pacific is, therefore,<br />
given at the beginning <strong>of</strong> each dossier. The key to Pacific Country<br />
abbreviations is: Fr P, French Polynesia; FSM, Federated States <strong>of</strong><br />
Micronesia; Kiri, Kiribati; Mar Is, Marshall Islands; N Cal, New Caledonia;<br />
PNG, Papua New Guinea; A Sam, American Samoa; Sam, Western Samoa;<br />
Sol Is, Solomon Islands; Tok, Tokelau; Tong, Tonga; Tuv, Tuvalu; Van,<br />
Vanuatu; W&F, Wallis and Futuna. The key to <strong>Southeast</strong> <strong>Asian</strong> countries is:<br />
Myan, Myanmar (Burma); Thai, Thailand; Laos; Camb, Cambodia; Viet,<br />
Vietnam; Msia, Malaysia; Sing, Singapore; Brun, Brunei; Indo, Indonesia;<br />
Phil, Philippines.<br />
In any biological control program it is essential that appropriate<br />
procedures are adopted in relation to the selection <strong>of</strong> suitably host-specific<br />
natural enemies, the gaining <strong>of</strong> approval for introduction and release from<br />
the national authorities and safe procedures for eliminating unwanted fellow<br />
travellers. Simple Guidelines for biological control projects in the Pacific<br />
(Waterhouse 1991) are available from the South Pacific Commission,<br />
Noumea and FAO has a Draft Code <strong>of</strong> Conduct for the Import and Release <strong>of</strong><br />
<strong>Biological</strong> <strong>Control</strong> Agents (1993) .<br />
Because there is a considerable lack <strong>of</strong> uniformity in the names applied<br />
to many <strong>of</strong> the insects involved, a separate index is included listing the<br />
preferred scientific names. These have been used in the text, replacing where
Introduction 7<br />
necessary those used by the authors quoted. Where the name <strong>of</strong> an insect<br />
used in a publication is no longer preferred by taxonomists, the superseded<br />
name, x, is shown thus (= x), but this usage is not intended to convey any<br />
other taxonomic message. Indeed, the superseded name may still be valid,<br />
but simply not applicable to the particular species referred to by the author.<br />
I am most grateful for assistance from many colleagues during the<br />
preparation <strong>of</strong> this book. It is not possible to name them all, but special<br />
thanks are due to a number <strong>of</strong> CSIRO colleagues, in particular to Dr K.R.<br />
Norris for editorial assistance, Dr M. Carver for valuable advice on the Aphis<br />
dossiers, J. Prance for bibliographic assistance and to several taxonomists,<br />
including Dr M. Carver (Hemiptera), Dr P. Cranston (Diptera), E.D.<br />
Edwards (Lepidoptera), Dr I.D. Naumann (Hymenoptera) and T. Weir<br />
(Coleoptera). Others who have provided valuable information include D.<br />
Smith (Queensland Department <strong>of</strong> Primary Industries), Dr P. Cochereau<br />
(ORSTOM, Noumea) and Dr C. Klein Koch (Chile).<br />
Continuing warm support has been provided by Dr P. Ferrar, Research<br />
Program Coordinator, Crop Sciences, ACIAR, Canberra.<br />
It is again a pleasure to acknowledge, warmly, the expert assistance <strong>of</strong><br />
Mrs Audra Johnstone in converting my manuscripts into presentable form.<br />
It would certainly not have been possible to continue with these<br />
biological control activities long into retirement without the unfailing<br />
support, encouragement and forbearance <strong>of</strong> my wife, to whom my very<br />
special thanks are due.
4 Target insect pests<br />
4.1 Agrius convolvuli<br />
India<br />
20°<br />
Myanmar<br />
P Laos<br />
P<br />
0°<br />
20°<br />
China<br />
++<br />
Thailand<br />
+<br />
Cambodia<br />
+<br />
Vietnam<br />
++<br />
P<br />
+ Brunei<br />
Malaysia<br />
Singapore<br />
++<br />
Indonesia<br />
Taiwan<br />
++<br />
Philippines<br />
P<br />
Australia<br />
Papua<br />
New Guinea<br />
+<br />
The moth Agrius convolvuli is widespread in the tropics and subtropics, except for the<br />
Americas where it does not occur.<br />
It is an important pest, sporadically, <strong>of</strong> sweet potato and also attacks several important<br />
legumes. For most <strong>of</strong> the time its populations are maintained at subeconomic levels,<br />
apparently by several trichogrammatid egg parasitoids. These could be considered for<br />
introduction as biological control agents where they do not already occur. The cause <strong>of</strong><br />
sporadic outbreaks is unknown.<br />
20°<br />
0°<br />
20°<br />
9
10 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Agrius convolvuli (Linnaeus)<br />
Rating<br />
Origin<br />
Distribution<br />
Biology<br />
Lepidoptera: Sphingidae<br />
sweet potato hawk moth, sweet potato hornworm<br />
Synonym: Herse convolvuli<br />
<strong>Southeast</strong> Asia China Southern and Western Pacific<br />
++ Viet, Indo ++ + N Cal, PNG<br />
7 + Thai, Camb<br />
Msia<br />
2<br />
P Myan, Brun P Widespread<br />
Very widespread in tropical and subtropical areas <strong>of</strong> the world, except for<br />
the Americas.<br />
Southern Europe: Azores, Crete, Malta, Sicily, Yugoslavia. Africa:<br />
Algeria, Angola, Benin, Burundi, Cape Verde Is, Congo, Egypt, Ethiopia,<br />
Ghana, Ivory Coast, Kenya, Libya, Madagascar, Madeira, Mali, Mauritius,<br />
Morocco, Mozambique, Niger, Nigeria, Rwanda, St Helena, Senegal,<br />
Seychelles, Sierra Leone, Somalia, South Africa, Sudan, Swaziland,<br />
Tanzania, Togo, Tunisia, Uganda, Upper Volta, Zambia, Zimbabwe. Asia:<br />
Andaman Is, Bangladesh, Bhutan, Cambodia, China, Christmas Is, Cyprus,<br />
India, Indonesia, Iran, Iraq, Israel, Japan, Laos, Malaysia, Myanmar,<br />
Pakistan, Philippines, Saudi Arabia, Singapore, Sri Lanka, Syria, Thailand,<br />
Turkey, Vietnam. Australasia and Pacific Islands:<br />
Australia, Cook Is, Fiji,<br />
Hawaii, Kiribati, Mariana Is, Marquesas Is, New Caledonia, New Zealand,<br />
Niue, Norfolk Is, Papua New Guinea, Samoa, Solomon Is, Tonga, Tuvalu,<br />
Vanuatu (CIE Map No 451, 1983).<br />
The smooth eggs are laid singly on stems and leaves and, in common with<br />
most other Lepidoptera, A. convolvuli larvae have 5 instars. There is a green,<br />
a black and a brown form <strong>of</strong> larvae, which have, at the posterior end, a<br />
uniformly curved, tapering, smooth dorsal horn. Fully grown larvae attain a<br />
length <strong>of</strong> 9 cm. Pupation occurs in earthern cells several centimetres below<br />
the soil surface. The pupa has a very characteristic proboscis, which is<br />
enclosed in a looped tube not fused to the body (Kalshoven 1981; Common
Host plants<br />
Damage<br />
4.1<br />
Agrius convolvuli<br />
1990). The mean development period at 25¡C in Japan was 21.2 days<br />
(Setokuchi et al. 1985). In Egypt at 30¡C and 61% RH average<br />
developmental periods were: larvae 14.4 days, prepupae 1.9 days and pupae<br />
13 days (Awadallah et al. 1976). The moths <strong>of</strong>ten enter houses in the evening<br />
and, when at rest, resemble pieces <strong>of</strong> bark. There are at least two generations<br />
during summer, and winter is passed as a pupa.<br />
An artificial diet containing powdered sweet potato leaf has been<br />
developed (Kiguchi and Shimoda 1994). On this at 27¡C and with a day<br />
length <strong>of</strong> 16 hours, A. convolvuli larvae moulted to the 5th instar 12 to 14<br />
days after hatching, pupated at 21 to 26 days and adults emerged at 36 to 41<br />
days. The 5th instar larvae grew to 8 cm in length and 11 to 12 g in weight<br />
(Shimoda et al. 1994). Consumption <strong>of</strong> sweet potato leaves was greatest at<br />
30¡C, the last instar eating 88% <strong>of</strong> the total dry weight (5 g) consumed<br />
(Setokuchi et al. 1986).<br />
The main commercial host is sweet potato ( Ipomoea batatas),<br />
but larvae also<br />
attack other Ipomoea species [e.g. I. pescapreae,<br />
I. cairica,<br />
I. indica<br />
(morning glory) I. hederifolia,<br />
(Moulds 1981)] and other Convolvulaceae<br />
[e.g. Merremia dissecta;<br />
bindweed, Convolvulus arvensis;<br />
Awadallah et al.<br />
1976; (Moulds 1981)]. Several pulses are attacked [e.g. wild mung, Vigna<br />
vexillata (Govindan et al. 1989); moth bean, V. aconitifolia (Bhat et al.<br />
1990); mung bean, V. radiata and urd bean, V. mungo (Shaw et al. 1989);<br />
and also Phaseolus spp. (Nagarkatti 1973)]. A strain <strong>of</strong> moth bean (IPCMO<br />
131) showed good resistance to attack (Bhat et al. 1990). In Papua New<br />
Guinea taro is also recorded as a host (Smee 1965).<br />
A. convolvuli larvae can defoliate sweet potato vines and, even when damage<br />
is less severe, harvest is delayed, increasing the likelihood <strong>of</strong> major attack by<br />
the sweet potato weevil, Cylas formicarius.<br />
Defoliation <strong>of</strong> pulses results in<br />
partial or complete crop failure.<br />
Natural enemies<br />
These are shown in Table 4.1.1.<br />
11
Table 4.1.1<br />
Natural enemies <strong>of</strong> Agrius convolvuli<br />
Species<br />
DIPTERA<br />
PHORIDAE<br />
Country Reference<br />
Megaselia rufipes<br />
TACHINIDAE<br />
Ireland Flemying 1918<br />
Sturmia dilabida<br />
Zimbabwe Cuthbertson 1934<br />
Zygobothria (= Argyrophylax = Sturmia)<br />
atropivora<br />
Malaysia<br />
Zimbabwe<br />
Zygobothria ciliata (= Sturmia macrophallus)<br />
Indonesia<br />
Oman<br />
Philippines<br />
HYMENOPTERA<br />
Corbett & Miller 1933<br />
Cuthbertson 1934<br />
Baran<strong>of</strong>f 1934<br />
Whitcombe & Erzinclioglu 1989<br />
Kalshoven 1981<br />
BRACONIDAE<br />
Apanteles spp.<br />
EULOPHIDAE<br />
China Wu 1983<br />
species<br />
ICHNEUMONIDAE<br />
China Wu 1983<br />
Amblyteles fuscipennis<br />
Central Europe Fahringer 1922<br />
England Morley & Rait-Smith 1933<br />
Charops bicolor<br />
China Wu 1983<br />
Hadrojoppa cognatoria<br />
Japan Uchida 1924, 1930<br />
Trogus exaltatorius<br />
SCELIONIDAE<br />
England Morley & Rait-Smith 1933<br />
Telenomus sp. India Nagarkatti 1973<br />
12 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong>
Table 4.1.1<br />
(contÕd) Natural enemies <strong>of</strong> Agrius convolvuli<br />
Species<br />
HYMENOPTERA<br />
Country Reference<br />
TRICHOGRAMMATIDAE<br />
Trichogramma achaeae<br />
India Nagarkatti 1973<br />
Trichogramma agriae<br />
India Nagarkatti 1973<br />
Trichogramma australicum<br />
India Nagarkatti 1973<br />
Trichogramma chilonis<br />
Guam Nafus & Schreiner 1986<br />
Trichogramma confusum<br />
India Nagarkatti & Nagaraja 1978<br />
Trichogramma ?minutum<br />
Indonesia Leefmans 1929; Kalshoven 1981<br />
Trichogramma sp.<br />
FUNGI<br />
Philippines Shibuya & Yamashita 1936<br />
Entomophthora sp. Ôgrylli'Õ<br />
type Japan Kushida et al. 1975<br />
4.3<br />
Agrius convolvuli<br />
13
14 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Attempts at classical biological control<br />
CHINA<br />
GUAM<br />
INDIA<br />
A species <strong>of</strong> Trichogramma,<br />
possibly T. australicum, (Nagarkatti 1973), has<br />
been imported on two occasions (Table 4.1.2) to attack the eggs <strong>of</strong> pest<br />
Lepidoptera, including Agrius convolvuli,<br />
but the resulting impact on<br />
populations <strong>of</strong> the sweet potato hawk moth is not recorded.<br />
Table 4.1.2 Attempts at classical biological control <strong>of</strong> A. convolvuli<br />
Species From To Year Result Reference<br />
HYMENOPTERA<br />
TRICHOGRAMMATIDAE<br />
Trichogramma<br />
?australicum<br />
Trichogramma<br />
?australicum<br />
USA Indonesia before<br />
1929<br />
+ Leefmans 1929;<br />
Nagarkatti 1973<br />
Philippines Japan 1929 + Shibuya &<br />
Yamashita 1936;<br />
Nagarkatti 1973<br />
In Fujian Province, A. convolvuli larvae were parasitised by Charops bicolor<br />
(Ichneumonidae), Apanteles spp. (Braconidae) and eulophid wasps (Wu<br />
1983).<br />
A. convolvuli is a minor pest <strong>of</strong> sweet potato on Guam. When sweet potato<br />
was intercropped with maize, A. convolvuli eggs were parasitised to the<br />
extent <strong>of</strong> 70 to 100% by Trichogramma chilonis.<br />
This parasitoid attacks the<br />
eggs <strong>of</strong> a range <strong>of</strong> sphingids and noctuids, including Ostrinia furnacalis,<br />
less<br />
than 20% <strong>of</strong> whose eggs on maize were parasitised. A. convolvuli colonises<br />
new sweet potato plantings as soon as cuttings strike and, by the 4th week,<br />
30 to 60% <strong>of</strong> its eggs are parasitised. Each large egg produces 13±<br />
7<br />
parasitoids, which emerge about 10 days after the host egg is parasitised. It<br />
was concluded that T. chilonis is a major mortality factor for the sweet potato<br />
hornworm (Nafus and Schreiner 1986).<br />
A. convolvuli is an occasional pest <strong>of</strong> sweet potato, Vigna mungo and Vigna<br />
radiata.<br />
Eggs are also laid on the leaves <strong>of</strong> Colocasia antiquorum and<br />
Clerodendrum chinense,<br />
but no significant feeding occurs on these latter<br />
plants.<br />
Four species <strong>of</strong> parasitoid attack the eggs <strong>of</strong> A. convolvuli near<br />
Bangalore: Trichogramma australicum, T. achaeae, T. agriae and a species<br />
<strong>of</strong> Telenomus (Eulophidae). The abundance <strong>of</strong> each parasitoid varied with<br />
the plant species on which the eggs were laid. T. agriae was the commonest<br />
species in eggs collected on Colocasia, followed by T. achaeae and
4.1 Agrius convolvuli 15<br />
Telenomus sp., up to a total <strong>of</strong> 43.6%. T. australicum, followed by<br />
Telenomus sp., were the main species emerging from eggs on<br />
Clerodendrum, up to a total <strong>of</strong> 63.9%. At no time were T. achaeae or<br />
T. agriae reared from eggs on Clerodendrum. Furthermore, T. australicum<br />
was reared only twice from Agrius eggs on Colocasia. These results<br />
highlight the difficulty <strong>of</strong> reaching decisions on host specificity on the basis<br />
<strong>of</strong> laboratory trials in a non-natural environment.<br />
Up to 49 Trichogramma individuals were reared from a single<br />
A. convolvuli egg and only in two instances were more than 1 species reared<br />
from a single egg. These were 7 T. agriae and 4 T. australicum on one<br />
occasion and 7 T. achaeae and 11 T. australicum in the second. Eggs<br />
parasitised by Telenomus sp. usually produced 3 to 5 adults and at no time<br />
did a Trichogramma emerge from the same egg as a Telenomus (Nagarkatti<br />
1973). Later, an additional parasitoid (Trichogramma confusum) was<br />
recorded from the eggs <strong>of</strong> A. convolvuli on Clerodendrum chinense<br />
(Nagarkatti and Nagaraja 1978).<br />
Nagarkatti (1973) suggested that the 4 former species might be<br />
introduced where A. convolvuli is a pest and where they do not already<br />
occur.<br />
INDONESIA<br />
Leefmans (1929) reported the parasitisation <strong>of</strong> A. convolvuli eggs by<br />
Trichogramma minutum imported from America. However, Nagarkatti<br />
(1973) suggests that, from the distribution <strong>of</strong> T. minutum at that time, it must<br />
have been T. australicum or some other species <strong>of</strong> Trichogramma.<br />
IRELAND<br />
An adult A. convolvuli produced, soon after capture, many small puparia,<br />
from which 76 Megaselia rufipes (Diptera, Phoridae) emerged (Flemying<br />
1918).<br />
JAPAN<br />
A species <strong>of</strong> Trichogramma that parasitises the eggs <strong>of</strong> Chilo<br />
suppressalis (= C. simplex) was imported in 1929 from the Philippines. It<br />
was shown to parasitise also the eggs <strong>of</strong> A. convolvuli and 10 other species <strong>of</strong><br />
Lepidoptera belonging to several families (Shibuya and Yamashita 1936).<br />
Nagarkatti (1973) suggests that the species was Trichogramma australicum.<br />
OMAN<br />
Adults <strong>of</strong> the tachinid Zygobothria ciliata emerged from puparia from a<br />
larva collected on Ipomoea (Whitcombe and Erzinclioglu 1989).
16 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
SOUTH AFRICA<br />
A. convolvuli is a common pest in the eastern part <strong>of</strong> South Africa. Although<br />
it is generally not abundant, from time to time large areas <strong>of</strong> sweet potatoes<br />
have been almost completely defoliated by it. There are 3 generations a year<br />
and overwintering occurs as the pupa. Natural enemies include the whitebellied<br />
stork (Ciconia nigra) which, on occasion, destroys large numbers <strong>of</strong><br />
larvae (Anon. 1927).<br />
ZIMBABWE<br />
A. convolvuli larvae on sweet potato were parasitised by the tachinids<br />
Zygobothria atropivora and Sturmia dilabida, both <strong>of</strong> which are widely<br />
distributed in South Africa. The latter parasitoid also attacks larvae <strong>of</strong><br />
Spodoptera exigua (Cuthbertson 1934).<br />
Discussion<br />
The majority <strong>of</strong> the parasitoids recorded as attacking A. convolvuli also<br />
attack the eggs or larvae <strong>of</strong> a range <strong>of</strong> other Lepidoptera living in the same<br />
environment. Many <strong>of</strong> these are pest species. Lack <strong>of</strong> parasitoid specificity is<br />
a significant advantage when dealing with a strong flying species, such as<br />
A. convolvuli, which can travel long distances, since the parasitoids are more<br />
likely to be already present on some other host when adult moths arrive to<br />
oviposit at a new site. On the other hand, lack <strong>of</strong> specificity is a disadvantage<br />
if the non-target species that are attacked include environmentally important<br />
species, the lowering <strong>of</strong> whose population density is considered undesirable.<br />
In the present instance it is clear, from the information outlined earlier<br />
under India that, whereas the Trichogramma egg parasitoids involved attack<br />
the eggs <strong>of</strong> a range <strong>of</strong> species <strong>of</strong> Lepidoptera, they do so only when the eggs<br />
are laid on particular host plants. In this sense they, indeed, display a<br />
valuable degree <strong>of</strong> specificity, which should be taken into consideration<br />
when deciding whether or not to proceed with introductions.<br />
With these qualifications it is clear that the establishment, in areas where<br />
they do not already occur, <strong>of</strong> any or all <strong>of</strong> 4 Trichogramma species<br />
(T. achaeae, T. agriae, T. australicum, T. chilonis) is highly likely to lead to<br />
a reduction to (or at least towards) subeconomic levels in the population <strong>of</strong><br />
A. convolvuli.<br />
The underlying causes <strong>of</strong> the sporadic outbreaks <strong>of</strong> A. convolvuli are<br />
unknown. Comparatively little work also has been carried out on the<br />
parasitoids and more detailed studies may well reveal attractive new options<br />
to pursue.
4.2 Anomis flava<br />
India<br />
20°<br />
Myanmar<br />
P Laos<br />
P<br />
0°<br />
20°<br />
China<br />
++<br />
Thailand<br />
+<br />
Cambodia<br />
+<br />
Vietnam<br />
+++<br />
++<br />
Malaysia<br />
Singapore<br />
Brunei<br />
+<br />
Indonesia<br />
Taiwan<br />
Philippines<br />
Australia<br />
Papua<br />
New Guinea<br />
The noctuid moth Anomis flava occurs widely in Africa, Asia and Oceania, where its larvae<br />
sporadically, but seriously, damage cotton, okra, kenaf and other Malvaceae: its adults are<br />
fruit-sucking moths. Its sporadic occurrence suggests that it may be under effective<br />
biological control for much <strong>of</strong> the time.<br />
It is attacked by non-specific predators and by a number <strong>of</strong> parasitoids. Many <strong>of</strong> the<br />
latter attack other Lepidoptera in the same plant environment and appear to be specific to<br />
larvae in that environment rather than to individual species inhabiting it.<br />
Further studies are needed to provide information on what the prospects are for<br />
classical biological control.<br />
17<br />
20°<br />
0°<br />
20°
18 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Anomis flava (Fabricius)<br />
Rating<br />
Origin<br />
Distribution<br />
Biology<br />
Lepidoptera: Noctuidae: Ophiderinae<br />
cotton semi looper, green semi looper, okra semi looper<br />
Synonyms: Cosmophila flava,<br />
Cosmophila indica.<br />
Cosmophila is<br />
now regarded as a subgenus <strong>of</strong> Anomis.<br />
A. flava does not occur in<br />
the Americas, where its equivalent is Anomis ( Cosmophila)<br />
erosa<br />
(Pearson 1958). Records <strong>of</strong> A. erosa in the <strong>Asian</strong> continent should<br />
be referred to A. flava.<br />
<strong>Southeast</strong> Asia Southern China Pacific<br />
+++ Viet<br />
10 ++ Msia ++<br />
+ Thai, Camb,<br />
Indo<br />
P Myan, Laos, Phil present, but not important<br />
Unclear: could be Africa or Asia. Information available on specific or<br />
reasonably specific parasitoids possibly favours Africa.<br />
Africa:<br />
Central and southern countries, including Angola, Benin,<br />
Cameroun, Chad, Congo, Ethiopia, Gambia, Ghana, Ivory Coast, Kenya,<br />
Madagascar, Malawi, Mali, Mauritius, Niger, Nigeria, Senegal, Somalia,<br />
Sudan, Tanzania, Togo, Uganda, Upper Volta, Zambia, Zimbabwe. Asia:<br />
Cambodia, China, India, Indonesia, Japan, Korea, Laos, Malaysia,<br />
Myanmar, Pakistan, Philippines, Sri Lanka, Taiwan, Thailand, Vietnam.<br />
Australasia and Pacific Islands:<br />
Australia, Cook Is, Fiji, Mariana Is,<br />
Marquesas, New Caledonia, Papua New Guinea, Samoa, Solomon Is.,<br />
Tonga, Vanuatu (CIE 1978), Rapa Is, Hawaii (Common 1990).<br />
Most A. flava eggs are laid on the undersurface <strong>of</strong> leaves, the young larvae<br />
are green, those <strong>of</strong> the last instar measure up to 35 mm in length and bear<br />
short, lighter-green longitudinal lines and spots. Young larvae skeletonise<br />
leaves, older larvae consume narrow leaf (roselle) cotton leaves and eat<br />
irregular holes in broader leaves. Larval survival and growth are greater on
Host plants<br />
Damage<br />
4.2<br />
Anomis flava<br />
hirsutum than on desi cotton (Sidhu and Dhawan 1979; Kalshoven 1981).<br />
Pupation occurs in a cocoon spun between leaves. Development times have<br />
been recorded on a number <strong>of</strong> occasions (for examples see Table 4.2.1, also<br />
Schmitz 1968; Yu and Tu 1969), egg to adult taking about 3 weeks or a little<br />
longer and the number <strong>of</strong> eggs laid ranging from 158 to 476, depending, in<br />
part, upon the larval food plant. Groups <strong>of</strong> larvae normally pass through 5<br />
moults whereas, when reared singly, up to 22% pass through 6 moults<br />
(Kirkpatrick 1963; Essien and Odebiyi 1991). There are 5 overlapping<br />
generations a year in Hunan Province, China, but fewer in some other<br />
regions (Chen et al. 1991).<br />
Adults rest in foliage by day and are active in the evening: they are<br />
attracted to light.<br />
A. flava is a major, but sporadic, pest <strong>of</strong> cotton. Larvae also attack many<br />
other plants, mainly in the family Malvaceae. These include, especially, okra<br />
( Hibiscus esculentus),<br />
but also kenaf or Deccan hemp ( H. cannabinus),<br />
jute<br />
( H. sabadariffa),<br />
bele ( H. manihot),<br />
muskmallow an important medicinal<br />
plant ( H. abelmoschus),<br />
sho<strong>of</strong>lower ( H. rosa-sinensis),<br />
hollyhock ( Althaea<br />
rosea),<br />
Arbutilon spp., Sida spp. and Urena spp. (all Malvaceae). However<br />
they also attack tomato ( Lycopersicon esculentum:<br />
Solanaceae); cowpea<br />
( Vigna unguiculata)<br />
and green gram ( Vigna radiata):<br />
Fabaceae; sweet<br />
potato ( Ipomoea batatas):<br />
Convolvulaceae; as well as melon ( Citrullus<br />
lanatus),<br />
Macadamia,<br />
Ricinus,<br />
Leea and Amaranthus spp. (Kalshoven 1981;<br />
Yein and Singh 1981; Croix and Thindwa 1986; Gatoria and Singh 1988;<br />
Essien and Odebiyi 1991).<br />
Okra and hemp (kenaf) were the most favoured larval food plants,<br />
whereas cotton and okra were the most favourable in terms <strong>of</strong> pupal weight<br />
and adult fecundity (Rao and Patel 1973).<br />
When abundant, A. flava larvae are capable <strong>of</strong> causing serious damage by<br />
destroying the leaves and buds <strong>of</strong> cotton and other Malvaceous crops.<br />
A. flava belongs to the subfamily Ophiderinae <strong>of</strong> noctuids, the adults <strong>of</strong><br />
which are <strong>of</strong>ten fruit-piercing species. In southern China, A. flava is reported<br />
to be a serious pest <strong>of</strong> citrus fruit (Li et al. 1997) and, in Korea, A. flava is one<br />
<strong>of</strong> a group <strong>of</strong> fruit-sucking moths observed to damage grapes and pears (Lee<br />
et al. 1970).<br />
19
Table 4.2.1<br />
Average figures (days) for development <strong>of</strong> Anomis flava<br />
Stage Rao & Patel 1973 Kalshoven 1981<br />
Author<br />
Ferino et al. 1982a Chen et al. 1991 Essien & Odebiyi 1991<br />
egg development 2 2Ð3 4Ð5 2Ð7<br />
1st instar larva 3 2.4<br />
2nd 1.8 1.8<br />
3rd 1.9 11 12Ð16 2.2<br />
4th 2 2.2<br />
5th 3 2.3<br />
prepupa 1 1.4<br />
pupa 6.2 6Ð11 7<br />
eggÐadult 21.1 21 19Ð23 22Ð29 28<br />
female longevity 31 28 10 4Ð7 19.8<br />
number <strong>of</strong> eggs 158 350 492 476<br />
preÐoviposition 1.25 3 3.3<br />
oviposition 6 7 12.4<br />
20 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong>
4.2<br />
Anomis flava<br />
A. flava is regarded as <strong>of</strong> only minor importance in the Pacific, which is<br />
not surprising since none <strong>of</strong> its major larval host plants is <strong>of</strong> much economic<br />
importance there.<br />
Natural enemies<br />
Those reported in the literature are listed in Table 4.2.2.<br />
Egg parasitoids are Trichogramma spp., which on occasion can be<br />
effective: in Mali, 92% parasitisation by Trichogramma sp. was recorded in<br />
untreated cotton (Pierrard 1970), 12.1% to 15% in the Philippines (Ferino et<br />
al. 1982a) and 60 to 80% <strong>of</strong> eggs on cotton were attacked by T. dendrolimi in<br />
China (Wang et al. 1985, 1988).<br />
As for larval parasitoids, Apanteles anomidis parasitised 27.5% in China<br />
(Xie 1984), Aleiodes aligharensi and Aleiodes sp. together 5.2% in Chad<br />
(Silvie et al. 1989), Charops bicolor 10.2% in China (Xie 1984), Meteorus<br />
pulchricornis 4.9% in China (Xie 1984), Meteorus sp. 50 to 69.4% in Nepal<br />
(Neupane 1977) and Winthemia dasyops 2.5% in Chad (Silvie et al. 1989).<br />
Most other records did not indicate effectiveness or, if they did, it was lower<br />
than 2.5% parasitisation.<br />
A. flava pupae are attacked by at least 5 species <strong>of</strong> Brachymeria<br />
(Chalcididae). In Madagascar, B. multicolor and B. tibialis parasitised 98%<br />
<strong>of</strong> pupae in some fields (Steffan 1958).<br />
Further details are provided in the country summaries. It is not easy to<br />
discern a pattern from these although, under some conditions, parasitoids are<br />
clearly able to have a major impact on A. flava populations.<br />
Less is known about the effectiveness <strong>of</strong> predators, although pentatomid,<br />
carabid, coccinellid, vespid and spider predators have been reported and the<br />
Indian mynah bird consumed large numbers <strong>of</strong> larvae when they were<br />
abundant (Khan 1956).<br />
Unexplained disappearance <strong>of</strong> larvae is <strong>of</strong>ten attributed to predation,<br />
although heavy rainfall may sometimes be responsible.<br />
Bacillus thuringiensis has been recorded in the field from A. flava larvae<br />
(Yin et al. 1991) and has given promising control on a number <strong>of</strong> occasions<br />
(Angelini and Couilloud 1972; Delattre 1973; Anon 1976b; Wilson 1981;<br />
Chen et al. 1991).<br />
Both granulosis and polyhedrosis viruses have been recorded in the field<br />
(Table 4.2.2) and it is possible that virus preparations might be used for<br />
control.<br />
21
Table 4.2.2<br />
Natural enemies <strong>of</strong> Anomis flava<br />
Species<br />
DERMAPTERA<br />
CARCINOPHORIDAE<br />
Country Reference<br />
Euborellia pallipes<br />
HEMIPTERA<br />
ANTHOCORIDAE<br />
China Yang 1985a<br />
Orius minutus<br />
LYGAEIDAE<br />
China Wu et al. 1981<br />
Geocoris sp.<br />
NABIDAE<br />
China Wu et al. 1981<br />
Nabis sin<strong>of</strong>erus<br />
PENTATOMIDAE<br />
China Wu et al. 1981<br />
Cermatulus nasalis<br />
Australia Kay & Brown 1991<br />
Eucanthecona (= Cantheconidia) furcellata<br />
China Wu et al. 1981<br />
Oechalia schellembergii<br />
NEUROPTERA<br />
CHRYSOPIDAE<br />
Australia Wilson 1981<br />
Chrysopa sp.<br />
DIPTERA<br />
TACHINIDAE<br />
China Wu et al. 1981<br />
?Isyropa<br />
India Maheswariah & Puttarudriah 1956<br />
Cadurcia (= Sturmia) auratocaudata Nigeria, Gold Coast Curran 1934<br />
Camplyocheta (= Elpe) sp. Cameroun Deguine 1991<br />
Carcelia (= Zenilla) cosmophilae Australia Curran 1934, 1938<br />
Carcelia kockiana India Sohi 1964<br />
Carcelia illota (= Zenilla noctuae) Australia Curran 1934, 1938; Kay & Brown 1991<br />
Cylindromya (= Ocyptera) sp. Senegal Risbec 1950<br />
22 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong>
Table 4.2.2 (contÕd) Natural enemies <strong>of</strong> Anomis flava<br />
Species<br />
DIPTERA<br />
Country Reference<br />
TACHINIDAE (contÕd)<br />
Exorista apicalia India Sohi 1964<br />
Exorista sorbillans Australia Kay & Brown 1991<br />
Palexorista inconspicua (= Sturmia bimaculata) Africa Pearson 1958<br />
Palexorista quadrizonula Africa<br />
Tanzania<br />
Crosskey 1970<br />
Robertson 1973<br />
Sericophoromyia marshalli South Africa Taylor 1930<br />
unidentified Chad Silvie et al. 1989<br />
Philippines Ferino et al. 1982a<br />
Winthemia dasyops Chad Silvie et al. 1989<br />
Zygobothria ciliata (= Sturmia macrophallus) India Thompson 1944; Sohi 1964<br />
HYMENOPTERA<br />
BRACONIDAE<br />
Aleiodes aligharensi Chad Silvie et al. 1989; Silvie 1991<br />
Aleiodes sp. Chad Silvie et al. 1989; Silvie 1991<br />
Philippines Ferino et al. 1982a<br />
Apanteles anomidis China<br />
Vietnam Xie 1984; Xiong et al. 1994;<br />
van Lam 1996<br />
Apanteles spp. India<br />
Philippines Maheswariah & Puttarudriah 1956; Sohi1964;<br />
Ferino et al. 1982a<br />
Apanteles syleptae Chad Silvie et al. 1989; Silvie 1991<br />
Cotesia (= Apanteles) ruficrus China Woo & Hsiang 1939<br />
Fiji Lever 1943<br />
Philippines Ferino et al. 1982a<br />
4.2 Anomis flava 23
Table 4.2.2 (contÕd) Natural enemies <strong>of</strong> Anomis flava<br />
Species<br />
HYMENOPTERA<br />
Country Reference<br />
BRACONIDAE (contÕd)<br />
Disophrys lutea Tanzania Robertson 1973<br />
Meteorus pulchricornis (= M. japonicus) China Chu 1934; Xie 1984<br />
Meteorus sp. nr fragilis Nepal Neupane 1977<br />
Nyereria sp. Chad Silvie et al. 1989<br />
Parapanteles sp. Chad Silvie et al. 1989<br />
Protomicroplitis sp. Chad Silvie et al. 1989<br />
Sigalphus nigripes<br />
CHALCIDIDAE<br />
China He & Chen 1993<br />
Brachymeria nr aliberti Chad Silvie et al. 1989<br />
Brachymeria lasus (= B. obscurata) China Chu & Hsia 1935; Woo & Hsiang 1939<br />
Philippines Ferino et al. 1982a<br />
Vietnam van Lam 1996<br />
Brachymeria madecassa Mauritius Vaissayre 1977<br />
Brachymeria multicolor Madagascar Steffan 1958<br />
Brachymeria paolii Tanzania Robertson 1973<br />
Brachymeria sp. Australia Kay & Brown 1991<br />
Brachymeria tibialis<br />
EULOPHIDAE<br />
Madagascar Steffan 1958<br />
Euplectrus manilae Philippines Ferino et al. 1982a; Otanes & Butac 1935;<br />
Otanes 1935<br />
Tetrastichus howardi (= T. ayyari) India Maheswariah & Puttarudriah 1956, Sohi 1964<br />
24 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong>
Table 4.2.2 (contÕd) Natural enemies <strong>of</strong> Anomis flava<br />
Species<br />
HYMENOPTERA<br />
Country Reference<br />
EUMENIDAE<br />
Delta (= Eumenes) pyriforme Philippines Ferino et al. 1982a<br />
Eumenes campaniformis<br />
ICHNEUMONIDAE<br />
Philippines Ferino et al. 1982a,b<br />
Charops bicolor China Xie 1984<br />
Charops sp. Senegal Risbec 1950<br />
Echthromorpha agrestoria Australia Kay & Brown 1991<br />
Enicospilus ?samoana Kay & Brown 1991<br />
Enicospilus dolosus Chad Silvie et al. 1989; Silvie 1991<br />
Enicospilus sp. Tanzania Robertson 1973<br />
Mesochorus sp. China Xie 1984<br />
Metopius sp. Vietnam van Lam 1996<br />
Xanthopimpla punctata China Woo & Hsiang 1939<br />
Zacharops narangae<br />
TRICHOGRAMMATIDAE<br />
China Chu 1934; Woo & Hsiang 1939<br />
Trichogramma chilonis Vietnam Nguyen & Nguyen 1982<br />
Trichogramma dendrolimi China Wang et al. 1985, 1988<br />
Trichogramma minutum India Maheswariah & Puttarudriah 1956; Sohi 1964<br />
Philippines Otanes & Butac 1935<br />
Trichogramma japonicum Vietnam Nguyen & Nguyen 1982<br />
Trichogramma sp. Australia Twine & Lloyd 1982<br />
sp. Mali Pierrard 1970<br />
spp. Philippines Ferino et al. 1982b<br />
4.2 Anomis flava 25
Table 4.2.2 (contÕd) Natural enemies <strong>of</strong> Anomis flava<br />
Species<br />
HYMENOPTERA<br />
Country Reference<br />
VESPIDAE<br />
Polistes jokahamae China Anon. 1976a<br />
Polistes sp. China Anon. 1976a<br />
COLEOPTERA<br />
CARABIDAE<br />
Calosoma schayeri Australia Twine & Lloyd 1982<br />
Lissauchenius venator Cameroun Deguine 1991<br />
COCCINELLIDAE<br />
Coccinella septempunctata China Wu et al. 1981<br />
ARACHNIDA<br />
Erigonidium graminicolum China Wu et al. 1981<br />
Misumenops tricuspidatus China Wu et al. 1981<br />
sp. (Oxyopidae) Philippines Ferino et al. 1982a<br />
sp. (Thomisidae)<br />
NEMATODA<br />
Philippines Ferino et al. 1982a<br />
MERMITHIDAE India Mundiwale et al. 1978<br />
not specified<br />
BACTERIA<br />
Chad Silvie 1991<br />
Bacillus thuringiensis wuhanensis<br />
VIRUSES<br />
China Yin et al. 1991<br />
Granulosis China Yin et al. 1991<br />
Polyhedrosis Australia Bishop et al. 1978<br />
Cameroun Delattre 1973<br />
China Liang et al. 1981<br />
Mali Atger & Chevalet 1975<br />
AVES<br />
Vietnam van Cam et al. 1996<br />
Acridotheres tristis India Khan 1956<br />
26 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong>
4.2 Anomis flava 27<br />
Attempts at classical biological control<br />
There appear to have been only two attempts (Table 4.2.3). The pentatomid<br />
bug Podisus maculiventris, a general predator <strong>of</strong> lepidopterous larvae, was<br />
introduced from USA (where A. flava does not occur) and liberated in Anhui<br />
Province, China in 1984. However, it failed to become established, possibly<br />
due to adverse climatic conditions (Wang and Gong 1987). Trichogramma<br />
minutum from USA was established, in the Philippines in 1934, but its<br />
impact is not recorded (Otanes and Butac 1935).<br />
Table 4.2.3 Attempts at biological control <strong>of</strong> Anomis flava<br />
Species<br />
HEMIPTERA<br />
PENTATOMIDAE<br />
From To Year Result Reference<br />
Podisus maculiventris<br />
HYMENOPTERA<br />
TRICHOGRAMMATIDAE<br />
USA China 1984 Ð Wang &<br />
Gong 1987<br />
Trichogramma minutum USA Philippines 1934 + Otanes &<br />
Butac 1935<br />
AUSTRALIA<br />
Regular releases <strong>of</strong> Trichogramma nr praetiosum at the rate <strong>of</strong> 50000 adults/<br />
ha were made from November to March on 8 ha <strong>of</strong> cotton in south eastern<br />
Queensland. The resulting mean rate <strong>of</strong> egg parasitisation (49.4%) was<br />
inadequate to control damage by Helicoverpa spp. and the few eggs <strong>of</strong><br />
A. flava collected were not parasitised, although high levels <strong>of</strong> parasitisation<br />
had been reported following the release <strong>of</strong> the same Trichogramma species<br />
in northern Western Australia (Twine and Lloyd 1982). Good control on<br />
cotton in northern New South Wales was obtained with a mixture <strong>of</strong> Bacillus<br />
thuringiensis and chlordimeform at a time at which, except for coccinellids,<br />
natural enemies were scarce, although low numbers <strong>of</strong> spiders and <strong>of</strong> the<br />
pentatomid predator Oechallia schellembergii were present (Wilson 1981).<br />
A. flava is one <strong>of</strong> two major pests <strong>of</strong> kenaf in northern Queensland and<br />
the Ord Irrigation Area <strong>of</strong> Western Australia, although natural enemies can<br />
produce valuable control (Kay and Brown 1991). The tachinids Carcelia<br />
cosmophilae, C. illota and Exorista sorbillans attack larvae <strong>of</strong> A. flava and<br />
other noctuids. Larvae are also attacked by the predator Cermatulus nasalis<br />
(Pentatomidae) and the parasitoids Brachymeria sp. (Chalcididae),<br />
Echthromorpha agrestoria and Enicospilus ?samoana (both Ichneumonidae)<br />
(Curran 1938; Kay and Brown 1991).
28 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
CHAD<br />
Eleven species <strong>of</strong> parasitoid, 3 species <strong>of</strong> hyperparasitoid and nematodes<br />
were reared from A. flava larvae on cotton (Silvie et al. 1989; Silvie 1991).<br />
Total parasitisation never exceeded 25% and, in 1987, 15.7% <strong>of</strong> 485 A. flava<br />
larvae were parasitised. Details are shown in Table 4.2.4. The commonest<br />
parasitoid was Aleiodes aligharensi which, together with Aleiodes sp.<br />
accounted for nearly a third <strong>of</strong> all larvae parasitised. Three hyperparasitoids<br />
were recorded, about half emerging from species <strong>of</strong> Aleiodes. The most<br />
abundant was Mesochorus (= Stictopisthus) africanus (Ichneumonidae)<br />
followed by Nesolynx phaeosoma (Eulophidae) and Eurytoma syleptae<br />
(Eurytomidae). All three species were also reared from parasitised larvae <strong>of</strong><br />
other host species (Silvie et al. 1989; Silvie 1991).<br />
Table 4.2.4 Natural enemies <strong>of</strong> A. flava larvae on cotton in Chad<br />
Species % <strong>of</strong> total larvae<br />
parasitised<br />
Primary parasitoids<br />
DIPTERA<br />
Other hosts<br />
TACHINIDAE<br />
Winthemia dasyops 15.8 Chrysodeixis acuta<br />
HYMENOPTERA<br />
BRACONIDAE<br />
Aleiodes aligharensi 32.9 Earias sp.<br />
Aleiodes sp. Helicoverpa armigera<br />
Apanteles syleptae 1.3 Syllepte derogata<br />
Nyereria sp. 1.3 Syllepte derogata<br />
Parapanteles sp. 5.3<br />
Protomicroplitis sp.<br />
CHALCIDIDAE<br />
3.9<br />
Brachymeria nr aliberti<br />
ICHNEUMONIDAE<br />
1.3<br />
Enicospilus dolosus 9.2<br />
NEMATODA<br />
Hyperparasitoids<br />
ICHNEUMONIDAE<br />
1.3<br />
Mesochorus (= Stictopisthus) africanus 4.0<br />
Dead parasitoids 23.7
4.2 Anomis flava 29<br />
CHINA<br />
Since 1970 the cultivation <strong>of</strong> bluish dogbane (Apocynum venotum) has<br />
increased greatly in Zhejiang Province, where A. flava is its most important<br />
pest and 43.7% <strong>of</strong> semilooper larvae were parasitised. There were two<br />
braconids, Apanteles anomidis (27.5% parasitisation) and Meteorus<br />
pulchricornis (4.9%); two ichneumonids, Charops bicolor (10 to 15%) and<br />
Mesochorus sp. (2.4%); and an unidentified species (0.54%). Mesochorus<br />
sp. acted as a hyperparasitoid <strong>of</strong> Apanteles anomidis, but itself parasitised<br />
about 1% <strong>of</strong> A. flava larvae (Xie 1984).<br />
Trichogramma chilonis was reared from the eggs <strong>of</strong> A. flava on cotton in<br />
Shanxi (Huo et al. 1988). Inoculative releases <strong>of</strong> T. dendrolimi in vegetable<br />
gardens adjacent to cotton fields infested with A. flava resulted in 61 to 81%<br />
parasitisation <strong>of</strong> its eggs. By comparison, in pesticide-treated fields nearby,<br />
parasitisation ranged from 2.5 to 30%. Inundative releases directly in cotton<br />
fields led to 30 to 80% parasitisation and no additional control measures<br />
were required (Wang et al. 1985, 1988).<br />
Polistes jokahamae and Polistes sp. were observed in Hunan Province<br />
preying on A. flava, the late instar larvae being preferred (Anon. 1976a).<br />
In Hubei Province, the earwig predator Euborellia pallipes was reported<br />
to reduce A. flava larval populations by 38 to 65% (Yang 1985a).<br />
INDIA<br />
Although A. flava is generally a minor pest, serious outbreaks occur<br />
sporadically. In Hyderabad State more than 1.5 million acres <strong>of</strong> cotton were<br />
affected in one outbreak, with up to 30 larvae per plant consuming<br />
everything except branches and bolls. Large numbers <strong>of</strong> the common mynah<br />
were reported eating the larvae (Khan 1956). In Mysore 70% <strong>of</strong> A. flava<br />
larvae on cotton in the field were parasitised by tachinid flies and Apanteles<br />
spp. In the laboratory, eggs were attacked by Trichogramma minutum and<br />
pupae by Tetrastichus howardi (Maheswariah and Puttarudiah 1956).<br />
MADAGASCAR<br />
The non-specific Brachymeria multicolor was recorded as producing more<br />
than 95% parasitisation <strong>of</strong> A. flava larvae on cotton (Steffan 1958; Delattre<br />
1973). B. madecassa was also credited with 50 to 90% parasitisation <strong>of</strong><br />
larvae in 1956 and 1957 (Vaissayre 1977).<br />
NEPAL<br />
The most important parasitoid <strong>of</strong> A. flava larvae, Meteorus sp. nr fragilis<br />
(Braconidae), was responsible for 50 and 69.4% parasitisation in 1973 and<br />
1974 respectively. There were no pupal parasitoids (Neupane 1977).
30 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
PHILIPPINES<br />
High temperatures inhibited and moderate rainfall favoured high<br />
populations <strong>of</strong> A. flava on seed cotton, yield being significantly reduced only<br />
at densities <strong>of</strong> 6 to 8 larvae or greater per plant or at damage rates involving<br />
at least 60% defoliation. Ten species <strong>of</strong> natural enemies were recorded, the<br />
most important being 2 Trichogramma egg parasitoids, a eulophid larval<br />
parasitoid, a larval and pupal predator (Delta (= Eumenes) pyriforme) and a<br />
pupal parasitoid (Brachymeria lasus). Larval disappearance was attributed<br />
to predators, including Eumenes campaniformis and 2 species <strong>of</strong> spiders.<br />
Egg and pupal parasitisation were generally high during the wet season,<br />
whereas larval and pupal predation were higher in the dry season. The major<br />
mortality occurred during the larval stage, followed by pupal mortality, with<br />
egg mortality being least important. Larval disappearance, suspected to be<br />
due to predation, was more important than parasitisation (Ferino et al.<br />
1982a,b).<br />
TAIWAN<br />
A. flava larvae feed on the leaves and buds <strong>of</strong> kenaf and heavy infestation<br />
reduces top growth. There are 3 generations a year, <strong>of</strong> which the 3rd occurs<br />
in July and is the most injurious. In Taiwan the main hosts are cotton and<br />
kenaf, although other Malvaceae are attacked (Yu and Tu 1969). The<br />
biology <strong>of</strong> Eucanthecona furcellata, a pentatomid predator <strong>of</strong> A. flava<br />
larvae, was studied by Chu and Chu (1975).<br />
TANZANIA<br />
Four species <strong>of</strong> parasitoid were reared from A. flava larvae collected from<br />
cotton and kenaf. In 1963, 7% and, in 1964, 13.8% <strong>of</strong> larvae were<br />
parasitised. The species involved were the tachinid fly Palexorista<br />
quadrizonula, which produced 1 to 5 puparia from each parasitised larva and<br />
had an average pupal period <strong>of</strong> 8 days; the ichneumonid Enicospilus sp.<br />
producing 1 pupa, with an average pupal period <strong>of</strong> 13 days; and, <strong>of</strong> lesser<br />
importance, the braconid Disophrys lutea (1 pupa, 5 days) and the chalcid<br />
Brachymeria paolii (1 pupa, 11 days). Palexorista quadrizonula was also<br />
reared from Spodoptera exigua, S. littoralis and Xanthodes graellsii (all<br />
Noctuidae); Enicospilus sp. from Helicoverpa armigera; and Disophrys<br />
lutea from Earias biplaga, Spodoptera exigua and S. littoralis (Robertson<br />
1973).<br />
VIETNAM<br />
<strong>Control</strong> <strong>of</strong> A. flava is particularly good in some years due to two naturally<br />
occurring egg parasitoids, Trichogramma chilonis and T. japonicum, 93%<br />
parasitisation <strong>of</strong> eggs being reported (Nguyen and Nguyen 1982; Nguyen<br />
1986).
Major natural enemies<br />
4.2 Anomis flava 31<br />
Apanteles anomidis Hym.: Braconidae<br />
A. anomidis is an important endoparasite <strong>of</strong> A. flava in China. It has one<br />
generation a year. A mated female lays an average <strong>of</strong> 109 eggs and prefers to<br />
lay in 1st to 3rd instar host larvae. Adults fed on 10% aqueous sugar solution<br />
lived about 1.5 days at 29¡C (Xiong et al. 1994). An average <strong>of</strong> 13.7 pupae <strong>of</strong><br />
A. anomidis were obtained from each parasitised A. flava larva (Xie 1984).<br />
Palexorista quadrizonula Dip.: Tachinidae<br />
This parasitoid was the most important <strong>of</strong> 4 species attacking A. flava in<br />
Tanzania. It is widespread in Africa south <strong>of</strong> the Sahara and occurs also in<br />
the Seychelles and St Helena. It attacks a range <strong>of</strong> lepidopterous larvae,<br />
especially species belonging to the Noctuidae, but also to the Arctiidae,<br />
Geometridae, Pyralidae and Tortricidae. In A. flava it produces 1 to 5<br />
puparia from each larva, with an average developmental period <strong>of</strong> 7.9 days<br />
(Crosskey 1970; Robertson 1973).<br />
Discussion<br />
Many natural enemies <strong>of</strong> A. flava have been reported, although there have<br />
been few studies detailed enough to indicate their true effectiveness. Most <strong>of</strong><br />
the parasitoids are unlikely to be specific to A. flava, but to attack also other<br />
lepidopterous larvae feeding on the same host plants. Most <strong>of</strong> these other<br />
hosts are themselves pest species, whose abundance it is desirable to lower.<br />
Specificity in these circumstances is rather to lepidopterous larvae in a<br />
particular habitat and the parasitoids may thus be sufficiently restricted in<br />
their attack on non-target species to be seriously considered as agents for<br />
classical biological control. Indeed, for a sporadic pest such as A. flava, it is<br />
highly desirable that there should be readily available a reservoir <strong>of</strong> natural<br />
enemies present continuously, so as to be in place when populations <strong>of</strong><br />
A. flava start to increase.<br />
The reasons for sporadic outbreaks have not been identified, although<br />
Brader (1966) suggested that it might well be due to the application <strong>of</strong><br />
insecticides resulting in the death <strong>of</strong> natural enemies.
Table 3<br />
Order No.<br />
<strong>of</strong> +s<br />
26<br />
1. 41<br />
2. 35<br />
3. 34<br />
4. 32<br />
5. 31<br />
6. 30<br />
7. = 29<br />
7. = 29<br />
9. 27<br />
10. 27<br />
11. 26<br />
12. 25<br />
13. 25<br />
14. = 24<br />
14. = 24<br />
16. 24<br />
17. 22<br />
18. 22<br />
19. 21<br />
20. 20<br />
21. = 18<br />
21. = 18<br />
23. 17<br />
24. 17<br />
25. 17<br />
26. 16<br />
27. 16<br />
28. 15<br />
29. 15<br />
30. 15<br />
a<br />
Walker 1993.<br />
Aggregated ratings <strong>of</strong> the major invertebrate pests <strong>of</strong> agriculture in the region.<br />
D.F. Waterhouse<br />
Pest and<br />
+ scores<br />
30 and over<br />
No. times<br />
in top 10<br />
Dossier<br />
available?<br />
Any biological<br />
control successes?<br />
Attractiveness<br />
as a target<br />
Bactrocera spp. 13 + + +<br />
Cosmopolites sordidus<br />
Spodoptera litura<br />
Aphis gossypii<br />
Cylas formicarius<br />
Plutella xylostella<br />
25–29<br />
Crocidolomia pavonana<br />
4 + + + +<br />
4 + – –<br />
6 + + + + +<br />
7 + – –<br />
9 + + + + + +<br />
4 + – +<br />
Liriomyza spp. 4 + + + +<br />
Othreis fullonia<br />
Helicoverpa armigera<br />
Pentalonia nigronervosa<br />
8 + + + + + +<br />
4 + + +<br />
4 + – +<br />
Epilachna spp. 4 + + + +<br />
Aulacophora spp. 2 + – –<br />
20–24<br />
Nacoleia octasema<br />
Maruca vittrata<br />
Polyphagotarsonemus<br />
latus<br />
Agonoxena argaula<br />
Brontispa longissima<br />
Tarophagus proserpina<br />
Aleurodicus dispersus<br />
15–19<br />
Phyllocnistis citrella<br />
Unaspis citri<br />
3 + + + +<br />
3 + – +<br />
1 + – –<br />
5 + + + + +<br />
4 + + + + + +<br />
3 + – + + + +<br />
3 + + + + + +<br />
2 + + + + +<br />
2 – + + + +<br />
Papuana spp. 5 + – +<br />
Adoretus versutus<br />
Dysmicoccus brevipes<br />
Euscepes postfasciatus<br />
Halticus tibialis<br />
Oryctes rhinocerus<br />
Thrips palmi<br />
Coccus viridis<br />
4 + – –<br />
– + + + +<br />
3 – – –<br />
2 – – –<br />
3 + + + + + +<br />
3 a (+)<br />
– –<br />
1 – ? + +<br />
(cont’d over)
Table 3<br />
(cont’d)<br />
Order No.<br />
<strong>of</strong> +s<br />
31. 14<br />
32. 14<br />
32. = 14<br />
32. = 14<br />
35. 13<br />
36. = 12<br />
36. = 12<br />
38. 12<br />
39. 11<br />
40. 11<br />
41. 10<br />
42. 10<br />
43. = 10<br />
43. = 10<br />
43. = 10<br />
46. 9<br />
b<br />
De Barro 1995.<br />
Aggregated ratings <strong>of</strong> the major invertebrate pests <strong>of</strong> agriculture in the region.<br />
10–14<br />
Achatina fulica<br />
Phyllocoptrupa oleivora<br />
Hellula spp.<br />
Nezara viridula<br />
Aspidiotus destructor<br />
Graeffea crouanii<br />
Planococcus pacificus<br />
Earias vittella<br />
Pest and<br />
+ scores<br />
Aphis craccivora<br />
Tetranycus lambi<br />
Bemisia tabaci<br />
Ceroplastes rubens<br />
Hippotion celerio<br />
Rhabdoscelus obscurus<br />
Tetranycus marianae<br />
Still invading<br />
Bemisia argentifolii<br />
No. times<br />
in top 10<br />
Dossier<br />
available?<br />
2 + + + + + +<br />
– – – –<br />
– + – –<br />
– + + + + + +<br />
2 + + + + + +<br />
2 + + + + +<br />
2 – – +<br />
1 – – –<br />
1 + + + +<br />
– – – –<br />
3 b (+)<br />
+ +<br />
1 – + + + +<br />
– – – –<br />
– – + + +<br />
– – – –<br />
3 b (+)<br />
Any biological<br />
control successes?<br />
Attractiveness<br />
as a target<br />
+ +<br />
The Major Invertebrate <strong>Pests</strong> and Weeds <strong>of</strong> Agriculture and Plantation Forestry in the Southern and Western Pacific<br />
27
4.4 Aphis gossypii<br />
India<br />
Myanmar<br />
++<br />
20°<br />
Laos<br />
+<br />
0°<br />
20°<br />
China<br />
+++<br />
Thailand<br />
+++<br />
Cambodia<br />
++<br />
Vietnam<br />
++<br />
+<br />
++ Brunei<br />
Malaysia<br />
+<br />
Singapore<br />
++<br />
Indonesia<br />
Taiwan<br />
++<br />
+++<br />
Philippines<br />
Australia<br />
Papua<br />
New Guinea<br />
+<br />
The comments under the map <strong>of</strong> Aphis craccivora apply also to A. gossypii.<br />
45<br />
20°<br />
0°<br />
20°
46 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
4.4 Aphis gossypii Glover<br />
Rating<br />
Origin<br />
Distribution<br />
Biology<br />
Hemiptera: Aphididae<br />
cotton aphid, melon aphid<br />
<strong>Southeast</strong> Asia China Southern and Western Pacific<br />
+++ Thai, Phil +++ +++ Fiji, Guam, Tong, Van<br />
19 ++ Myan, Camb, Viet,<br />
32 ++ Cook Is, FSM, Fr P, Kiri,<br />
Msia, Indo<br />
Niue, Sam, Tuv<br />
+ Laos, Sing, Brun + N Cal, PNG, A Sam, Sol<br />
Is, Tok, W & F<br />
P P Tuv, Van<br />
Unclear. Starù (1967a) suggests Ôprobably steppe areas <strong>of</strong> the Palaearctic<br />
regionÕ, possibly inferring southeastern Europe and adjoining regions. The<br />
taxonomic status <strong>of</strong> A. gossypii is complex and there are a number <strong>of</strong><br />
biotypes.<br />
A. gossypii is now very widespread throughout warm temperate, subtropical<br />
and tropical regions <strong>of</strong> the world.<br />
The cotton aphid varies greatly in colour, usually from light green or dark<br />
green to almost black but, for older, overcrowded larvae (nymphs) and at<br />
high temperatures it is yellow to almost white and the aphids are smaller than<br />
on young growth. Wingless females (apterae) vary from 0.9 to 1.8 mm in<br />
length and winged females (alatae) 1.1 to 1.8 mm.<br />
In Europe, there is no sexual reproduction, but there is in East Africa,<br />
USA, China and Japan. However, in Japan, there are also parthenogenetic<br />
overwintering populations (Komazaki 1993). The young generally moult 4<br />
times (range 3 to 5). Their rate <strong>of</strong> development is influenced by the host<br />
plant, cotton being superior to squash. On cotton and squash it takes an<br />
average <strong>of</strong> 4.5 and 6.7 days respectively to the adult stage at about 27¡C:<br />
there is then a period <strong>of</strong> about 2 days before nymphs are produced. In this<br />
series <strong>of</strong> experiments females on cotton produced an average <strong>of</strong> 27 nymphs<br />
(range 9 to 43), whereas those on squash produced an average <strong>of</strong> 14 (range 2
Host plants<br />
Damage<br />
4.4<br />
Aphis gossypii<br />
to 35) (Khalifa and Sharaf 1964). Life history data on cucumber is provided<br />
by van Steenis and El Khawass (1995).<br />
In U.K., apterae lived 16 days and each produced about 40 <strong>of</strong>fspring. In<br />
founding colonies without competition, the 40 progeny could be produced in<br />
about 7 days and the total population increased about 10 fold each<br />
subsequent week. The rate was reduced as crowding occurred and only then<br />
was it possible for the rate <strong>of</strong> parasitoid increase to exceed that <strong>of</strong> the aphid<br />
(Hussey and Bravenboer 1971).<br />
A. gossypii is widely polyphagous. Cotton, in particular, can carry very<br />
heavy infestations, as also can various cucurbits (e.g. cucumber, squash,<br />
watermelon). In many parts <strong>of</strong> the world it is one <strong>of</strong> the most serious <strong>of</strong> the<br />
aphids on citrus. A. gossypii also infests beans, egg plant, guava, mango,<br />
okra, paprika, potato, taro and numerous ornamentals. In Central and South<br />
America it also damages c<strong>of</strong>fee and cocoa.<br />
As its common and scientific names imply, cotton can be seriously damaged<br />
by A. gossypii.<br />
It can be a major problem and even cause death <strong>of</strong> the plant at<br />
early stages <strong>of</strong> growth; and a further serious attack may occur when the plant<br />
is near maturation and copious production <strong>of</strong> honeydew can contaminate the<br />
cotton lint.<br />
On all <strong>of</strong> its many hosts, severely attacked leaves curl and young growth<br />
is stunted. As populations build up, the upper surface <strong>of</strong> leaves and fruit<br />
becomes contaminated with honeydew, leading to growth <strong>of</strong> sooty moulds,<br />
which is unsightly and interferes with photosynthesis.<br />
For many crops, virus transmission is far more important than direct<br />
damage, since even small numbers <strong>of</strong> migrating aphids can cause serious<br />
problems, whereas even large colonies may cause only moderate leaf<br />
deformation (Barbagallo and Patti 1983). Although, formerly, it was not an<br />
effective vector <strong>of</strong> citrus tristeza virus, it has now become a dangerous one in<br />
USA and Israel. Both adults and nymphs can transmit the virus (Komazaki<br />
1993). A. gossypii is also an important vector <strong>of</strong> a very wide range <strong>of</strong> other<br />
plant viruses.<br />
47
48 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Natural enemies<br />
Two groups <strong>of</strong> hymenopterous parasitoids attack (but are restricted to)<br />
aphids, a larger one consisting <strong>of</strong> species belonging to the family Aphidiidae<br />
and a smaller group belonging to the family Aphelinidae. Both groups occur<br />
worldwide as solitary endoparasitoids. Although many <strong>of</strong> the species are<br />
recorded as having an extensive host range, there is almost always a<br />
significant degree <strong>of</strong> host restriction. Hosts are frequently some (but not all)<br />
<strong>of</strong> the species in a particular aphid genus or several closely related genera.<br />
There is good evidence that there are biotypes within some parasitoid<br />
species, since populations from some hosts or some areas parasitise a<br />
narrower range <strong>of</strong> hosts than the species as a whole. There may also be<br />
differences between biotypes in their preference for the host aphid when<br />
feeding on a particular host plant or in a particular habitat. When a parasitoid<br />
is abundant on a preferred host it may occasionally attack a nearby nonpreferred<br />
host, as with Diaeretiella rapae,<br />
a major parasitoid <strong>of</strong> the cabbage<br />
aphid Brevicoryne brassicae,<br />
which has occasionally been recorded from<br />
both A. craccivora and A. gossypii,<br />
but for which it exhibits a low preference<br />
(Dhiman et al. 1983). There are some species (or biotypes <strong>of</strong> species) that<br />
have been found capable <strong>of</strong> generally causing high levels <strong>of</strong> parasitisation <strong>of</strong><br />
A. craccivora.<br />
Those selected by Starù (1967a, b) are shown in bold italics in<br />
table 4.4.1 and might be considered first as potential species for biological<br />
control introductions to areas where they do not already occur. Valuable<br />
reviews <strong>of</strong> the effectiveness <strong>of</strong> aphid parasitoids are provided by Carver<br />
(1989), Hagen and van den Bosch (1968) and Hughes (1989).<br />
Although many coccinellids, syrphids, chrysopids, hemerobiids and a<br />
few predator species from other insect families attack aphids, their impact in<br />
regulating populations is generally regarded as disappointing, although they<br />
must certainly at times limit economic damage. The efficiency <strong>of</strong> a predator<br />
depends upon its searching ability and effectiveness in capturing prey. The<br />
numbers <strong>of</strong> predators seem to be greatest when aphid numbers are already<br />
declining after a peak in abundance and, thus, their apparently great impact<br />
at that time may actually have little significance in population regulation.<br />
Predators can increase rapidly in numbers only after their prey has become<br />
sufficiently abundant, so there is an important time lag between prey and<br />
predator numbers (Hemptinne and Dixon 1991).<br />
Coccinellids have been used successfully for the biological control <strong>of</strong><br />
several, relatively sessile, coccid pests, whereas results have generally been<br />
poor against aphids. One <strong>of</strong> the reasons is that adult coccinellids and their<br />
larvae are poor at capturing other than first instar aphids (Dixon 1989).<br />
Indeed, the survival <strong>of</strong> newly-hatched beetle larvae is very dependent upon
4.4<br />
Aphis gossypii<br />
the abundance <strong>of</strong> young aphids, so there is a need for coccinellids to lay eggs<br />
very early in the development <strong>of</strong> aphid colonies. Oviposition late in aphid<br />
population development may result in older larvae starving from lack <strong>of</strong><br />
food and the comparatively poor searching ability <strong>of</strong> coccinellids for low<br />
aphid populations aggravates the situation (Hemptinne and Dixon 1991).<br />
Another reason is that coccinellids disperse when prey populations fall to<br />
low levels.<br />
Adults <strong>of</strong> most aphidophagous syrphids are attracted to, and lay their<br />
eggs in or close to, large aphid colonies, the number <strong>of</strong> eggs deposited<br />
increasing as aphid density increases (Chandler 1967). Syrphid larvae also<br />
generally become abundant when the aphid colony is already declining. The<br />
larvae <strong>of</strong> the aphidophagous cecidomyiid Aphidoletes aphidimyza appear to<br />
have adequate host specificity to be acceptable for biological control<br />
introductions. The species has a high degree <strong>of</strong> density dependence, kills<br />
more aphids than it consumes and is less affected than many other predators<br />
by insecticides (Meadow et al. 1985).<br />
Chrysopids and hemerobiids are more effective than many other<br />
predators at capturing prey and are likely to be more efficient predators at<br />
low aphid densities.<br />
A particular problem with most predators is that they are highly<br />
polyphagous. They will almost always attack a very wide range <strong>of</strong> nontarget<br />
insects, some <strong>of</strong> which are likely to be <strong>of</strong> environmental concern.<br />
Regulatory authorities responsible for approving import permits to a country<br />
are becoming increasingly reluctant to do so, unless an adequate degree <strong>of</strong><br />
specificity has been demonstrated and this is occasionally possible.<br />
For the above reasons, no attempt has been made to assemble lists <strong>of</strong> the<br />
many generalist predators recorded as attacking (or probably attacking)<br />
A. craccivora or A. gossypii in the field, although a few facts about their<br />
activities are recorded in the segments dealing with individual countries in<br />
order to provide an entry into the literature. Abstracts <strong>of</strong> many additional<br />
papers are available in CABIÕs Review <strong>of</strong> Agricultural Entomology and its<br />
predecessor Review <strong>of</strong> Applied Entomology, Series A. The major<br />
parasitoids <strong>of</strong> A. gossypii are listed in Table 4.4.1.<br />
49
Table 4.4.1<br />
Parasitoids <strong>of</strong> Aphis gossypii<br />
Country Reference<br />
HYMENOPTERA<br />
APHELINIDAE<br />
Aphelinus abdominalis<br />
China Shi 1980<br />
(= Aphelinus sp. nr flavipes)<br />
Guam<br />
Fulmek 1956<br />
India<br />
Ramaseshiah & Dharmadhikari 1969<br />
(= Aphelinus flavipes)<br />
Shanghai Shi 1980<br />
Aphelinus asychis<br />
Italy Ferrari & Nicoli 1994<br />
Aphelinus chaoniae<br />
Italy Ferrari & Nicoli 1994<br />
Aphelinus gossypii (= Aphelinus kashmiriensis)<br />
Australia<br />
Carver et al. 1993<br />
Cook Is<br />
Walker & Deitz 1979<br />
Hawaii<br />
Timberlake 1924; Yoshimoto 1965<br />
India<br />
Bhat 1987<br />
Japan<br />
Takada & Tokomaku 1996<br />
Tonga<br />
Carver et al. 1993;<br />
Stechmann & Všlkl 1988<br />
Aphelinus humilis<br />
Australia M. Carver pers. comm.<br />
Aphelinus mali<br />
China<br />
Shi 1985<br />
India<br />
Ramaseshiah & Dharmadhikari 1969<br />
Senegal<br />
Risbec 1951; Fulmek 1956<br />
Shanghai<br />
Shi 1980<br />
Taiwan<br />
Takada 1992<br />
Trinidad<br />
Bennett 1985<br />
Aphelinus paramali<br />
Israel Zehavi & Rosen 1988<br />
Aphelinus semiflavus<br />
USA Hartley 1922; Spencer 1926;<br />
Oatman et al. 1983b; Trumble & Oatman 1984<br />
Aphelinus varipes (= A. nigritus)<br />
Transcaucusus<br />
Fulmek 1956<br />
USA<br />
Wharton 1983<br />
50 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong>
Table 4.4.1 (contÕd)<br />
HYMENOPTERA<br />
APHELINIDAE (contÕd)<br />
Aphelinus sp.<br />
2 ´ spp.<br />
APHIDIIDAE<br />
* Aphidius colemani<br />
Aphidius ervi<br />
Aphidius floridaensis<br />
Parasitoids <strong>of</strong> Aphis gossypii<br />
Country Reference<br />
Colombia<br />
India<br />
Japan<br />
Angola<br />
Argentina<br />
Australia<br />
Chile<br />
China<br />
Egypt<br />
India<br />
Japan<br />
Kenya<br />
Mozambique<br />
Pakistan<br />
RŽunion<br />
Tonga<br />
Uruguay<br />
Venezuela<br />
Morocco<br />
Uzbekistan<br />
USA, West Indies Starù 1967a,b<br />
Ramirez & Zuluaga 1995<br />
Ramaseshiah & Dharmadhikari 1969<br />
Takada 1992<br />
Starù & van Harten 1972<br />
Starù 1967a, 1972<br />
Carver & Starù 1974; Room & Wardhaugh 1977<br />
Prado 1991, Starù 1975<br />
Xi & Zhu 1984<br />
Selim et al. 1987<br />
Starù 1972; Agarwala et al. 1981<br />
Starù 1967a<br />
Starù & Schmutterer 1973<br />
Starù & van Harten 1972<br />
Starù 1975<br />
Starù 1975<br />
Carver et al. 1993<br />
Starù 1975<br />
Cermeli 1989<br />
Fulmek 1956<br />
Starù 1979<br />
4.4<br />
Aphis gossypii<br />
51
Table 4.4.1 (contÕd)<br />
HYMENOPTERA<br />
APHIDIIDAE (contÕd)<br />
* Aphidius gifuensis<br />
China<br />
Hawaii<br />
India<br />
Japan<br />
Korea<br />
Taiwan<br />
Aphidius urticae (= Aphidius lonicerae)<br />
Bšrner et al. 1957<br />
Aphidius matricariae (= Aphidius phorodontis)<br />
Brazil<br />
Canada<br />
Chile<br />
Germany<br />
India<br />
Italy<br />
Lebanon<br />
Pakistan<br />
Peru<br />
Tunisia<br />
USA<br />
Aphidius picipes<br />
Aphidius similis<br />
Aphidius sonchi<br />
Aphidius uzbekistanicus<br />
Parasitoids <strong>of</strong> Aphis gossypii<br />
Country Reference<br />
Shi 1980; Takada 1992<br />
Mackauer & Starù 1967; Takada 1968<br />
Raychaudhuri 1990<br />
Mackauer & Starù 1967;<br />
Takada 1968, 1992<br />
Mackauer & Starù 1967; Takada 1992<br />
Mackauer & Starù 1967<br />
Starù 1967a<br />
Starù 1967a<br />
Prado 1991<br />
Mackauer 1962b<br />
Agarwala et al. 1981; Agarwala 1983<br />
Starù 1976<br />
Tremblay et al. 1985<br />
Starù & Ghosh 1983<br />
Starù 1967a<br />
Halima-Kamel 1993<br />
Starù 1967a<br />
Bšrner et al. 1957<br />
China Li & Wen 1988; Xi & Zhu 1984<br />
India Agarwala et al. 1981<br />
Bšrner et al. 1957<br />
India Raychaudhuri 1990; Takada 1992<br />
52 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong>
Table 4.4.1 (contÕd)<br />
HYMENOPTERA<br />
APHIDIIDAE (contÕd)<br />
Aphidius spp. India Agarwala et al. 1981<br />
Cristicaudus nepalensis<br />
India Raychaudhuri 1990; Takada 1992<br />
Diaeretiella rapae<br />
Ephedrus nacheri<br />
* Ephedrus persicae<br />
Ephedrus plagiator<br />
* Lipolexis gracilis<br />
Parasitoids <strong>of</strong> Aphis gossypii<br />
Country Reference<br />
India, Japan<br />
Tunisia<br />
USA<br />
Uzbekistan<br />
Agarwala et al. 1981; Takada 1992<br />
Halima-Kamel 1993<br />
Starù 1967a<br />
Starù 1979<br />
China, Japan, Europe Takada 1968, 1992<br />
India<br />
Iraq<br />
Korea<br />
Lebanon<br />
Taiwan<br />
USA<br />
USSR<br />
India<br />
Uzbekistan<br />
Japan,<br />
Korea, Taiwan<br />
USSR<br />
China<br />
Europe<br />
Hong Kong<br />
India<br />
Japan<br />
Lebanon<br />
Shanghai<br />
Taiwan<br />
Agarwala et al. 1981, Takada 1992<br />
Al-Azawi 1970<br />
Takada 1972b; Chou 1981; Paik 1975<br />
Tremblay et al. 1985<br />
Chou 1981b<br />
Schlinger & Hall 1960<br />
Starù 1970<br />
Raychaudhuri 1990<br />
Starù 1979<br />
Takada 1992<br />
Paik 1975; Chou 1981b<br />
Starù 1970<br />
Shi 1980; Xi & Zhu 1984; Takada 1992,<br />
Starù 1970<br />
Takada 1992<br />
Raychaudhuri 1990<br />
Takada 1992<br />
Tremblay et al. 1985<br />
Shi 1980<br />
Chou 1981b, Takada 1992<br />
4.4<br />
Aphis gossypii<br />
53
Table 4.4.1 (contÕd) Parasitoids <strong>of</strong> Aphis gossypii<br />
HYMENOPTERA<br />
APHIDIIDAE (contÕd)<br />
Lipolexis scutellaris (= Lipolexis pseudoscutellaris) Hong Kong<br />
India<br />
Malaysia<br />
Philippines<br />
Vietnam<br />
Lysaphidus schimitscheki India Raychaudhuri 1990<br />
*Lysiphlebia japonica China, Japan<br />
Korea, Taiwan<br />
Country Reference<br />
Takada 1992<br />
Starù & Ghosh 1975; Agarwala et al. 1981; Pramanik &<br />
Raychaudhuri 1984; Raychaudhuri 1990; Takada 1992<br />
Ng & Starù 1986; Takada 1992;<br />
V.J. Calilung pers. comm. 1995<br />
Starù & Zelenù 1983<br />
Takada 1968, 1992; Paik 1975; Chou 1981; Tian et al.<br />
1981; Xi & Zhu 1984<br />
Lysiphlebia mirzai Vietnam Starù & Zelenù 1983; Takada 1992<br />
*Lysiphlebus fabarum<br />
Algeria, Bulgaria, Corsica, Starù et al. 1975; Starù 1976<br />
(= Lysiphlebus ambiguus<br />
Israel, Italy<br />
= Lysiphlebus cardui<br />
Egypt<br />
Selim et al. 1987<br />
= Lysiphlebus confusus)<br />
Europe<br />
Starù 1970<br />
Greece<br />
Santas 1978<br />
Iraq<br />
Al-Azawi 1966<br />
Japan<br />
Takada 1992<br />
Lebanon<br />
Tremblay et al. 1985<br />
Morocco<br />
Starù 1967a<br />
Pakistan<br />
Hamid et al. 1977<br />
Tunisia<br />
Halima-Kamel 1993<br />
USA<br />
Starù 1967a<br />
USSR<br />
Starù 1967a; Lyashova 1992<br />
Uzbekistan<br />
Starù 1979<br />
Lysiphlebus shaanxiensis China Chou & Xiang 1982<br />
54 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong>
Table 4.4.1 (contÕd) Parasitoids <strong>of</strong> Aphis gossypii<br />
HYMENOPTERA<br />
APHIDIIDAE (contÕd)<br />
*Lysiphlebus testaceipes<br />
Chile<br />
Colombia<br />
Cuba<br />
France<br />
Guadeloupe<br />
Haiti<br />
Hawaii<br />
Italy<br />
Mexico<br />
Spain<br />
Portugal<br />
Trinidad<br />
USA<br />
Venezuela<br />
West Indies<br />
Lysiphlebus sp. Argentina<br />
Colombia<br />
Hawaii<br />
India<br />
Pakistan<br />
Portugal<br />
USA<br />
Country Reference<br />
Prado 1991<br />
Fulmek 1956; Vergara & Galeano 1994<br />
Starù 1967b, 1981<br />
Starù et al. 1988a,b<br />
Starù et al. 1987<br />
Fulmek 1956<br />
Starù 1967a<br />
Tremblay & Barbagallo 1982<br />
Starù & Remaudi re 1982<br />
Starù et al. 1988a,b;<br />
Costa & Starù 1988; Starù et al. 1988c<br />
Bennett 1985<br />
Spencer 1926; Schlinger & Hall 1960; Starù 1970;<br />
Oatman et al. 1983b; Trumble & Oatman 1984<br />
Cermeli 1989<br />
Starù 1967b<br />
Fulmek 1956<br />
Ramirez & Zuluaga 1995<br />
Fulmek 1956<br />
Agarwala et al. 1981<br />
Mohyuddin & Anwar 1972, 1973<br />
Boelpaepe et al. 1992<br />
Fulmek 1956<br />
Praon abjectum India Raychaudhuri 1990; Takada 1992<br />
Praon absinthii India Agarwala et al. 1981<br />
Praon exsoletum Uzbekistan Starù 1979<br />
4.4 Aphis gossypii 55
Table 4.4.1 (contÕd) Parasitoids <strong>of</strong> Aphis gossypii<br />
Country Reference<br />
HYMENOPTERA<br />
APHIDIIDAE (contÕd)<br />
Praon myzophagum India Agarwala et al. 1981<br />
Praon volucre<br />
Lebanon<br />
Tremblay et al. 1985<br />
Tajikistan, Uzbekistan Starù 1979<br />
Praon sp.<br />
India<br />
Agarwala et al. 1981<br />
Uzbekistan<br />
Starù 1979<br />
Toxares macrosiphophagum India Raychaudhuri 1990; Takada 1992<br />
Trioxys acalephae<br />
China<br />
Takada 1992,<br />
India<br />
Agarwala et al. 1981<br />
Trioxys angelicae<br />
Corsica, Greece, Iraq, Italy, Starù 1976; Santas 1978<br />
Israel, Morocco<br />
Al-Azawi 1970<br />
Israel<br />
Rosen 1967b<br />
Lebanon<br />
Hussein & Kawar 1984; Tremblay et al. 1985<br />
Tunisia<br />
Halima-Kamel 1993<br />
Trioxys asiaticus Iran, Tajikistan, Uzbekistan Starù 1979<br />
Trioxys auctus<br />
USSR<br />
Uzbekistan<br />
Fulmek 1956; Mackauer & Starù 1967<br />
Starù 1979<br />
Trioxys basicurvus India Agarwala et al. 1981; Raychaudhuri 1990; Takada 1992<br />
Trioxys communis China<br />
Japan, Korea, Taiwan<br />
Philippines<br />
Shi 1980, 1985<br />
Paik 1976; Chou 1981a; Lu & Lee 1987; Takada 1992<br />
V.J. Calilung pers. comm. 1995<br />
Trioxys complanatus Uzbekistan Starù 1979<br />
Trioxys equatus India Samanta et al. 1985; Raychaudhuri 1990<br />
56 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong>
Table 4.4.1 (contÕd) Parasitoids <strong>of</strong> Aphis gossypii<br />
HYMENOPTERA<br />
APHIDIIDAE (contÕd)<br />
*Trioxys indicus China<br />
Tian et al. 1981, Xi & Zhu 1984<br />
India<br />
Starù & Ghosh 1975; Agarwala et al. 1981; Agarwala<br />
1983,1988; Raychaudhuri 1990; Takada 1992<br />
Taiwan<br />
Chou 1981b, Takada 1992<br />
Trioxys nr pallidus Morocco Fulmek 1956<br />
Trioxys rietscheli China, India Shi 1980; Raychaudhuri 1990<br />
Trioxys rubicola India Agarwala et al. 1981<br />
Trioxys sinensis Pakistan Mohyuddin et al. 1972<br />
Trioxys sp.<br />
Portugal<br />
Taiwan, USA<br />
ENCYRTIDAE<br />
Aphidencyrtus sp. Malaysia Yunus & Ho 1980<br />
PTEROMALIDAE<br />
Pachyneuron aphidis China Shi 1987<br />
DIPTERA<br />
CECIDOMYIIDAE<br />
Aphidoletes aphidimyza Chile, Europe,<br />
Nth America<br />
Endaphis maculans Trinidad<br />
USA<br />
Country Reference<br />
Bšrner et al. 1957; Boelpaepe et al. 1992<br />
Fulmek 1956<br />
Harris 1973; Kocourek et al. 1993,<br />
Meadow et al. 1985; Prado 1991<br />
Kirkpatrick 1954<br />
Tang et al. 1994; Yokomi et al. 1994<br />
4.4 Aphis gossypii 57
Table 4.4.1 (contÕd) Parasitoids <strong>of</strong> Aphis gossypii<br />
ACARINA<br />
Country Reference<br />
TROMBIDIIDAE<br />
Allothrombium pulvinum China Dong et al. 1992<br />
Xu et al. 1993<br />
Zhang et al. 1993; Zhang & Chen 1993<br />
*Starù (1967a, 1970) selected these species (bold type) for possible introduction to areas where they do not occur.<br />
Also recorded from Aphis craccivora<br />
58 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong>
4.4 Aphis gossypii 59<br />
Under humid conditions, high aphid mortality may result from fungal<br />
infection. The two species commonly reported are Neozygites fresenii and<br />
Cephalosporium (= Verticillium) lecanii (Table 4.4.2), although about a<br />
dozen species may be involved (Hagen and van den Bosch 1968). Effective<br />
use <strong>of</strong> the above fungi has been made under glasshouse conditions and<br />
V. lecanii is available commercially for this purpose. However, in Florida<br />
this fungus has performed poorly on A. gossypii compared with against<br />
Myzus persicae (Osborne et al. 1994).<br />
Table 4.4.2 Fungi attacking Aphis gossypii and/or A. craccivora<br />
Country Reference<br />
Arthrobotrys sp. USA OÕBrien et al. 1993<br />
Beauveria bassiana USSR Pavlyushin & Krasavina 1987<br />
Cephalosporium lecanii Japan<br />
Netherlands<br />
USA<br />
USSR<br />
Venezuela<br />
Masuda & Kikuchi 1992;<br />
Saito 1988<br />
Yokomi & Gottwald 1988;<br />
Sopp et al. 1990; Vehrs &<br />
Parrella 1991; Schelt 1993<br />
Cermeli 1989<br />
Pavlyushin & Krasavina 1987<br />
Entomophthora exitialis India Kranz et al. 1977<br />
Entomophthora sp. Chile Prado 1991<br />
Neozygites fresenii Australia<br />
Chad<br />
China<br />
Cuba<br />
Milner & Holdom 1986<br />
Silvie & Papierok 1991<br />
Zhang 1987<br />
Hernandez & Alvarez 1985<br />
USA Steinkraus et al. 1991, 1995,<br />
1996; OÕBrien et al. 1993;<br />
Steinkraus & Slaymaker 1994;<br />
Smith & Hardee 1996<br />
Paecilomyces fumosoroseus USSR Pavlyushin & Krasavina 1987
60 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Attempts at biological control<br />
There have been many intentional and unintentional transfers <strong>of</strong> aphid<br />
parasitoids, which have influenced the populations <strong>of</strong> A. craccivora and<br />
A. gossypii in different regions <strong>of</strong> the world. However, the majority <strong>of</strong><br />
intentional transfers have been aimed at other target aphid pests. Some<br />
deliberate attempts against these two species have been unsuccessful (Table<br />
4.4.3). Overall, however, there is little doubt that, where parasitoids have<br />
become established, the situation is better, sometimes significantly better,<br />
than if they were not present, even if the level <strong>of</strong> control may not be as<br />
effective as is desirable. There is little doubt that, in many regions, an<br />
improved situation is likely to occur if additional parasitoid species are<br />
established.<br />
AUSTRALIA<br />
Aphidius colemani is capable <strong>of</strong> producing rapid decreases in populations <strong>of</strong><br />
A. gossypii on cotton and, <strong>of</strong> carrying this to extinction in association with<br />
high densities <strong>of</strong> Harmonia octomaculata (= Coccinella arcuata) and<br />
Coccinella transversalis (= C. repanda) (Room and Wardhaugh 1977).<br />
Three species <strong>of</strong> parasitoid were imported in the hope that, as polyphagous<br />
species, they would contribute to the biological control <strong>of</strong> several species <strong>of</strong><br />
pest aphid. The principal target for two <strong>of</strong> the species was A. craccivora,<br />
which is very sporadic in occurrence in Australia. It was hoped that, in the<br />
unpredictable absence <strong>of</strong> A. craccivora, the parasitoids would continue to<br />
breed and survive in other hosts.<br />
Lysiphlebus testaceipes (from Aphis nerii in California) and L. fabarum<br />
(from Greece and Turkey) were imported as biological control agents <strong>of</strong><br />
A. craccivora on legumes, mass reared, and released in 1982 and 1983 in<br />
New South Wales and Victoria. Both parasitoids readily parasitised<br />
A. craccivora, A. gossypii and some other aphid species in the laboratory.<br />
The releases coincided with a prolonged drought during which there were no<br />
A. craccivora available on legume crops. Releases were, therefore, made on<br />
A. gossypii infesting Hibiscus bushes. No parasites were recovered the<br />
following year from either A. craccivora or A. gossypii, although<br />
L. testaceipes became established in Aphis nerii on oleander (Nerium)<br />
(Hughes 1989). L. fabarum was not recovered. It was concluded that the<br />
parasitoids must have been unsuitable biotypes (Carver 1984, 1989).
Table 4.4.3 Releases for the biological control (inter alia) <strong>of</strong> Aphis craccivora and/or A. gossypii<br />
Parasitoid From To Year Result Reference<br />
Aphelinus varipes South Carolina California + Wharton 1983<br />
Aphelinus abdominalis India U.K. + Hussey & Bravenboer 1971<br />
Aphidius colemani S. Brazil France 1982 + Rabasse 1986<br />
Australia Tonga 1990 + Carver et al. 1993;<br />
Wellings et al. 1994<br />
Lysiphlebus fabarum France, Italy, Greece, Australia 1982,<br />
- Carver 1984, 1989<br />
Turkey<br />
1983<br />
Lysiphlebus testaceipes USA<br />
China<br />
1983<br />
+ Zheng & Tang 1989<br />
USA<br />
Hawaii 1923<br />
+ Beardsley 1961<br />
USA<br />
India<br />
1966<br />
? Ramaseshiah et al. 1969;<br />
Sankaran 1974<br />
USA<br />
Australia 1982<br />
+ Carver 1984; Hughes 1989<br />
Cuba<br />
France, Italy 1973<br />
+ Rabasse 1986; Starù et al. 1988a,b<br />
Czechoslovakia Tonga<br />
- Stechmann & Všlkl 1988; Všlkl et al.<br />
1990<br />
Hawaii<br />
Pakistan 1972<br />
? Anwar 1974;<br />
Mohyuddin & Anwar 1972, 1973,<br />
Mohyuddin et al. 1971<br />
Mexico<br />
USSR<br />
1989<br />
+ Shiiko et al. 1991<br />
USA<br />
China<br />
1983<br />
+ Zheng & Tang 1989<br />
Trioxys indicus India Australia 1986 - Carver 1989; Sandow 1986<br />
4.4 Aphis gossypii 61
62 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
CHINA<br />
Praon volucre was imported from the Mediterranean area for the<br />
biological control <strong>of</strong> Hyperomyzus lactucae, a vector <strong>of</strong> lettuce necrotic<br />
yellows. It was mass reared and released in 1981 and 1982, mainly in New<br />
South Wales and Victoria. It has not been recovered from mainland<br />
Australia, but is reported to be present in Tasmania. In laboratory trials it<br />
successfully parasitised a number <strong>of</strong> pest aphids that occur in Australia,<br />
including A. craccivora.<br />
In 1986 Trioxys indicus was introduced from India and released in<br />
Western Australia, Victoria and New South Wales against A. craccivora<br />
(Carver 1989), but establishment did not occur.<br />
Although he gives no further details Mohammad (1979) states that<br />
parasitisation <strong>of</strong> A. craccivora in Adelaide soon after colonisation<br />
frequently prevented the establishment <strong>of</strong> a colony.<br />
Zhang (1992) recorded for A. gossypii on cotton in China more than 48<br />
species <strong>of</strong> natural enemy (belonging to 19 families in 9 orders). Coccinellids,<br />
spiders and lacewings were the most important predators <strong>of</strong> this and other<br />
pests in cotton fields. Nan et al. (1987) record attack on A. gossypii and other<br />
cotton pests by 5 species <strong>of</strong> pentatomid, 9 species <strong>of</strong> coccinellid, 4 species <strong>of</strong><br />
lacewing and 36 species <strong>of</strong> spider. The dominant predators studied by Wu<br />
(1986) were found to vary according to the season and included the<br />
coccinellids Coccinella septempunctata, Hippodamia variegata, Propylea<br />
japonica, Harmonia axyridis, the lacewings Chrysopa (= Chrysoperla)<br />
sinica, Chrysopa formosa, C. pallens (= C. septempunctata), C. intima and<br />
the spiders Erigonidium graminicolum, Misumenops tricuspidatus and<br />
Xysticus croceus. Zhang (1985) carried out laboratory tests on the daily<br />
consumption <strong>of</strong> A. gossypii by Scymnus h<strong>of</strong>fmanni, Chrysopa sinica and the<br />
spider Erigonidium graminicolum. Ma and Liu (1985) reported on the<br />
effectiveness and seasonal fluctuations <strong>of</strong> Propylea japonica and Yang<br />
(1985b) on its laboratory rearing. Ding and Chen (1986) examined the<br />
predation pattern <strong>of</strong> Chrysopa sinica. Propylea japonica, Scymnus<br />
h<strong>of</strong>fmanni and spiders (especially Theridion octomaculatum and<br />
Erigonidium graminicolum) were major enemies <strong>of</strong> A. gossypii on cotton in<br />
Hunan Province. The spiders (2.6 to 26 per 100 plants ) were present from<br />
late June to late August and were relatively unaffected by the weather.<br />
Coccinellid populations fluctuated somewhat with the season. Reproduction<br />
<strong>of</strong> A. gossypii was inhibited and its damage reduced when the ratio <strong>of</strong> total<br />
natural enemies to aphids was 1:50 or the ratio <strong>of</strong> coccinellids was 1:140.<br />
Since 1978, cotton fields over large areas have not been treated with<br />
insecticides before August, in order to safeguard the natural enemies which<br />
now hold the aphids in check (Mao and Xia 1983). Zhao et al. (1989) report
4.4 Aphis gossypii 63<br />
that the lycosid spider Pardosa astrigata is an important predator in cotton<br />
fields and Dong (1988) that the coccinellid Harmonia axyridis was an<br />
effective natural enemy when present in adequate numbers. Other predators<br />
include the anthocorid bug Orius minutus (Miao & Sun 1987), the<br />
coccinellids Scymnus h<strong>of</strong>fmanni (Zhao and Holling 1986), Propylea<br />
japonica and Harmonia axyridis (Zou et al. 1986; Lei et al. 1987 ).<br />
The aphidiid Aphidius picipes (= A. avenae) parasitised more than 80%<br />
<strong>of</strong> A. gossypii individuals on Chinese cabbage growing near cotton fields (Li<br />
& Wen 1988). Laboratory studies showed that Trioxys communis was more<br />
effective than Aphelinus mali in suppressing A. gossypii populations (Shi<br />
1985). Of the 5 species <strong>of</strong> parasitoid recorded by Xi and Zhu (1984) on<br />
A. gossypii on cotton in Jiangsu Province, Lysiphlebia japonica and Trioxys<br />
indicus were dominant and each accounts for about 45% <strong>of</strong> all parasitoids. In<br />
the laboratory, female L. japonica laid an average <strong>of</strong> 120 eggs and produced<br />
a parasitisation rate <strong>of</strong> up to 14%. In the field it overwinters as larvae inside<br />
A. gossypii, A. craccivora or Myzus persicae. In an earlier study Tian et al.<br />
(1981) recorded the same two parasitoids on both A. craccivora and<br />
A. gossypii with a combined parasitisation rate <strong>of</strong> 13%. This did not provide<br />
effective control and it was pointed out there was heavy attack by<br />
hyperparasitoids, such as Aphidencyrtus sp. (Encyrtidae).<br />
Larvae <strong>of</strong> the mite Allothrombium pulvinum have been observed<br />
attacking A. gossypii in cotton fields (Zhang and Chen 1993; Zhang et al.<br />
1993).<br />
COLOMBIA<br />
Aphelinus sp. caused 2.2% and Lysiphlebus sp. 0.3% parasitisation <strong>of</strong><br />
A. gossypii on cotton in the field (Ramirez and Zuluaga 1995).<br />
CUBA<br />
Aphis craccivora is a common pest <strong>of</strong> vegetables and many other crops and<br />
also occurs on wayside trees, such as Gliricidia. In beans and other annual<br />
crops it occurs for a short period only, whereas on wayside trees it is present<br />
more or less continuously. The native parasitoid Lysiphlebus testaceipes<br />
parasitises the aphid heavily on Gliricidia, but poorly or not at all on young<br />
beans. This appears to be primarily a matter <strong>of</strong> the relative rates <strong>of</strong> dispersal<br />
<strong>of</strong> the aphid and its parasitoid (Starù 1970).<br />
EAST ASIA<br />
A list <strong>of</strong> parasitoids recorded from A. gossypii in East Asia is shown in table<br />
4.4.1 in which most entries from this region are based on Takada (1992). The<br />
most comprehensive information within this region is available from<br />
Taiwan (Starù and Schlinger 1967; Tao and Chiu 1971; Chou 1984), Japan<br />
(Takada 1968; Takada and Yamauchi 1979; Takada unpublished) and India<br />
(Raychaudhuri 1990). For <strong>Southeast</strong> Asia, there are two records from
64 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
FRANCE<br />
INDIA<br />
Vietnam and one from Malaysia. In so far as one can argue from such scanty<br />
data, the principal parasitoids <strong>of</strong> A. gossypii in the Far East (Trioxys<br />
communis, Lysiphlebia japonica and the particular species involved <strong>of</strong><br />
Aphelinus) do not occur in India (Takada 1992). According to Takada, the<br />
principal species attacking A. gossypii in India is Trioxys indicus, which is<br />
recorded from Taiwan, but apparently not from Japan. Another species<br />
attacking A. gossypii in India is Lipolexis scutellaris, which also occurs in<br />
Vietnam, Malaysia and Hong Kong, but is not recorded elsewhere in the Far<br />
East (Takada 1992). Two widely distributed and effective parasitoids that<br />
attack A. gossypii in other parts <strong>of</strong> the world (Aphidius colemani and<br />
Lysiphlebus testaceipes) do not yet seem to be present in <strong>Southeast</strong> Asia,<br />
although A. colemani is recorded from the field in Pakistan (Starù 1975).<br />
Takada (1992) comments on the habitat specialisations in Japan <strong>of</strong><br />
parasitoids <strong>of</strong> A. gossypii, which occurs in both open and lightly wooded<br />
habitats: Trioxys communis, and Aphelinus species prefer the open habitat,<br />
whereas Lysiphlebia japonica, Ephedrus nacheri, E. persicae and<br />
E. plagiator prefer the lightly-wooded habitat. Thus, the parasitoid complex<br />
on A. gossypii on cucumber, egg plant or taro is quite different from that on<br />
Hibiscus or Rhamnus in a garden.<br />
Starù et al. (1973) reviewed the parasitoids <strong>of</strong> aphids in France. A South<br />
American strain <strong>of</strong> Aphidius colemani, which is adapted to warm subtropical<br />
conditions and is highly polyphagous, was introduced from southern Brazil<br />
and released against Toxoptera aurantii in France near Antibes in 1982. It is<br />
reported to be established (Rabasse 1986; Tardieux and Rabasse 1986). In<br />
1973Ð74 Lysiphlebus testaceipes was introduced into France and released<br />
near Antibes and in Corsica. It was recovered soon after, and later in Italy. It<br />
was also sent to eastern Spain where it established and spread to become the<br />
predominant parasitoid in the regions where it occurs. It attacks A. gossypii<br />
on citrus and a number <strong>of</strong> other aphids on other host plants (Starù et al.<br />
1988a).<br />
Including the widespread Trioxys indicus, 14 parasitoids were recorded from<br />
A. gossypii (a preferred host) and 8 from A. craccivora (Agarwala 1983).<br />
Ephedrus persicae is reported to be confined in India to A. craccivora and 3<br />
parasitoids, Praon absinthii, Trioxys basicurvus and T. rubicola confined to<br />
the A. gossypii complex (Agarwala et al. 1981).<br />
The impact <strong>of</strong> the widespread parasitoid Trioxys indicus on<br />
A. craccivora feeding on pigeon pea was studied in the laboratory and the<br />
field. The parasitoid had a high searching ability and exhibited a density<br />
dependent relationship with its host. A single female oviposited in 100 to<br />
150 aphids in 3 to 5 days after emergence and the life cycle occupied 15 to 20
IRAQ<br />
ISRAEL<br />
4.4 Aphis gossypii 65<br />
days. Early in the season 9.4% <strong>of</strong> the aphids were parasitised, rising to a peak<br />
<strong>of</strong> 64.6% two months later and resulting in suppression <strong>of</strong> the aphid<br />
population. Trioxys indicus has a relatively narrow host range, which<br />
includes A. gossypii, A. craccivora and the oleander aphid A. nerii. Up to<br />
2.4% <strong>of</strong> the parasitoids were hyperparasitised by a cynipoid wasp. Singh and<br />
Sinha (1983) concluded from these studies that T. indicus had most <strong>of</strong> the<br />
necessary attributes <strong>of</strong> a potentially effective biological control agent for<br />
A. craccivora and A. gossypii.<br />
Ramaseshiah and Dharmadhikari (1969) found Aphelinus abdominalis<br />
(= A. flavipes) to be one <strong>of</strong> the important parasitoids <strong>of</strong> A. gossypii in India;<br />
furthermore, that Aphelinus sp. nr abdominalis had A. craccivora as a<br />
preferred host, but also attacked A. gossypii.<br />
Many generalist predators attack A. gossypii and other aphids in India,<br />
and Agarwala and Saha (1986) record 8 species attacking it there on cotton.<br />
Seven species <strong>of</strong> predatory coccinellid, 2 syrphids and a chrysopid were<br />
recorded preying on A. gossypii on potatoes. Coccinella septempunctata and<br />
Cheilomenes (= Menochilus) sexmaculata were the most important (Raj<br />
1989). A. gossypii was reported to be controlled on sunflower by<br />
coccinellids (Goel and Kumar 1990). The aphidophaghous coccinellid<br />
Micraspis discolor showed a preference for A. craccivora (Agarwala et al.<br />
1988).<br />
The biology <strong>of</strong> the predatory Leucopis species (Diptera) attacking both<br />
A. craccivora and A. gossypii on chrysanthemums is described by Kumar et<br />
al. (1988). Larvae <strong>of</strong> Chrysopa orestes had a substantial effect on A. gossypii<br />
on eggplant (Bhagat and Masoodi 1986).<br />
Raychaudhuri et al. (1979) record three predatory spiders: Cyclosa<br />
insulana (Araneidae) attacking A. craccivora and A. gossypii; Theridion sp.<br />
(Theridiidae) attacking A. craccivora; and Uloborus sp. (Uloboridae)<br />
attacking A. gossypii.<br />
A. gossypii is parasitised on melons, cotton and okra by Lysiphlebus<br />
fabarum which, in turn, is attacked by the hyperparasitoids Pachyneuron<br />
aphidis (Pteromalidae), Dendrocerus (= Lygocerus) sp. (Megaspilidae),<br />
Aphidencyrtus sp. (Encyrtidae) and Alloxysta (= Charips) sp. (Charipidae)<br />
(Al-Azawi 1966).<br />
A. gossypii occasionally infests citrus in the vicinity <strong>of</strong> cotton fields. Based<br />
on small samples, it was found to be attacked by only one parasitoid, Trioxys<br />
angelicae (Rosen 1967a,b).
66 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
ITALY<br />
A. gossypii in particular, but also A. craccivora, are two <strong>of</strong> the 10 aphid<br />
species attacking citrus. Eleven species <strong>of</strong> aphidiine parasitoids provide a<br />
considerable measure <strong>of</strong> biological control (Tremblay 1980). Lysiphlebus<br />
testaceipes (particularly) and L. fabarum are the commonest species and<br />
together may attain a parasitisation rate <strong>of</strong> 90% to 100%.<br />
The most important predators are coccinellids, <strong>of</strong> which Scymnus spp.<br />
are common amongst colonies <strong>of</strong> A. gossypii. Together with Coccinella<br />
septempunctata and other natural enemies, they may quickly suppress a<br />
cotton aphid population. Chrysopid, syrphid and cecidomyiid larvae are less<br />
effective although, in the absence <strong>of</strong> coccinellid larvae, syrphid larvae may<br />
be important. A. gossypii is under biological control on citrus in orchards<br />
where pest management procedures are adopted (Barbagallo and Patti 1983;<br />
Starù 1964). Recent papers on natural enemies <strong>of</strong> A. gossypii in Italy have<br />
been published by Ferrari and Burgio (1994) and Ferrari & Nicoli (1994).<br />
JAPAN<br />
Amongst numerous predators reported on A. gossypii by many authors are<br />
Coccinella septempunctata (Nozato and Abe 1988) and Scymnus h<strong>of</strong>fmanni<br />
(Kawauchi 1987).<br />
KOREA<br />
The consumption <strong>of</strong> A. gossypii by larvae <strong>of</strong> the coccinellid Harmonia<br />
axyridis was studied by Choi and Kim (1985).<br />
Eight species <strong>of</strong> parasitoid (and 6 <strong>of</strong> hyperparasitoid) <strong>of</strong> A. craccivora<br />
were reported by Chang and Youn (1983). The more important species (and<br />
rates <strong>of</strong> parasitisation) were Lysiphlebus fabarum (31.6%), Lipolexis<br />
scutellaris (18.8%), Lysiphlebia japonica (16.7%) and Adialytus salicaphis<br />
(11.4%). Of 509 field collected, mummified aphids 44.8% produced<br />
parasitoids and 43.8% hyperparasitoids giving an overall parasitisation rate<br />
<strong>of</strong> 88.6%.<br />
MALAYSIA<br />
The only record <strong>of</strong> parasitoids <strong>of</strong> A. craccivora or A. gossypii appears to be<br />
that <strong>of</strong> Ng and Starù (1986). Lipolexis scutellaris, an oriental species with a<br />
wide distribution in India and extending to Vietnam, southern China and<br />
Taiwan, was recorded from both aphid hosts. Trioxys communis, also an<br />
oriental species, was rarely found, but only on another aphid species (Aphis<br />
spiraecola). At least 3 unidentified species <strong>of</strong> aphelinids were bred and in<br />
large numbers, but no other information on these is provided.<br />
NETHERLANDS<br />
A valuable review <strong>of</strong> the biological control <strong>of</strong> A. gossypii in glasshouses,<br />
with special reference to the situation in the Netherlands, is provided by van<br />
Steenis (1992). van Steenis (1995) evaluated 4 aphidiine parasitoids for<br />
biological control <strong>of</strong> A. gossypii on glasshouse cucumbers. Aphidius<br />
colemani performed the best (72 to 80% parasitisation), followed by
4.4 Aphis gossypii 67<br />
Lysiphlebus testaceipes (26%), Ephedrus cerasicola (23%) and Aphidius<br />
matricariae (less than 5%). The general principles <strong>of</strong> selection and<br />
establishment <strong>of</strong> species are relevant elsewhere, although only the first <strong>of</strong> the<br />
three species selected (Lysiphlebus testaceipes, Aphidius matricariae,<br />
Ephedrus cerasicola) would appear to be particularly relevant to <strong>Southeast</strong><br />
Asia or the Pacific.<br />
PAKISTAN<br />
In investigations from 1967 to 1970, 6 parasitoid species and 22 predator<br />
species were recorded attacking Aphis craccivora on a range <strong>of</strong> leguminous<br />
crops and weeds in 5 climatologically different zones <strong>of</strong> Pakistan (Hamid et<br />
al. 1977). Alate A. craccivora were found on soybean (Glycine max), but<br />
colonies <strong>of</strong> apterous aphids never developed, possibly due to the presence <strong>of</strong><br />
abundant plant hairs.<br />
Aphelinus abdominalis (= A. basalis), which was widespread and active<br />
throughout the year, parasitised from 0.6% to 17.6% <strong>of</strong> A. craccivora, the<br />
level depending upon host plant, location (hills, foothills or plains) and<br />
season. Perhaps due to its small size, A. abdominalis was the only parasite<br />
that attacked A. craccivora under the covering <strong>of</strong> hairs on Phaseolus aureus.<br />
The period from oviposition in the aphid to mummy formation was 3 to 4<br />
days and adult wasps then emerged in 5 days. Trioxys ?sinensis parasitised<br />
up to 21.3% <strong>of</strong> aphids. Development from oviposition in the aphid to<br />
mummy formation took 14 to 16 days and adults emerged 4 to 7 days later.<br />
Lysiphlebus fabarum parasitised between 1.3% and 72.9% <strong>of</strong> available<br />
hosts, Ephedrus nr cerasicola 1.3% to 1.4% and Aphidius absinthii 11%. A<br />
low level <strong>of</strong> attack on most <strong>of</strong> the parasitoids was recorded by the<br />
hyperparasitoids Alloxysta sp. and Pachyneuron sp.<br />
Ants (Monomorium indicum and Pheidole sp.) were associated with over<br />
90% <strong>of</strong> the aphid colonies. The size <strong>of</strong> A. craccivora on Vicia faba nursed by<br />
ants was far greater than those not attended and aphid mortality was higher<br />
when ants were absent. The most abundant <strong>of</strong> 22 predator species (Table<br />
4.4.2) were Cheilomenes sexmaculata, Coccinella septempunctata and<br />
Syrphus spp.<br />
In the same environment Trioxys sinensis and Lysiphlebus fabarum<br />
parasitised Aphis gossypii on cucumbers and on Hibiscus esculentus (Hamid<br />
et al. 1977).<br />
PHILIPPINES<br />
A. craccivora is commonly attacked by Lipolexis scutellaris; and A. gossypii<br />
by this species and also Trioxys communis. Both parasitoids also attack other<br />
species <strong>of</strong> aphids, but not the banana aphid Pentalonia nigronervosa (V.J.<br />
Calilung pers. comm. 1995).
68 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
RƒUNION<br />
A. craccivora is parasitised by Aphidius colemani on Gliricidia maculata<br />
and by Aphelinus sp. on Vigna unguiculata. In turn Aphelinus sp. is<br />
parasitised by Syrphophagus africanus and Pachyneuron vitodurense.<br />
A. craccivora is also attacked by the coccinellid predators Scymnus<br />
constrictus and Platynaspis capicola (Quilici et al. 1988).<br />
SHANGHAI<br />
The main parasitoids <strong>of</strong> A. gossypii on cotton were Trioxys communis,<br />
T. rietscheli and Lipolexis gracilis. Next in importance was Aphidius<br />
gifuensis, and there was occasional attack by Aphelinus abdominalis and<br />
A. mali. The highest total parasitisation recorded was about 27%. Parasitoids<br />
constituted 22.7% <strong>of</strong> the emergences from aphid mummies and the<br />
hyperparasitoids Syrphophagus aphidivora 45.2%, Alloxysta sp. 15.1%,<br />
Pachyneuron aphidis 14.7% and Dendrocerus 2.3%. The number <strong>of</strong><br />
parasitoids only exceeded that <strong>of</strong> hyperparasitoids during the first half <strong>of</strong><br />
August (Shi 1980, 1987).<br />
TONGA<br />
Aphidius colemani and Lysiphlebus testaceipes were introduced from<br />
cultures in Czechoslovakia for the biological control <strong>of</strong> the banana aphid<br />
Pentalonia nigronervosa (Stechmann and Všlkl 1988, 1990; Všlkl et al.<br />
1990), but there is no indication that they became established. A. colemani<br />
from a culture originating from a garden in Canberra was introduced again in<br />
1990 and recovered in 1992 from Aphis gossypii on taro, but not from the<br />
banana aphid. The further introduction <strong>of</strong> Aphidiidae, which are obligate<br />
parasitoids <strong>of</strong> aphids, was recommended since they could assist in the<br />
control <strong>of</strong> pest aphids and not pose a threat to non-target insects (Carver et al.<br />
1993). Although 15 aphid species are recorded in Tonga including<br />
A. craccivora, by 1993 no aphids other than A. gossypii had been recorded<br />
as hosts <strong>of</strong> A. colemani, although recent monitoring has not been possible.<br />
Two other primary parasitoids were recorded, Aphelinus gossypii (from<br />
Aphis gossypii) and Lipolexis scutellaris, from a single female, free on a<br />
banana sucker (Carver et al. 1993; Wellings et al. 1994).<br />
Three common and widespread aphid predators were recorded (the<br />
syrphid Ischiodon scutellaris, the coccinellid Harmonia octomaculata and<br />
the hemerobiid Micromus timidus) in addition to 11 tramp species <strong>of</strong> ants<br />
(Carver et al. 1993).
USA<br />
USSR<br />
4.4 Aphis gossypii 69<br />
Lysiphlebus testaceipes parasitised 74.5% <strong>of</strong> Aphis gossypii on strawberries<br />
and Aphelinus semiflavus a smaller number. Seven hyperparasitoids were<br />
reared (Oatman et al. 1983b). L. testaceipes was considered by Schlinger<br />
and Hall (1960) to be the most effective aphid parasitoid in southern<br />
California and to give excellent control there <strong>of</strong> Aphis gossypii. At least 8<br />
hyperparasitoid species were also reared (Schlinger and Hall 1960).<br />
Entomopathogen infection was the primary cause <strong>of</strong> a reduction in<br />
A. gossypii population that occurred during the week after peak aphid<br />
abundance on cotton in Mississippi and continued pathogen activity,<br />
combined with predation, maintained aphids at a low density for the<br />
remainder <strong>of</strong> the season. Early in the season parasitisation and predation<br />
may have reduced aphid population growth (Weathersbee and Hardee 1993,<br />
1994).<br />
In untreated cotton plots small predators (spiders and Geocoris spp.:<br />
Hemiptera, Lygaeidae) had the greatest impact on A. gossypii populations<br />
and the parasitoid Lysiphlebus testaceipes was never abundant. Fungi killed<br />
many aphids and constituted the most important natural enemy factor in<br />
insecticide treated plots (Kerns and Gaylor 1993). Fungi attacking<br />
A. gossypii in USA include Neozygites fresenii (Steinkraus et al. 1992,<br />
1993a,b,c; Sanchez-Pena 1993, Smith and Hardee 1993) and<br />
Cephalosporium (= Verticillium) lecanii (Sopp et al. 1990, Yokomi and<br />
Gottwald 1988). The coccinellid predators Hippodamia convergens and<br />
Scymnus louisianae, the chrysopid Chrysoperla carnea, and Syrphus sp.<br />
were effective in reducing populations <strong>of</strong> A. gossypii in Texas (Vinson and<br />
Scarborough 1989). In Alabama the hemerobiid Micromus posticus is an<br />
important predator (Miller and Cave 1987).<br />
In southern USSR coccinellid beetles are important predators <strong>of</strong> aphids.<br />
Adult Coccinella undecimpunctata and larvae <strong>of</strong> Hippodamia variegata are<br />
the most voracious and prefer A. gossypii, whereas Coccinella<br />
septempunctata prefers A. craccivora (Belikova and Kosaev 1985).<br />
There are many papers dealing with the control <strong>of</strong> A. gossypii and<br />
associated pests in glasshouses. The lacewing Chrysoperla carnea was<br />
effective only when released at a predator: aphid ratio <strong>of</strong> 1:20, whereas<br />
Chrysopa sinica was effective at 1:50 (Shuvakhina 1983). Other predators<br />
utilised include Chrysopa perla (Ushchekov 1989) and the cecidomyiid<br />
Aphidoletes aphidimyza (Begunov and Storozhkov 1986). Under conditions<br />
<strong>of</strong> high humidity, high aphid mortality was caused by the fungi<br />
Cephalosporium lecanii, Beauveria bassiana and Paecilomyces<br />
fumosoroseus (Pavlyushin and Krasavina 1987).
70 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
VIETNAM<br />
A. gossypii was one <strong>of</strong> four aphids surveyed for parasitoids by Starù and<br />
Zelenù (1983), but the number <strong>of</strong> aphidiid species found (2) was surprisingly<br />
low. Lipolexis scutellaris was commoner than Lysiphlebia mirzai. The<br />
former parasitises a number <strong>of</strong> other Aphis species, including Aphis<br />
spiraecola (= A. citricola), A. craccivora and A. nerii. Some unidentified<br />
aphelinid parasitoids were also present.<br />
Starù and Zelenù (1983) suggest that Lipolexis scutellaris may be a<br />
valuable species for transfer elsewhere and also that Vietnam would benefit<br />
from the introduction <strong>of</strong> additional parasitoid species.<br />
The major parasitoid species<br />
General features <strong>of</strong> the Aphidiidae<br />
Different populations <strong>of</strong> many Aphidiidae have differing biological<br />
properties and are <strong>of</strong>ten known as biotypes, i.e. contrasting groups, each<br />
consisting <strong>of</strong> individuals <strong>of</strong> the same species. Biotypes are recognised by<br />
biological function rather than morphology and consist <strong>of</strong> those individuals<br />
that behave similarly as far as our immediate interests are concerned<br />
(Mackauer and Way 1976).<br />
The members <strong>of</strong> this family attack aphids exclusively and are probably<br />
the most commonly observed cause <strong>of</strong> aphid mortality in the field. In<br />
Europe, many, if not all, aphid colonies come to include some mummified<br />
individuals (i.e. dead aphids containing a fully-grown parasitoid larva or<br />
pupa) (Starù 1970). Aphidiidae may hibernate as prepupae within host<br />
mummies. All are solitary endoparasitoids. All aphid stages are attacked<br />
except the eggs, but alatae are least <strong>of</strong>ten attacked. The parasitoid egg is<br />
usually inserted anywhere in the aphid abdomen. The preferred aphid larval<br />
instar varies with the parasitoid species, but younger instars are usually<br />
chosen. If adult aphids are parasitised the parasitoid larva may not complete<br />
its development before the insect dies, so that the parasitoid perishes also.<br />
Oviposition into an aphid does not ensure the successful development <strong>of</strong> a<br />
parasitoid, since the host may be unsuitable or it may already be parasitised:<br />
defence and immune responses are common. However parasitised hosts are<br />
usually distinguished by the parasitoid and receive no further eggs.<br />
If an aphid is parasitised during the last larval instar, a mummy is formed<br />
after it moults to the adult. The first and second instars <strong>of</strong> aphidiid<br />
parasitoids generally feed on haemolymph, but the last instar attacks the<br />
alimentary canal and other organs, ultimately killing its host. The parasitoid<br />
larva then spins a cocoon and pupates inside the empty aphid skin. The adult<br />
emerges through a small circular hole usually cut dorsally or apically near
4.4 Aphis gossypii 71<br />
the posterior <strong>of</strong> the mummy. Aphidiid mummies are round and usually<br />
straw-coloured to brown and in some genera (e.g. Ephedrus) always black<br />
and parchment-like. Aphelinid larvae do not spin cocoons, and their<br />
mummies are usually slender and black (Takada 1992).<br />
The chain <strong>of</strong> events that determines host specificity includes, in<br />
sequence, host habitat finding, host finding, host acceptance by the<br />
parasitoid and host suitability.<br />
The last larval instar <strong>of</strong> some parasitoids provokes their aphid hosts to<br />
move away from the plant on which they were feeding. With Lysiphlebus<br />
fabarum, Ephedrus plagiator and Trioxys angelicae this migration <strong>of</strong> premummies<br />
is connected with diapause (under conditions <strong>of</strong> a short day) or<br />
with aestivation (under conditions <strong>of</strong> a long day). Parasitoids usually emerge<br />
without delay from aphids which become mummies where they have been<br />
feeding (Behrendt 1968).<br />
Starù (1970) provides additional details <strong>of</strong> many <strong>of</strong> the species <strong>of</strong><br />
Aphidiidae that follow below and HŒgvar and H<strong>of</strong>svang (1991) a<br />
comprehensive review <strong>of</strong> their biology, host selection and use in biological<br />
control.<br />
Aphelinus abdominalis (= A. flavipes) Hym.: Aphelinidae<br />
This species is widespread in Europe and is recorded also from USSR, India,<br />
Australia and Israel. It is extensively distributed in India as one <strong>of</strong> the<br />
important parasitoids <strong>of</strong> Myzus persicae and A. gossypii. It becomes active<br />
in late May and is abundant during June and July. The incubation time for<br />
eggs is 2 days and adults emerge after 13 days in September (Ramaseshiah<br />
and Dharmadhikari 1969).<br />
When an Indian strain was liberated in a U.K. glasshouse at 23¡C, one<br />
week after artificially infesting plants with A. gossypii, it was unable to<br />
overtake the pest population because the rate <strong>of</strong> increase <strong>of</strong> the pest was<br />
scarcely affected by the parasitoid. Only when aphid overcrowding occurred<br />
and rate <strong>of</strong> increase was self-limited, did the parasiteÕs rate <strong>of</strong> increase (6 ´ per<br />
week) exceed that <strong>of</strong> the aphid. Reducing the glasshouse temperature to 19¡C<br />
slowed the rate <strong>of</strong> aphid increase and permitted the parasitoids to contain the<br />
pest before severe leaf-distortion occurred. On the other hand, when<br />
parasitoids were present at the time that A. gossypii was introduced, aphid<br />
reproduction was suppressed and effective control resulted (Hussey and<br />
Bravenboer 1971).<br />
Aphelinus gossypii Hym.: Aphelinidae<br />
This species was described from Hawaii and is recorded from Australia,<br />
New Zealand, India, Japan and also from Tonga, where it was reared in<br />
abundance from Aphis gossypii on taro (Colocasia esculenta). It is probably
72 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
present elsewhere under other names (Carver et al. 1993). Aphelinus<br />
gossypii lays in some host eggs encountered, but also kills many others by<br />
probing and then feeding on exuding fluids (Takada and Tokumaku 1996).<br />
Parasitisation is reduced when the host is protected by the presence <strong>of</strong> ants<br />
(Stechman et al. 1996). Aphelinus gossypii was parasitised in Tonga to the<br />
extent <strong>of</strong> 30% to 60% by the cynipoid hyperparasitoid Alloxysta darci. This<br />
parasitises species <strong>of</strong> Aphelinus, but not <strong>of</strong> Aphidiidae so it is most unlikely<br />
to parasitise Aphidius colemani, a recently established parasitoid attacking<br />
Aphis gossypii there. Alloxysta darci was earlier incorrectly identified as<br />
Alloxysta brevis (Carver 1992; Carver et al. 1993). Aphelinus gossypii is an<br />
effective parasitoid at low A. gossypii density and hence an important<br />
candidate for consideration for introduction (P. Wellings, pers. comm.).<br />
Aphelinus mali Hym.: Aphelinidae<br />
This well known North American parasitoid <strong>of</strong> above-ground stages <strong>of</strong> the<br />
woolly apple aphid Eriosoma lanigerum has been introduced intentionally<br />
or inadvertently into almost every country where its host has established<br />
itself as a pest. It is a very effective parasitoid in moderately warm climates<br />
(Rosen 1967b) and has occasionally been reported from other hosts,<br />
including Aphis gossypii, although in such instances it has probably been<br />
confused with the very similar Aphelinus gossypii (M. Carver pers. comm.).<br />
The life cycle occupies almost 20 days in summer (egg 3, larva 10 to 12 and<br />
pupa 6 to 7 days respectively). Parasitised aphids have a strong tendency to<br />
seek sheltered places before death (Clausen 1978).<br />
Aphelinus semiflavus Hym.: Aphelinidae<br />
This species has a wide host range and occurs in USA, Hawaii, India and<br />
Europe. It overwinters as a pupa in its host. Single eggs are laid, generally in<br />
the dorsal surface <strong>of</strong> the host abdomen. Young aphids are preferred hosts, but<br />
even adults are parasitised, in which case fewer young are produced by the<br />
aphid before it is killed. Over 600 eggs may be produced by a female which<br />
<strong>of</strong>ten feeds on haemolymph exuding from oviposition punctures. Males are<br />
rare and females can produce <strong>of</strong>fspring parthenogenetically. Developmental<br />
periods are: egg 3 days, larva 6 to 11 days and pupa 7 to 8 days. Although<br />
Myzus persicae is preferred, A. semiflavus also parasitises Aphis gossypii as<br />
one <strong>of</strong> 15 or so other hosts. In USA it was attacked by 3 hyperparasitoids,<br />
Asaphes lucens (= A. americana) (Pteromalidae), Alloxysta sp. (Charipidae)<br />
and Syrphophagus aphidivora (= Aphidencyrtus aphidiphagus) (Encyrtidae)<br />
(Hartley 1922; Ramaseshiah and Dharmadhikari 1969; Schlinger and Hall<br />
1959).
4.4 Aphis gossypii 73<br />
Aphidius colemani Hym.: Aphidiidae (= A. platensis,<br />
= A. transcaspicus)<br />
Starù (1975) postulated that this species originated in India or nearby<br />
(possibly the Eastern Mediterranean). It is now widely distributed in<br />
Mediterranean Europe, Asia Minor, Central Asia, India, Pakistan, Africa,<br />
South America, Australia, New Zealand and New Caledonia (Starù 1972). In<br />
addition there have been intentional introductions to California, U.K.,<br />
Czechoslovakia, Kenya (Starù1975) and Tonga (Carver et al. 1993). It is<br />
rather strange, if it originated in India, that it appears to be absent from<br />
Japan, China and possibly some <strong>of</strong> <strong>Southeast</strong> Asia (Starù 1975; Takada<br />
1992). A. colemani is restricted to the family Aphididae. Hosts consist <strong>of</strong> at<br />
least 9 species <strong>of</strong> Aphis, including A. craccivora and A. gossypii and at least<br />
30 species in other genera (Elliott et al. 1994; Starù 1975).<br />
In the field in Australia A. colemani is known to parasitise many species<br />
in the aphid tribes Aphidini and Myzini, but rarely species in the<br />
Macrosiphini and even more rarely species in other subfamilies (Carver et<br />
al. 1993).<br />
There are significant differences between countries both in the range <strong>of</strong><br />
hosts attacked by A. colemani and the preference for particular host species<br />
(e.g. Messing and Rabasse 1995). This indicates that the species that is<br />
known as A. colemani is a complex <strong>of</strong> closely-related species or biotypes.<br />
For example, A. colemani parasitises Melanaphis donacis and Hyalopterus<br />
pruni in Mediterranean Italy and France, but none <strong>of</strong> the many other aphids<br />
present; in Central Asia only the latter aphid is attacked and in Iraq both<br />
aphids are attacked, in addition to Aphis zizyphi and A. punicae. An Italian<br />
population from Hyalopterus pruni was successfully reared on both Aphis<br />
craccivora and A. fabae in the laboratory. Furthermore, a French population<br />
from Melanaphis donacis was readily reared in the laboratory on Aphis<br />
craccivora, A. fabae and Myzus persicae (Starù1975). As another example, a<br />
strain (from Brazil) <strong>of</strong> Aphidius colemani successfully parasitised the<br />
oleander aphid Aphis nerii in France whereas another strain (from France)<br />
failed to do so (Tardieux and Rabasse 1986, 1988). In Mediterranean regions<br />
A. colemani parasitised A. gossypii successfully at 20¡C, but at temperatures<br />
above 27¡ it frequently failed to do so (Guenaoui 1991). The number <strong>of</strong> eggs<br />
laid per female A. colemani was 302 at 20¡C and 388 at 25¡C and<br />
development time to adult was 12.7 days and 10.0 days respectively. The<br />
intrinsic rate <strong>of</strong> increase <strong>of</strong> the parasitoid was similar to that <strong>of</strong> A. gossypii,<br />
suggesting that it is a promising parasitoid (van Steenis 1993). The optimum<br />
scheme for introducing A. colemani into glasshouses for control <strong>of</strong><br />
A. gossypii on cucumbers has been investigated by van Steenis et al. (1996).<br />
Chou (1984) recorded A. colemani from TaiwanÑthe first record <strong>of</strong> this<br />
species from east AsiaÑwith H. pruni as its only host.
74 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
The care needed in selecting a biotype appropriate to the target pest is<br />
also illustrated by the following example. A. colemani from Aphis nerii<br />
mummies on a garden plant (Tweedia coerulia) in Canberra was readily<br />
reared for some generations in an insectary on the banana aphid Pentalonia<br />
nigronervosa before release in Tonga for biological control <strong>of</strong> that species. It<br />
has not been recovered from the banana aphid, but is now well established on<br />
A. gossypii attacking cucurbits (Carver et al. 1993; Wellings et al. 1994).<br />
When P. nigronervosa colonies are small they are mainly located deep in the<br />
leaf sheaths and they only extend into more exposed areas as they increase in<br />
size. Stadler and Všlkl (1991) found that Lysiphlebus testaceipes searched<br />
mainly in exposed areas for hosts, whereas A. colemani searched both<br />
exposed and concealed areas. This suggests that A. colemani would be the<br />
more appropriate <strong>of</strong> the parasitoids for hosts in concealed situations and,<br />
interestingly, it has been reported from P. nigronervosa in the field in<br />
northern New South Wales (M. Carver pers. comm.), where it is rare and was<br />
not encountered in recent searches (P.W. Wellings pers. comm.).<br />
A. colemani (<strong>of</strong>ten under one <strong>of</strong> its synonyms) has been introduced to<br />
several countries for the biological control <strong>of</strong> a range <strong>of</strong> aphid species (Starù<br />
1975).<br />
Aphidius gifuensis Hym.: Aphidiidae<br />
This species is native to the Oriental region. Details <strong>of</strong> its fecundity,<br />
oviposition period and longevity are provided by Fukui and Takada (1988).<br />
Aphidius matricariae Hym.: Aphidiidae (= A. phorodontis)<br />
This species is native to the temperate zones <strong>of</strong> the Palearctic region and has<br />
been recorded from more than 40 aphid species in Europe, North Africa, the<br />
Middle East, Israel, Mongolia and North and South America. It has a<br />
preference for the green peach aphid Myzus persicae in Israel (Rosen<br />
1967a,b) and California (Schlinger and Mackauer 1963). After contact with<br />
honeydew or an aphid host the time spent in searching that region for hosts<br />
increased (Masum 1994).<br />
Ephedrus persicae Hym.: Aphidiidae<br />
This is an almost cosmopolitan species, which is probably native to the<br />
Middle East or Central Asia, and now occurs in the Far East, Europe, South<br />
Africa, Madagascar, Australia and North America. It prefers leaf-curling<br />
aphid hosts, mainly belonging to the Myzinae and, less frequently, to the<br />
Aphidinae (Aphis spp.) (Mackauer 1963, 1965; Starù 1966). A review <strong>of</strong> the<br />
taxonomy and biology <strong>of</strong> Ephedrus persicae and E. plagiator is provided by<br />
GŠrdenfors (1986).
4.4 Aphis gossypii 75<br />
Ephedrus plagiator Hym.: Aphidiidae<br />
This aphid is native to the far eastern deciduous forests and steppes <strong>of</strong> the<br />
Palearctic region and is widely distributed in India. It has many hosts<br />
amongst species <strong>of</strong> Aphis and Myzus (Starù 1967a).<br />
Lipolexis gracilis Hym.: Aphidiidae<br />
This is a European or Far Eastern species with hosts in a number <strong>of</strong> aphid<br />
genera, including Aphis (Starù 1967a).<br />
Lipolexis scutellaris Hym.: Aphidiidae<br />
This is an oriental species (Raychaudhuri 1990) and is known from southern<br />
China, Japan and Taiwan and also from India, Pakistan and Tonga. It has a<br />
wide host range and an apparent preference for Aphis species (Carver et al.<br />
1993).<br />
Lysiphlebia japonica Hym.: Aphidiidae<br />
This is native to the Far East and is probably a well-adapted species for<br />
tropical climates. It typically occurs in forest or open woodland<br />
environments on many species in the genus Aphis in addition to those in a<br />
number <strong>of</strong> other related aphid genera (Starù 1967b).<br />
Lysiphlebia mirzai Hym.: Aphidiidae<br />
This species was described from India and is known also from Vietnam and<br />
China.<br />
Lysiphlebus fabarum Hym.: Aphidiidae<br />
(= Lysiphlebus confusus, L. ambiguus)<br />
This is a Palaearctic species (Europe, Asia Minor, Caucasus, Central Asia),<br />
which is now widespread and occurs also in Israel, a number <strong>of</strong> African<br />
countries and USA. It is the most abundant parasitoid <strong>of</strong> the black citrus<br />
aphid Toxoptera aurantii in Italy (Starù 1964) and Israel (Rosen 1967a,b). In<br />
some countries it is biparental but, in Israel, only females are known (Rosen<br />
1967a,b). It is recommended by Starù (1967b) as a species useful for<br />
biological control. L. fabarum is both biparental and parthenogenetic<br />
(Carver 1984) and 15 to 16 generations a year have been recorded in Italy<br />
(Tremblay 1964). It has a very extensive host range, with records from at<br />
least 144 species <strong>of</strong> aphids in 36 genera, 81 (56%) <strong>of</strong> these species belonging<br />
to the genus Aphis (Carver 1984).<br />
There is little doubt that L. fabarum refers to a complex <strong>of</strong> closely related<br />
sibling species or at least <strong>of</strong> host-specific biotypes. For example, in the<br />
laboratory L. fabarum bred from Aphis species readily parasitised other<br />
Aphis species, but not Brachycaudus sp.. However, in the field, colonies <strong>of</strong><br />
Brachycaudus cardui heavily parasitised by L. fabarum shared the same<br />
host plants as unparasitised Aphis fabae (Mackauer 1962a). The influence <strong>of</strong>
76 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
temperature and humidity on the development <strong>of</strong> L. fabarum in A. gossypii<br />
and A. craccivora has been reported by Davletshina and Gomolitskia (1975)<br />
and methods for its mass production by Tregubenko and Popushoi (1987).<br />
Lysiphlebus testaceipes Hym.: Aphidiidae<br />
This parasitoid has a natural range extending from North America through<br />
Central America to the northern parts <strong>of</strong> South America. It is now known<br />
also from Hawaii, Australia, Europe and East Africa. It is the commonest<br />
native parasitoid <strong>of</strong> aphids in Mexico (Starù and Remaudi re 1982). It has a<br />
very wide host range, having been reported from at least 79 aphid species (32<br />
in the genus Aphis) in 32 genera (Carver 1984).<br />
Oviposition generally occurs in the abdomen <strong>of</strong> half grown and<br />
unparasitised hosts and, when hosts are scarce, more than one egg may be<br />
deposited (Sekhar 1957). Up to 254 eggs may be laid. It is heavily attacked<br />
by hyperparasitoids in its natural range (eg. 6 species when attacking Aphis<br />
gossypii (Schlinger and Hall 1960) or 7 species (Oatman et al. 1983b)).<br />
L. testaceipes was present in Australia (New South Wales and South<br />
Australia) prior to its introduction as a biological control agent and attacked<br />
an indigenous aphid Aphis acaenovinae (Starù and Carver 1979).<br />
There are many examples to demonstrate that L. testaceipes consists <strong>of</strong><br />
biotypes. Thus, Californian L. testaceipes is unable to complete its<br />
development on Aphis spiraecola, whereas a Cuban strain does so<br />
successfully. The Californian strain did not attack Aphis nerii after<br />
introduction to Hawaii, although a Mexican strain subsequently introduced<br />
did so (Starù 1970). Then again, the biotype from A. craccivora on Robinia<br />
pseudacacia does not parasitise this same aphid on Phaseolus vulgaris;<br />
another biotype prefers A. gossypii on squash to this same aphid on hibiscus<br />
(Sekhar 1960; Tremblay and Barbagallo 1982). The effectiveness <strong>of</strong><br />
Lysiphlebus testaceipes as a parasitoid on A. gossypii is thus significantly<br />
affected by the host plant on which the aphid is feeding (Steinberg et al.<br />
1993).<br />
L. testaceipes is reported to attack A. gossypii in Cuba (Starù 1981),<br />
Mexico (Starù and Remaudi re 1982) and, after introduction to Europe, in<br />
Spain, France and Italy. In Europe it now attacks more than 26 aphid species<br />
including A. craccivora and A. gossypii, <strong>of</strong>ten with high levels <strong>of</strong><br />
parasitisation (Starù et al. 1988a,b,c).<br />
Lysiphlebus testaceipes was successfully introduced in 1923 from<br />
California to Hawaii for the biological control <strong>of</strong> aphids, including Aphis<br />
craccivora and A. gossypii. It soon spread widely throughout the islands,<br />
attacking these and other aphid species. In 1965, a further introduction from<br />
Mexico was made to control the oleander aphid Aphis nerii which had<br />
previously escaped attack. By 1927 several hyperparasitoids were recorded
4.4 Aphis gossypii 77<br />
as attacking L. testaceipes breeding in Rhopalosiphum maidis, an important<br />
virus vector on sugarcane (Timberlake 1927). According to Starù (1970), the<br />
introductions were partially to substantially effective, but importation <strong>of</strong><br />
additional species was recommended. L. testaceipes from Cuba was<br />
introduced to Czechoslovakia for the biological control <strong>of</strong> aphids in<br />
greenhouses and also <strong>of</strong> some pest aphids in some subtropical areas (Starù<br />
1970). L. testaceipes was introduced in 1956 and 1960 from Hawaii to the<br />
Philippines, but no recoveries have been recorded (Baltazar 1963).<br />
L. testaceipes was introduced in 1973 from Cuba into France and Corsica<br />
(Italy) to reduce the numbers <strong>of</strong> citrus aphids (Starù et al. 1988b). It became<br />
well established, heavily parasitising Aphis gossypii and several other aphids<br />
(Rabasse 1986).<br />
Although there do not seem to be comparable data for A. gossypii, Hall<br />
and Ehler (1980) found that L. testaceipes averaged 79.5% parasitisation <strong>of</strong><br />
Aphis nerii populations on oleander, with an average density <strong>of</strong> 12.4 aphids<br />
per shoot. When natural enemies were excluded, an average <strong>of</strong> 32.6 aphids<br />
were present per shoot, a clear indication that parasitisation was having a<br />
significant effect. A hyperparasitoid Pachyneuron sp. was active, but<br />
appeared to be generally unimportant in aphid population regulation. The<br />
average fecundity <strong>of</strong> L. testaceipes was found to be 128.2 eggs at 20¡C and<br />
180 eggs at 25¡C. Development from egg to female adult was completed in<br />
12.9 days at 20¡C and 9.5 days at 25¡C; and the life span <strong>of</strong> females was 2.7<br />
and 2.6 days at 20 and 25¡C respectively. At 20¡C the intrinsic rate <strong>of</strong><br />
increase was slightly lower than that <strong>of</strong> Aphis gossypii, but was the same at<br />
25¡C. It was concluded that, at temperatures below 25¡C, L. testaceipes<br />
might not be able to overtake an established population <strong>of</strong> A. gossypii (van<br />
Steenis 1994). Earlier Schlinger and Hall (1960) concluded that<br />
L. testaceipes was capable <strong>of</strong> producing excellent control <strong>of</strong> both<br />
A. craccivora and A. gossypii.<br />
Praon volucre Hym.: Aphidiidae<br />
This is a palearctic species and is known from the Middle East, North Africa,<br />
India and Central Asia. It has an extensive and diverse host range, having<br />
been recorded from at least 90 aphid species in 35 genera. There is good<br />
evidence that P. volucre exists as a complex <strong>of</strong> host-specific biotypes or<br />
sibling species (Mackauer 1959, 1962a,b). Its biology has been studied by<br />
Beirne (1942). As in other Praon spp., pupation takes place under the empty<br />
mummy <strong>of</strong> the parasitised host. P. volucre females have been observed to<br />
use their front legs to hold the host aphid during oviposition (Beirne 1942).<br />
Although it has been recorded from A. craccivora in the field (Starù 1967a)<br />
and the laboratory (Carver 1984), it does not seem to have been reported<br />
from A. gossypii.
78 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Trioxys angelicae Hym.: Aphidiidae<br />
This is widely distributed in Europe, Asia Minor and North Africa and has<br />
been reared from a wide range <strong>of</strong> hosts (Rosen 1967a,b).<br />
Trioxys communis Hym.: Aphidiidae<br />
This very important parasitoid <strong>of</strong> A. gossypii in China has been studied in a<br />
series <strong>of</strong> papers by Shi (1984, 1985, 1986). It develops in 8 days at 30¡C and<br />
in 16 days at 20¡C. When 4th instar A. gossypii are parasitised they<br />
mummify in the adult stage, having produced very few <strong>of</strong>fspring. It is<br />
hyperparasitised by the pteromalid Pachyneuron aphidis.<br />
T. communis has also been recorded from A. gossypii in Japan, Korea,<br />
Taiwan and India and has been taken rarely in Malaysia from Aphis<br />
spiraecola (= A. citricola), but apparently not from other aphids (Ng and<br />
Starù 1986).<br />
Trioxys indicus Hym.: Aphidiidae<br />
A valuable review <strong>of</strong> the biology, ecology and control efficiency <strong>of</strong> this<br />
parasitoid is presented by Singh and Agarwala (1992). Its biology has been<br />
studied by Subba Rao and Sharma (1962) and later in a long series <strong>of</strong> papers<br />
from India (e.g. Singh et al. 1979; Sinha and Singh 1980a,b; Singh and Sinha<br />
1980a,b,c, 1982a,b; Pandey et al. 1982, 1984; Kumar et al. 1983; Singh and<br />
Pandey 1986; Singh and Srivastava 1988a,b, 1991, Singh and Agarwala<br />
1992). Ghosh and Agarwala (1982) provide host, host plant records and<br />
information on its distribution in India.<br />
Recorded hosts <strong>of</strong> T. indicus belong to 24 species <strong>of</strong> aphids in 14 genera,<br />
<strong>of</strong> which species <strong>of</strong> Aphis are best represented. The majority <strong>of</strong> host aphids<br />
are polyphagous, 6 are oligophagous and 1 monophagous. T. indicus prefers<br />
hosts on cultivated and wild shrubs to those on herbaceous and woody<br />
plants. A. craccivora is parasitised to the extent <strong>of</strong> 87% on pigeon pea (Singh<br />
and Tripathi 1987) and A. gossypii to the extent <strong>of</strong> 60% on both bottle gourd<br />
(Singh and Bhatt 1988) and eggplant (Subba Rao and Sharma 1962), and<br />
30% on cotton (Agarwala 1988).<br />
Based on extensive field observations, the three main hosts <strong>of</strong> T. indicus<br />
are A. craccivora, A. gossypii and A. nerii, each <strong>of</strong> which is parasitised by a<br />
range <strong>of</strong> other polyphagous parasitoids.<br />
The native range <strong>of</strong> T. indicus is largely the Indian subcontinent,<br />
although it has also been recorded from Taiwan (Starù and Schlinger 1967).<br />
It is most abundant in tropical and subtropical regions, where its numbers are<br />
comparatively low in summer and in rainy months and higher in the cooler<br />
months (Agarwala 1988).<br />
T. indicus females prefer to oviposit in second and third instar host<br />
nymphs. Probing without oviposition is common with first and second
Diptera<br />
4.4 Aphis gossypii 79<br />
instars, leading to high aphid mortality (up to 80% for first instars). Hosts<br />
that are already parasitised are generally avoided. Fecundity varies, but the<br />
figure <strong>of</strong> 143 <strong>of</strong>fspring per female is quoted. The average time from<br />
oviposition to emergence is 10 days at 24 to 27¡C on A. gossypii (Subba Rao<br />
and Sharma 1962) and 16 to 18 days at 24 to 26¡C on A. craccivora.<br />
T. indicus lives less than 10 days in the laboratory (Pandey et al. 1982).<br />
Augmentation <strong>of</strong> T. indicus early in the season in pigeon pea fields in India<br />
was sufficient to control A. craccivora (Singh and Agarwala 1992). Extracts<br />
<strong>of</strong> A. craccivora sprayed on pigeon pea increased the fecundity <strong>of</strong> T. indicus<br />
and reduced the population doubling time (Singh and Srivastava 1991).<br />
A density-dependent relationship between T. indicus and its hosts has<br />
been reported by several authors (Subba Rao and Sharma 1962; Singh and<br />
Sinha 1980a; Saha and Agarwala 1986; Bhatt and Singh 1991a,b ).<br />
Eleven hyperparasitoids <strong>of</strong> T. indicus are known (Singh and Agarwala<br />
1992) and these should be rigorously excluded in any biological control<br />
transfers.<br />
Singh and Agarwala (1992) conclude that T. indicus is a very important<br />
parasitoid, especially <strong>of</strong> several species <strong>of</strong> Aphis in India, that it possesses<br />
most <strong>of</strong> the desirable attributes <strong>of</strong> a successful biological control agent and is<br />
therefore a promising natural enemy for introduction elsewhere against<br />
relevant pest aphids. It is also <strong>of</strong> value for inundative releases (Singh and<br />
Rao 1995).<br />
Endaphis maculans Dip.: Cecidomyiidae<br />
This aphid endoparasitoid attacked Toxoptera aurantii freely, but<br />
A. gossypii only lightly. It was seldom found in A. craccivora (Kirkpatrick<br />
1954). E. maculans lays its eggs on aphid-infested leaves and, upon<br />
hatching, the larva searches for aphids. When a host is encountered the larva<br />
penetrates the aphids dorsum and develops as an endoparasitoid, leaving as a<br />
mature larva via the aphidÕs anus. Average development time from egg to<br />
adult at 25¡ to 26¡C was 19.1 days (Tang et al. 1994).<br />
An aphid-specific predator<br />
Aphidoletes aphidimyza Dip.: Cecidomyiidae<br />
This is a common and widely distributed species throughout the northern<br />
hemisphere. It has been recorded from Japan, USSR, Czechoslovakia,<br />
Austria, Germany, Finland, France, Netherlands, U.K., Italy, Israel, Egypt,<br />
Sudan, Canada, USA and Hawaii. It is not recorded from Australia or New<br />
Zealand.
80 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Comments<br />
Larvae <strong>of</strong> Aphidoletes reportedly feed exclusively as predators on aphids<br />
and are hence more host-specific than many <strong>of</strong> the other predators:<br />
A. aphidimyza is the best known <strong>of</strong> the cecidomyiid predators. Adults<br />
emerge during the day from pupae in the soil. They generally fly between<br />
sunset and sunrise. Orange-coloured eggs are laid singly or in clusters <strong>of</strong> up<br />
to 40, usually on plants near aphid colonies. Females live up to 14 days in the<br />
laboratory and lay about 100 eggs. These hatch after 3Ð4 days and first instar<br />
larvae immediately seek out and attack aphids. They usually attack by<br />
piercing a leg joint or some other joint. A toxin is perhaps injected, since the<br />
aphid is rapidly immobilised before its body fluids are extracted. The<br />
shrivelled bodies <strong>of</strong> some aphids remain attached to the plant by the stylet.<br />
Larval development involves 3 instars and takes 7 to 14 days (Harris 1973).<br />
On the other hand, Herpai (1991) reports 21 days from egg to adult (egg 3<br />
days, larva 8 days, pupa 10 days).<br />
Roberti (1946) gave a figure <strong>of</strong> 60 to 80 Aphis gossypii attacked per day.<br />
Predator larvae usually drop to the soil to pupate. They construct small silk<br />
cocoons in the top few millimetres <strong>of</strong> soil, but occasionally cocoons may be<br />
spun on plants. Larvae pupate within a few days <strong>of</strong> cocoon construction and<br />
adults emerge after 1Ð3 weeks depending upon temperatures. The life cycle<br />
can be completed in about 3 weeks at temperatures above 21¡C (Harris 1973;<br />
Herpai 1991).<br />
Harris (1973) does not record it from A. craccivora. Two<br />
hyperparasitoids <strong>of</strong> A. aphidimyza are known in Africa, the platygasterid<br />
Synopeas rhanis and an unidentified braconid (Harris 1973).<br />
There are many reports in the literature that natural enemies play an<br />
important role in reducing (and probably regulating) the abundance <strong>of</strong> pest<br />
aphids. More than 100 biological control programs have been mounted<br />
against at least 26 aphid species and 48% <strong>of</strong> them have reported success<br />
(HŒgvar and H<strong>of</strong>svang 1991). Twenty three species <strong>of</strong> aphidiid parasitoids<br />
have been used in classical biological control <strong>of</strong> aphids and the parasitoids<br />
became established in 32 out <strong>of</strong> 55 attempts (Greathead 1989). Most pest<br />
aphids are attacked in their native range by many parasitoids and predators<br />
and by a few pathogenic fungi. However, many <strong>of</strong> the natural enemies have<br />
not accompanied their aphid hosts when these have spread into new regions.<br />
Indeed, they may not even be present throughout the presumed native range<br />
<strong>of</strong> their host. Since both the direct and indirect damage caused by aphids<br />
seem to be proportional to their numbers, any reduction is potentially<br />
beneficial. Even in the case <strong>of</strong> virus transmission, where the feeding (or
4.4 Aphis gossypii 81<br />
probing) <strong>of</strong> single infected aphids on a crop may lead to substantial loss,<br />
reduction in aphid numbers will more than proportionally reduce the<br />
probability <strong>of</strong> flying aphids migrating to an uninfected plant (Wellings<br />
1991), because reduced crowding <strong>of</strong> aphids usually results in a lower<br />
number forming wings. As indicated above, there have been a number <strong>of</strong><br />
attempts at classical biological control <strong>of</strong> aphids, and there are at least 7<br />
well-documented successes up to 1988 (Hughes 1989). None <strong>of</strong> these,<br />
however, involved A. craccivora or A. gossypii as the main target. These<br />
latter species have, however, been subjected to important attack by<br />
parasitoids introduced primarily against another pest aphid in the same<br />
general environment. For example, although Aphidius colemani failed to<br />
establish on Pentalonia nigronervosa in Tonga, it did so very successfully<br />
there on A. gossypii. Although A. craccivora is also present in Tonga,<br />
Aphidius colemani has not yet been recorded from it (Carver et al. 1993), but<br />
monitoring has been minimal.<br />
The best predictor <strong>of</strong> success in biological control is previous success<br />
with a natural enemy in a similar environment. If this experience is<br />
unavailable, the best chances appear to be with a climatically adapted,<br />
adequately host-specific, natural enemy that is known to attack the pest in its<br />
native or expanded range. Parasitic wasps appear to be the best natural<br />
enemies available for aphids, because they are generally far more host<br />
specific than predators and are <strong>of</strong>ten more efficient at searching for hosts at<br />
low aphid densities (Hughes 1989). Predators can be very effective in<br />
reducing aphid numbers at certain times <strong>of</strong> the year, but are <strong>of</strong>ten unable to<br />
prevent damage. Furthermore, their general lack <strong>of</strong> host specificity makes<br />
most <strong>of</strong> them unattractive to authorities responsible for approving<br />
introductions, so they have not been dealt with in any detail in this dossier.<br />
A feature that makes it difficult to generaliseÑand even to make clear<br />
recommendationsÑis that many <strong>of</strong> the identifications <strong>of</strong> some <strong>of</strong> the<br />
parasitoids are incorrect, particularly the earlier ones, but even some <strong>of</strong> the<br />
more recent ones made by non-specialists. The selection <strong>of</strong> appropriate<br />
natural enemies for an aphid biological control program requires a detailed<br />
knowledge <strong>of</strong> the ecology <strong>of</strong> the target aphid and <strong>of</strong>ten <strong>of</strong> other potential<br />
aphid hosts in the environment where it is causing problems; also <strong>of</strong> where to<br />
obtain parasitoid biotypes with appropriate host specificity and habitat<br />
adaptation. If an apparently good species fails to establish or, if established,<br />
to become effective, it is probably worth seeking the same enemy from a<br />
potentially more appropriate source, such as one with a better climate match;<br />
or a biotype that exhibits a special preference for the target aphid; or the first<br />
generation from field-collected material, rather than employing individuals<br />
bred for many generations in the laboratory (Hughes 1989).
82 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Both A. craccivora and A. gossypii are now almost cosmopolitan in their<br />
occurrence throughout the temperate, subtropical and tropical regions <strong>of</strong> the<br />
world and both are polyphagous. At least A. gossypii exists as a series <strong>of</strong><br />
biotypes with different spectra <strong>of</strong> host preferences and both it and<br />
A. craccivora owe a considerable amount <strong>of</strong> their economic importance to<br />
their ability to transmit an extensive range <strong>of</strong> important plant viruses.<br />
Both aphids are attacked by a wide range <strong>of</strong> parasitoids and share a<br />
number <strong>of</strong> these species. The majority <strong>of</strong> these parasitoids are polyphagous<br />
and attack many other (but not all) species <strong>of</strong> the genus Aphis and some<br />
species in related aphid genera. Two important parasitoids are the American<br />
Lysiphlebus testaceipes and the Indian Aphidius colemani. These parasitoids<br />
both exist in a number <strong>of</strong> biotypes with different host spectra and abilities to<br />
attack A. craccivora and A. gossypii on some plants, but not on others. Thus,<br />
when biological control is attempted, care must be taken to select a<br />
parasitoid biotype that is appropriate for the aphid biotype, the host plant and<br />
the prevailing environmental conditions. Laboratory comparison <strong>of</strong> the<br />
impact on A. gossypii <strong>of</strong> Aphidius colemani, Lysiphlebus testaceipes and<br />
Aphidius matricariae indicated that A. colemani was the most and<br />
A. matricariae the least effective (van Steenis 1992). The polyphagous<br />
nature <strong>of</strong> many effective parasitoids has the advantage that, in any region, a<br />
number <strong>of</strong> other aphid species will be parasitisedÑand hence serve as<br />
valuable reservoirs <strong>of</strong> parasitoids when the target pest population falls to a<br />
low level. The aims <strong>of</strong> aphid biological control are (i) as far as possible to<br />
maintain densities below those at which alates are formed due to crowding<br />
and (ii) if possible, to depress densities still further, so that sap removal,<br />
volume <strong>of</strong> honeydew produced and plant deformation become unimportant.<br />
Takada (1992) points out that the aphid parasitoid fauna in Far East Asia<br />
is quite different from that in India. Thus, the most important parasitoids <strong>of</strong><br />
A. gossypii in the Far East are Trioxys communis, Lysiphlebia japonica and<br />
Aphelinus sp., none <strong>of</strong> which occurs in India. On the other hand, the<br />
dominant parasitoid <strong>of</strong> A. gossypii in India is Trioxys indicus, which is<br />
present in Taiwan, but not in Japan or Korea. Another Oriental species,<br />
Lipolexis scutellaris occurs in Hong Kong, Vietnam and Malaysia. It<br />
appears that Indian parasitoids, rather than Far East <strong>Asian</strong> species are<br />
present in <strong>Southeast</strong> Asia. However inadequate information is available in<br />
<strong>Southeast</strong> Asia on the natural enemies <strong>of</strong> A. craccivora and A. gossypii<br />
present in the many different crop systems and environments in which these<br />
aphids occur. Even within a single country, it is necessary to examine the<br />
aphid population in the particular situations and the crops where they are<br />
causing important problems. This can be illustrated by the parasitoid<br />
complex <strong>of</strong> A. gossypii in Japan where it occurs in habitats ranging from
4.4 Aphis gossypii 83<br />
open fields with low vegetation to garden habitats with low shrubs. In both<br />
situations it is attacked by specialised and generalist parasitoids. Of the two<br />
specialised parasitoids, Trioxys communis prefers the open field whereas<br />
Lysiphlebia japonica the garden habitat. Of the generalist parasitoids<br />
Aphelinus sp. prefers the open field and Ephedrus sp. the garden habitat.<br />
Thus the parasitoid complex <strong>of</strong> A. gossypii on cucumber, eggplant or taro in<br />
an open field is quite different from that on Hibiscus or Rhamnus in a garden<br />
(Takada 1992). Hence, in any country, the aim would be to determine<br />
whether there are gaps in the range <strong>of</strong> parasitoids present that might be filled<br />
with species known to be effective elsewhere. If there appear to be important<br />
gaps, there are good reasons for believing that there could be considerable<br />
advantages in filling them. Nevertheless, there are greater complexities than<br />
for many other pests in making clear recommendations, largely because <strong>of</strong><br />
the range <strong>of</strong> biotypes that exist in both aphid hosts and their parasitoids.<br />
Although it is not possible to make specific recommendations for any<br />
country without knowing what parasitoids are already present and the<br />
crop(s) on which control is desired the following parasitoids merit<br />
consideration:<br />
Aphelinus gossypii Lysiphlebia japonica<br />
Aphidius colemani Lysiphlebus fabarum<br />
Aphidius gifuensis Lysiphlebus testaceipes<br />
Ephedrus persicae Trioxys communis<br />
Lipolexis scutellaris Trioxys indicus<br />
Even if any <strong>of</strong> these species is already in a country, but not attacking<br />
either A. craccivora or A. gossypii, a host-adapted strain should be<br />
considered for introduction. It is possible that some <strong>of</strong> the parasitoid species<br />
will compete directly with species that are already present. If this happens<br />
and one parasitoid species is displaced from an aphid host in some situations<br />
or on some crops, the final result will almost always be a lower overall aphid<br />
density.
4.5 Cosmopolites sordidus<br />
India<br />
20°<br />
Myanmar<br />
? Laos<br />
0°<br />
20°<br />
China<br />
P<br />
Thailand<br />
+<br />
Cambodia<br />
+<br />
Vietnam<br />
+++<br />
++<br />
++ Brunei<br />
Malaysia<br />
+<br />
Singapore<br />
++<br />
Indonesia<br />
+<br />
Taiwan<br />
++<br />
Philippines<br />
Australia<br />
Papua<br />
New Guinea<br />
++<br />
The banana weevil borer Cosmopolites sordidus is native to the Indo-Malaysian region.<br />
There have been many attempts at biological control, involving three predatory beetles, but<br />
the results have generally been disappointing. Many predators attack Cosmopolites larvae in<br />
their native range, especially the histerid beetles Plaesius javanus and Plaesius laevigatus in<br />
Indonesia and Dactylosternum hydrophiloides in Malaysia. The first two species have been<br />
established in Fiji, with some reduction in Cosmopolites abundance, but it remains an important<br />
pest there. In Cook Is, they appear to have had little impact. Unless beneficial effects from P.<br />
javanus and P. laevigatus can be demonstrated in Fiji or other countries, there would seem to be<br />
little point in introducing these species to any additional countries. In Kenya, Koppenhšfer et al.<br />
(1992) recorded two important predatory beetles, Dactylosternum abdominale and<br />
Thyreocephalus interocularis,<br />
capable <strong>of</strong> reducing larval abundance by 40% to 90%. In Cuba,<br />
the ant Tetramorium bicarinatum is reported to keep C. sordidus under control. This ant is<br />
widespread, but there is no information on its effectiveness elsewhere.<br />
It would be highly desirable to investigate whether the weevil is indeed absent or <strong>of</strong> very<br />
minor importance in some areas (e.g. Myanmar, southern China) and, if so, what part is played<br />
by resistant cultivars, cultural methods or natural enemies. Ants might be evaluated for their<br />
effects in the Solomon Is where Cosmopolites is unimportant or in Papua New Guinea where it is<br />
<strong>of</strong> local importance only.<br />
85<br />
20°<br />
0°<br />
20°
86 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Cosmopolites sordidus (Germar)<br />
Rating<br />
Origin<br />
Distribution<br />
Coleoptera: Curculionidae<br />
banana weevil borer<br />
<strong>Southeast</strong> Asia China Southern and Western Pacific<br />
+++ Viet +++ Cook Is, Fr. P, Fiji, Guam,<br />
New Cal, Niue, A Sam, Tong<br />
13 ++ Msia, Brun, Indo,<br />
Phil<br />
+ Thai, Sing + FSM<br />
P Camb P P Kir<br />
? Myan, Laos<br />
35 ++ PNG, Sol Is, Sam, Van,<br />
W & F<br />
This account brings up-to-date the chapter on C. sordidus in Waterhouse and<br />
Norris (1987) and increases its relevance to <strong>Southeast</strong> Asia.<br />
According to Purseglove (1972) the genus Musa has a centre <strong>of</strong> diversity in<br />
the Assam-Burma-Thailand area and it probably originated there. The<br />
banana weevil borer is also stated to be a native <strong>of</strong> the Indo-Malaysian region<br />
(Zimmerman 1968; Clausen 1978). Although this region seems a likely<br />
centre <strong>of</strong> origin <strong>of</strong> the weevil, and those investigating its biological control<br />
have consistently assumed so, there had already been, by the time <strong>of</strong><br />
GermarÕs 1824 description based on material from India, centuries <strong>of</strong><br />
intercontinental travel by Europeans, by means <strong>of</strong> which the weevil could<br />
have spread to many other lands in infested plants, thus obscuring its origin.<br />
There are only two species in the genus Cosmopolites,<br />
the lesser known<br />
C. pruinosus occurring in Borneo and the Philippines and, after introduction,<br />
in Micronesia (Zimmerman 1968).<br />
C. sordidus is present in virtually all banana-growing areas <strong>of</strong> the world,<br />
including most, if not all, <strong>of</strong> <strong>Southeast</strong> Asia and most <strong>of</strong> the Pacific.<br />
Exceptions in the Pacific are Marshall Is, Tuvalu and Tokelau (Anon. 1979b;<br />
Waterhouse and Norris 1987; Waterhouse 1997). In <strong>Southeast</strong> Asia no<br />
information is available from Laos and the situation in Myanmar is unclear.<br />
A recommendation was made in the standard work on the Ô<strong>Insect</strong> <strong>Pests</strong> <strong>of</strong><br />
BurmaÕ (Ghosh 1940) to guard against the introduction to that country <strong>of</strong><br />
C. sordidus and neither N. von Keyserlingk nor G. Pierrard (pers. comm.
Biology<br />
4.5<br />
Cosmopolites sordidus<br />
1992) were able to establish, when based in Yangon, that the species<br />
occurred in Myanmar. Its uncertain status in Myanmar and Laos and its<br />
status <strong>of</strong> Ôpresent, but unimportantÕ, in Cambodia and China clearly merits<br />
further investigation.<br />
The ovoid, 2 mm long, white eggs are laid singly, usually between the leafsheath<br />
scars on the crown <strong>of</strong> the banana rhizome, in small cavities chewed<br />
out by the female just above the ground surface. Laying also occurs on the<br />
pseudostems <strong>of</strong> fallen plants. The eggs hatch in about 8 days in summer and<br />
the larvae tunnel into the tissues. On reaching maturity, after about 20 days<br />
feeding in warm weather (Jepson 1914), the larvae tunnel to near the surface<br />
<strong>of</strong> the corm and form an oval chamber in which they pupate. The period from<br />
egg to adult may be as short as 29 days in the New South Wales summer or as<br />
long as 6 months in the cooler parts <strong>of</strong> the year (Hely et al. 1982). Fifty days<br />
is a more usual maximum for the life cycle in Fiji (Swaine 1971). The<br />
nocturnal adults also tunnel in banana tissues. During the day they generally<br />
hide in or around the rhizomes or between the leaf sheaths at or just above<br />
ground level. They are slow moving and will feign death when disturbed.<br />
They seldom fly, but walk over the soil surface and vegetation. Whalley<br />
(1957) found that adults dispersed slowly in Uganda. Of 400 marked weevils<br />
35% were recovered over an 8-month period within a radius <strong>of</strong> 7 m from the<br />
release point. In Colombia a few marked adults were recaptured after 2<br />
weeks at a distance <strong>of</strong> 6 to 8 m, but 23 months after release none were found<br />
in another plantation 20 m away (Cardenas and Arango 1986). Male<br />
C. sordidus release an aggregation pheromone, sordidin (Beauhaire et al.<br />
1995), from the hindgut which attracts both males and females, but females<br />
do not produce a pheromone attractive to either sex (Budenberg et al. 1993b;<br />
Ndiege et al. 1996). Both sexes were attracted to freshly cut rhizome and<br />
pseudostem and females to rotting pseudostem (Budenberg et al. 1993a).<br />
Eggs are laid throughout the year at a rate varying with temperature and up to<br />
100 a year. Adults may live as long as 2 years (Swaine 1971). They can<br />
survive in captivity for 14 weeks without food (Zimmerman 1968).<br />
Laboratory rearing <strong>of</strong> C. sordidus was studied by Afreh (1993) and<br />
Koppenhšfer and Reddy (1994).<br />
87
88 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Damage<br />
The status <strong>of</strong> C. sordidus as one <strong>of</strong> the most important pests <strong>of</strong> bananas is<br />
<strong>of</strong>ten reported (Swaine 1971; Purseglove 1972) and, indeed, many adults<br />
and larvae are <strong>of</strong>ten present. It is reported to be now the most important pest<br />
<strong>of</strong> bananas in Africa (Nahif et al. 1994; Ortiz et al. 1995). However, in order<br />
to place these reports in context, it is necessary to outline the stages <strong>of</strong><br />
growth <strong>of</strong> the plant. Bananas are propagated vegetatively by planting<br />
rhizomes (corms), which give rise to shoots after a few weeks. As the plant<br />
grows, a pseudostem is formed from the sheaths <strong>of</strong> the leaves which<br />
continue to develop internally until a flowering shoot emerges from the top<br />
<strong>of</strong> the pseudostem at a height <strong>of</strong> 2 to 4 m depending upon the variety. When<br />
each bunch <strong>of</strong> bananas is cut, the pseudostem bearing that bunch is also cut<br />
down. At the same time a healthy sucker growing from the same base is<br />
selected to succeed and other suckers removed. C. sordidus larvae tunnel in<br />
the rhizome and the base <strong>of</strong> the pseudostem, but do not attack the roots. This<br />
tunneling may kill young plants and greatly increases the susceptibility <strong>of</strong><br />
mature plants to wind damage. Adults cause little damage and feed mainly<br />
on rotting banana tissue. Injury by larvae to the rhizome can interfere with<br />
root initiation and sap flow within the plant and grossly infested plants may<br />
bear small bunches <strong>of</strong> undersized fruit (Wright 1976). Much <strong>of</strong> the damage<br />
attributed to C. sordidus is probably caused by rhizome rot or nematodes<br />
(Ostmark 1974). In East Africa the combined attack <strong>of</strong> nematodes and <strong>of</strong><br />
banana weevil borer is considered to be the main reason for the serious<br />
decline in productivity <strong>of</strong> bananas (De Langhe 1988), but the precise role <strong>of</strong><br />
the weevil is still to be established. Suckers infested with nematodes were<br />
found to be more than four times more likely to be attacked by C. sordidus<br />
than suckers without nematodes (Speijer et al. 1993). Although the banana<br />
weevil borer has occasionally been responsible for severe losses <strong>of</strong> newlyplanted<br />
rhizomes, extensive experiments in Central America agree with<br />
some reports from Australia (Smith 1993; Wallace 1937) that weevil<br />
damage is not as important as frequently claimed since the larvae have a<br />
strong preference for rhizomes <strong>of</strong> harvested plants over healthy rhizomes<br />
(Ostmark 1974). There are reports that some banana cultivars are<br />
comparatively resistant to attack by C. sordidus (Mesquita et al. 1984;<br />
Mesquita and Caldas 1987), but the mechanism <strong>of</strong> such resistance<br />
(repellency, toxicity, greater tolerance to damage, etc.) is not known.<br />
In Australia, Braithwaite (1963) concluded that the importance <strong>of</strong><br />
C. sordidus infestation is aggravated by poor culture, but that benefit could<br />
be derived from almost complete control with insecticides (Braithwaite<br />
1958). In the same region Loebel (1975) concluded that heavy weevil
Host plants<br />
4.5<br />
Cosmopolites sordidus<br />
infestation is a symptom, rather than a cause, <strong>of</strong> a declining plantation,<br />
because 2 years <strong>of</strong> effective use <strong>of</strong> chemicals failed to improve growth or<br />
yield in his experimental plots. In Costa Rica several insecticides were<br />
effective in controlling C. sordidus populations, but banana yields were not<br />
increased (Nanne and Klink 1975). Nevertheless C. sordidus is always likely<br />
to be <strong>of</strong> importance in areas that experience strong winds. The abundance <strong>of</strong><br />
adult weevils can be estimated by counting the number attracted to cut<br />
segments <strong>of</strong> pseudostem and <strong>of</strong> larvae by estimating the area damaged and<br />
counting the number <strong>of</strong> galleries exposed by slitting the rhizome or the<br />
pseudostem very near to its base (Vilardebo 1973; Delattre 1980; Mesquita<br />
1985; Smith 1993). In spite <strong>of</strong> this, an adequate relationship between adult<br />
and larval abundance and economic loss remains to be established. It must<br />
be added, however, that there continues to be a widespread view that<br />
C. sordidus is a major pest.<br />
The weevil attacks all banana ( Musa sapientum)<br />
cultivars, including<br />
plantain, and also Manila hemp ( Musa textilis).<br />
It has been recorded in<br />
earlier days (but not in recent years) from plants in other Orders, but these<br />
reports are almost certainly in error.<br />
Natural enemies<br />
Although many predators are known to attack C. sordidus eggs, larvae and<br />
pupae (Table 4.5.1), it is extraordinary that, with one possible exception, not<br />
a single parasitoid <strong>of</strong> any life history stage has been recorded. That exception<br />
is the early report from the Philippines <strong>of</strong> Cendana (1922), who found a<br />
chalcidid wasp in one <strong>of</strong> his C. sordidus breeding cages, but it may not have<br />
been parasitising the weevil. It is, perhaps, less surprising that the heavily<br />
sclerotised adult weevil has very few enemies. It is true that some weevils<br />
are effectively attacked by hymenopterous parasitoids and that some<br />
tachinid parasitoids are able to attack certain weevils by laying eggs in their<br />
food or beneath their mouth when they are feeding (Jacobs and Renner<br />
1988), but neither has been observed for C. sordidus.<br />
Koppenhšfer<br />
(1993a,b) estimated that 58% <strong>of</strong> the eggs were accessible to predators and<br />
presumably at least as many should be available to parasitoids if there were<br />
any. Most eggs were found in the surface layer <strong>of</strong> rhizomes, particularly in<br />
the crown, but some are also laid at the base <strong>of</strong> pseudostems and in the walls<br />
<strong>of</strong> abandoned larval tunnels in both pseudostems and rhizomes. As soon as<br />
eggs hatch the young larvae immediately tunnel deeper into the plant tissue<br />
and thus become far less available to natural enemies.<br />
89
Table 4.5.1<br />
<strong>Insect</strong> predators <strong>of</strong> Cosmopolites sordidus<br />
<strong>Insect</strong><br />
DERMAPTERA<br />
LABIIDAE<br />
Stage<br />
attacked<br />
Carcinophora (= Psalis)<br />
americana<br />
larvae Jamaica<br />
Puerto Rico<br />
Euborellia (= Anisolabis)<br />
annulipes<br />
egg, larva Jamaica<br />
Kenya<br />
Country Reference<br />
Edwards 1934<br />
Anon. 1939<br />
Edwards 1934<br />
Koppenhšfer et al. 1992; Sirjusingh<br />
et al. 1992<br />
Labia borellii<br />
egg, larva Kenya Koppenhšfer et al. 1992<br />
Labia curvicauda<br />
HEMIPTERA<br />
CYDNIDAE<br />
egg, larva Kenya Koppenhšfer et al. 1992<br />
Geotomus pygmaeus<br />
MIRIDAE<br />
egg Malaysia, widespread China 1935<br />
Fulvius nigricornis<br />
NABIDAE<br />
egg Malaysia China 1935<br />
Phorticus pygmaeus<br />
REDUVIIDAE<br />
egg Malaysia, Papua New Guinea China 1935<br />
Physoderes curculionis<br />
COLEOPTERA<br />
CARABIDAE<br />
larva Malaysia China 1935<br />
Abacetus?<br />
optimus<br />
Kenya Koppenhšfer et al. 1992<br />
Galerita (= Propagalerita)<br />
bicolor<br />
Sirjusingh et al. 1992<br />
Scarites sp. Sirjusingh et al. 1992<br />
90 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong>
Table 4.5.1 (contÕd) <strong>Insect</strong> predators <strong>of</strong> Cosmopolites sordidus<br />
<strong>Insect</strong> Stage<br />
attacked<br />
COLEOPTERA<br />
ELATERIDAE<br />
unidentified spp. Australia,<br />
New Caledonia<br />
HISTERIDAE<br />
Hister niloticus<br />
Hololepta (= Lioderma)<br />
quadridentata<br />
Country Reference<br />
Froggatt 1928a<br />
Jacques 1931<br />
larva Kenya Koppenhšfer et al. 1992<br />
Malaysia<br />
Trinidad<br />
Clement 1944<br />
Pea & Duncan 1991<br />
Hololepta striaditera<br />
larva, pupa Kenya Koppenhšfer et al. 1992<br />
Hololepta sp. St Vincent Sirjusingh et al. 1992<br />
Lioderma sp. Sirjusingh et al. 1992<br />
Plaesius (= Hyposolenus)<br />
laevigatus<br />
Indonesia Froggatt 1928b<br />
Plaesius javanus<br />
larva, pupa Malaysia<br />
Froggatt 1928b; Clement 1944;<br />
Indonesia<br />
Jepson 1914<br />
Thailand<br />
Charernsom 1992<br />
Platysoma abrupta<br />
larva Malaysia Corbett 1936; Lamas 1947<br />
Platysoma sp. Corbett 1936<br />
unidentified histerid sp.<br />
HYDROPHILIDAE<br />
egg, larva, pupa Kenya Koppenhšfer et al. 1992<br />
Dactylosternum abdominale<br />
larva Kenya,<br />
Corbett 1936; Edwards 1939,<br />
Philippines<br />
Koppenhšfer et al. 1992<br />
D. hydrophiloides larva Indonesia,<br />
Malaysia<br />
Corbett 1936<br />
D. intermedium larva Guinea CuillŽ 1950<br />
D. pr<strong>of</strong>undus San Thom Beccari 1967<br />
D. subdepressum Trinidad Cock 1985<br />
D. subquadratum larva Malaysia Corbett 1936<br />
Omicrogiton insularis larva Malaysia Corbett 1936<br />
4<br />
.5<br />
Cosmopolites sordidus<br />
91
Table 4.5.1 (contÕd) <strong>Insect</strong> predators <strong>of</strong> Cosmopolites sordidus<br />
<strong>Insect</strong> Stage<br />
attacked<br />
COLEOPTERA<br />
SILVANIDAE<br />
Cathartus sp. larva Indonesia Jepson 1914<br />
STAPHYLINIDAE<br />
Belonuchus ferrugatus larva Indonesia Jepson 1914; CuillŽ 1950<br />
B. quadratus larva Malaysia Corbett 1936; Lamas 1947<br />
Charichirus sp. egg. larva Kenya Koppenhšfer et al. 1992<br />
Eulissus sp. Kenya Reddy 1988<br />
Hesperus? sparsior egg, larva Kenya Koppenhšfer et al. 1992<br />
Priochirus (= Leptochirus) unicolor larva Indonesia Jepson 1914, CuillŽ 1950<br />
Thyreocephalus? interocularis egg, larva Kenya Koppenhšfer et al. 1992<br />
TENEBRIONIDAE<br />
Eutochia pulla egg Kenya Koppenhšfer et al. 1992<br />
DIPTERA<br />
RHAGIONIDAE<br />
Chrysopilus ferruginosus larva Indonesia,<br />
India,<br />
Philippines<br />
Country Reference<br />
Froggatt 1928b; Beccari 1967<br />
Jepson 1914<br />
Chrysopilus sp. Brazil Sirjusingh et al. 1992<br />
HYMENOPTERA<br />
FORMICIDAE<br />
Anochaetus sp. Kenya Reddy 1988<br />
Pheidole megacephala egg, larva Cuba Castineiras et al. 1991a<br />
Tetramorium bicarinatum<br />
(= T. guineense)<br />
egg, larva Cuba Roche & Abreu 1983<br />
92 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong>
4.5 Cosmopolites sordidus 93<br />
Because <strong>of</strong> the reported existence <strong>of</strong> natural enemies <strong>of</strong> C. sordidus in<br />
Indonesia and nearby countries, Jepson (1914) was sent from Fiji to<br />
investigate the situation in Java. The histerid beetle Plaesius javanus was<br />
commonly found preying on C. sordidus and other insects in the soil and leaf<br />
litter. Two staphylinid beetles Belonuchus ferrugatus and Priochirus<br />
(= Leptochirus) unicolor and a silvanid beetle Cathartus sp. were also<br />
shown to attack C. sordidus larvae, but they were not nearly as voracious.<br />
The predatory larvae <strong>of</strong> the rhagionid fly Chrysopilus ferruginosus attacked<br />
C. sordidus larvae in the laboratory, but not in the field.<br />
A later investigation in Java (Froggatt 1928b) failed to reveal any egg<br />
parasites, but P. javanus and C. ferruginosus were recorded, as well as two<br />
other species <strong>of</strong> Histeridae, one probably Plaesius (= Hyposolenus)<br />
laevigatus, one or two species <strong>of</strong> Staphylinidae and two species <strong>of</strong><br />
Hydrophilidae (all Coleoptera). Several species <strong>of</strong> Dermaptera were fairly<br />
common in the rotting banana plant. In southern China (Yunnan Province)<br />
two Dermaptera (one a forficulid) are reported to eat C. sordidus larvae and<br />
pupae, and a mite to attack larvae and adults; also a white muscardine fungus<br />
to infect 1% <strong>of</strong> larvae and pupae (Sun 1994).<br />
The only detailed study in recent times <strong>of</strong> the natural enemies <strong>of</strong><br />
C. sordidus is that <strong>of</strong> Koppenhšfer et al. (1992) in Kenya. Not surprisingly,<br />
the species (<strong>of</strong> predators) recorded were all polyphagous, since many were<br />
native species that had come to include an introduced pest amongst their<br />
prey. Twelve predators <strong>of</strong> eggs, larvae and pupae <strong>of</strong> C. sordidus were found.<br />
None <strong>of</strong> these attacked adults and no parasitoids were recorded<br />
(Koppenhšfer et al. 1992). Of these predators, the hydrophilid beetle<br />
Dactylosternum abdominale reduced weevil numbers in suckers by up to<br />
50% and in residual stumps <strong>of</strong> harvested suckers by 39%. In spent<br />
pseudostems, D. abdominale reduced numbers by 40% to 90% at different<br />
predator densities and the large staphylinid predator Thyreocephalus<br />
interocularis reduced numbers by 42%. Other predators (Table 4.5.1) were<br />
unimportant (Koppenhšfer and Schmutterer 1993).<br />
These particular predators are clearly non-specific, since they are native<br />
to East Africa and C. sordidus is not. Thus, although they may be attractive<br />
candidates for introduction (and Plaesius javanus was in this same<br />
category), the widely held view now is that, because <strong>of</strong> their impact on nontarget<br />
organisms, very careful consideration should be given before any<br />
highly non-specific predators are introduced to a new region.<br />
Extensive testing has been carried out <strong>of</strong> fungi (Metarhizium anisopliae<br />
and Beauveria bassiana) and <strong>of</strong> entomopathogenic nematodes<br />
(Heterorhabditis spp. and Steinernema spp.) as biological ÔpesticidesÕ<br />
against C. sordidus. Laboratory or field trials showed that strains <strong>of</strong> both
94 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
fungi were capable <strong>of</strong> killing adults and larvae (Delattre and Jean-Bart 1978;<br />
Filho et al. 1987; Busoli et al. 1989; Castineiras et al. 1991a,b; Pea and<br />
Duncan 1991; Ponce et al. 1992; Brenes and Carballo 1994; Carballo and de<br />
Lopez 1994; Pea et al. 1995). In the field the best results were obtained with<br />
application <strong>of</strong> fungi twice a year at a dose <strong>of</strong> 10 13 conidia per ha. This<br />
reduced trap catches <strong>of</strong> adults by 52% and rhizome damage by 65%, leading<br />
to a 25% yield increase. Parallel experiments with 9 colonies per ha <strong>of</strong> the<br />
predatory ant Pheidole megacephala yielded similar results (Castineiras et<br />
al. 1991a,b).<br />
Early laboratory tests in Guadelupe by Laumond et al. (1979) showed<br />
that C. sordidus is susceptible to the entomopathogenic nematode<br />
Steinernema carpocapsae (= S. feltiae), an observation since widely<br />
confirmed in Central America for this and other nematode species (e.g.<br />
Figueroa 1990; Pea and Duncan 1991). However, the most extensive recent<br />
work has been carried out in Australia and Tonga. Twenty-one different<br />
species <strong>of</strong> Steinernema and Heterorhabditis, 7 strains <strong>of</strong> Steinernema<br />
carpocapsae, 2 <strong>of</strong> S. feltiae, 4 <strong>of</strong> H. bacteriophora and 2 <strong>of</strong><br />
H. zealandica were tested against adult banana weevils. The best <strong>of</strong> these,<br />
S. carpocapsae BW strain, gave 85% infection in the laboratory (Parnitzki et<br />
al. 1990, 1998; Treverrow et al. 1991). Adult C. sordidus are highly resistant<br />
to entomopathogenic nematodes, due to the difficulty <strong>of</strong> nematodes entering<br />
the host via the mouth or anus. The large spiracles <strong>of</strong> the first abdominal<br />
segment <strong>of</strong>fer an effective site <strong>of</strong> entry for the nematodes if they are able to<br />
pass under the tightly fitting elytra. By adding paraffin oil to the nematode<br />
preparation to seal the elytra, the beetle is caused to raise them slightly to<br />
respire, simultaneously giving the nematodes access to the spiracles. Adult<br />
weevils are strongly attracted to holes or cuts in the rhizome or psuedostem,<br />
but they require a thigmotactic stimulus to remain long enough to become<br />
infected by nematodes. If a core <strong>of</strong> tissue is removed from two sites at the<br />
base <strong>of</strong> a residual corm using a desuckering tool, 250000 nematodes<br />
(S. carpocapsae BW) added to each hole and the core loosely inserted,<br />
nearly all adult weevils attracted are killed. In one large scale field trial in<br />
New South Wales 8% <strong>of</strong> plants in untreated plots suffered significant<br />
damage, 3% when prothiophos was added to the core, 1% when nematodes<br />
were added and 0% when prothiophos was applied to the soil around the base<br />
<strong>of</strong> the plant. It was concluded that control <strong>of</strong> banana weevil using<br />
entomopathogenic nematodes should now be economically feasible<br />
(Treverrow and Bedding 1992, 1993a,b). More recently, Treverrow (1994)<br />
has found that baiting, and stem injection with very small amounts <strong>of</strong><br />
insecticide, can reduce treatment costs to less than 1 cent per stool. This<br />
makes nematode applications against adults uncompetitive unless market
4.5 Cosmopolites sordidus 95<br />
premiums can be obtained for fruit produced in the absence <strong>of</strong> insecticides.<br />
However, targetting the highly susceptible larvae instead <strong>of</strong> the more<br />
resistant adult weevils may significantly reduce the costs <strong>of</strong> nematode<br />
applications. Applications <strong>of</strong> S. carpocapsae with a water-absorbing<br />
polyacrylamide gel into cuts or holes made in residual rhizomes gave<br />
significant mortality <strong>of</strong> C. sordidus larvae (Treverrow et al. 1991; Treverrow<br />
and Bedding 1993b). However, mortality at this stage may have limited<br />
effect on abundance, since a survey <strong>of</strong> 50 properties showed that 70% <strong>of</strong><br />
adult weevils had already emerged from pre-harvest corms (Treverrow and<br />
Bedding 1993a; Treverrow 1994). The importance <strong>of</strong> correct formulation for<br />
the nematodes is highlighted by the disappointing results obtained in<br />
Queensland, when an earlier formulation without a water-absorbing gel<br />
failed to give effective control <strong>of</strong> adults, possibly due to the accumulation <strong>of</strong><br />
excess water in the core holes (Smith 1993).<br />
Attempts at biological control<br />
FIJI<br />
C. sordidus became a very destructive pest <strong>of</strong> bananas following its<br />
introduction about 1901 and this led to the first attempt at its biological<br />
control. This consisted <strong>of</strong> the introduction <strong>of</strong> the predatory histerid beetle<br />
Plaesius javanus into Fiji in 1913 from Java (Jepson 1914). This was<br />
unsuccessful (Table 4.5.2), but a further introduction in 1918 led to its<br />
establishment (Veitch 1926; Bennett et al. 1976). Simmonds (1935) reported<br />
a marked reduction in weevil damage, an opinion supported by Anon.<br />
(1935), but Pemberton (1954) considered that only partial control had been<br />
achieved. It now seems probable that, in addition to P. javanus, the similar<br />
looking histerid Plaesius laevigatus was also introduced, at least to Cook Is<br />
(Walker and Deitz 1979).<br />
AUSTRALIA<br />
Cosmopolites sordidus is thought to have become established in Queensland<br />
about the end <strong>of</strong> the 19th century, having arrived possibly from Papua New<br />
Guinea, but it was not until about 1914Ð15 that it reached New South Wales<br />
as a result <strong>of</strong> an accidental introduction from Fiji (Wilson 1960). Following<br />
its successful introduction into Fiji, Plaesius javanus was introduced from<br />
Java and liberated in Queensland from 1921 to 1928, but became established<br />
only briefly. It was introduced into New South Wales in 1922 from Java and<br />
again in 1934 from Fiji, but again it did not become established. The fly<br />
Chrysopilus ferruginosus was also introduced from Java, but failed to<br />
become established.
Table 4.5.2 Introductions for the biological control <strong>of</strong> Cosmopolites sordidus<br />
Country and species Liberated From Result Reference<br />
AUSTRALIA<br />
Plaesius javanus 1921Ð28<br />
Java<br />
Ð Clausen 1978<br />
Weddell 1932<br />
1934<br />
Fiji<br />
Ð Wilson 1960<br />
Chrysopilus ferruginosus 1928 Java Ð Wilson 1960<br />
Dactylosternum hydrophiloides<br />
CAMEROON<br />
1939 Malaysia + Smith 1944; Wilson 1960<br />
Plaesius javanus 1952 Trinidad Ð Bennett et al. 1976<br />
Hololepta quadridentata 1952 Trinidad Ð Bennett et al. 1976<br />
COOK IS<br />
Plaesius javanus 1937Ð40 Fiji + Walker & Deitz 1979<br />
Plaesius laevigatus 1937Ð40 Fiji + Walker & Deitz 1979<br />
CUBA<br />
Plaesius laevigatus Ð Sirjusingh et al. 1992<br />
DOMINICA<br />
Plaesius javanus 1951 Trinidad Ð Cock 1985<br />
1958Ð59 ? Ð Clausen 1978<br />
Hololepta quadridentata 1951<br />
1958Ð59<br />
FIJI<br />
Trinidad<br />
?<br />
Ð<br />
Ð<br />
Cock 1985<br />
Clausen 1978<br />
Plaesius javanus 1913 Java Ð Jepson 1914<br />
1918 Java + Veitch 1926<br />
Plaesius laevigatus 1918 Java + Walker & Deitz 1979<br />
96 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong>
Table 4.5.2 (contÕd) Introductions for the biological control <strong>of</strong> Cosmopolites sordidus<br />
Country and species<br />
FRENCH POLYNESIA<br />
Liberated From Result Reference<br />
Plaesius javanus 1937 Fiji + Delobel 1977<br />
GRENADA<br />
Hololepta quadridentata 1949, 1951 Trinidad Ð Cock 1985<br />
Plaesius javanus 1949, 1951 Trinidad Ð Cock 1985<br />
HONDURAS<br />
Plaesius javanus 1942 ? Ð Greathead 1971<br />
JAMAICA<br />
Plaesius javanus 1918Ð19 Java Ð Cock 1985; Edwards 1934<br />
1937Ð38 Java via Fiji +<br />
Ð<br />
Cock 1985<br />
Sirjusingh et al. 1992<br />
Dactylosternum hydrophiloides 1918Ð19 Malaysia Ð Edwards 1934<br />
1937Ð38 Malaysia + Edwards 1934<br />
Dactylosternum abdominale 1937Ð38 Malaysia Ð Edwards 1934<br />
MARIANAS<br />
Plaesius javanus 1947 Fiji + Clausen 1978<br />
Hololepta quadridentata 1953 ? Ð Clausen 1978<br />
Hololepta minuta 1953 ? Ð Clausen 1978<br />
Hololepta sp. 1953 Trinidad Ð Clausen 1978<br />
MAURITIUS<br />
Plaesius javanus 1959 Trinidad Ð Bennett et al. 1976<br />
? Trinidad Ð Gomy 1983<br />
Hololepta quadridentata 1942 Trinidad Ð Bennett et al. 1976<br />
1959 ? Ð Clausen 1978<br />
4.5 Cosmopolites sordidus 97
Table 4.5.2 (contÕd) Introductions for the biological control <strong>of</strong> Cosmopolites sordidus<br />
Country and species<br />
MEXICO<br />
Liberated From Result Reference<br />
Plaesius javanus<br />
NEW CALEDONIA<br />
1955 ? + Barrera & Jiminez 1994;<br />
Greathead 1971<br />
Plaesius javanus<br />
PALAU IS<br />
1949 Fiji + Dumbleton 1957<br />
Dactylosternum hydrophiloides 1948 Malaysia Ð Dumbleton 1957<br />
Hololepta sp. 1953 ? Ð Dumbleton 1957<br />
PUERTO RICO<br />
Plaesius javanus 1936 ? Ð Greathead 1971<br />
SAMOA<br />
Plaesius javanus 1957 Fiji + Dale and Herring 1959<br />
SEYCHELLES<br />
Plaesius javanus 1952 Trinidad Ð Greathead 1971<br />
Hololepta quadridentata 1952 Trinidad Ð Greathead 1971<br />
ST LUCIA<br />
Hololepta quadridentata 1950Ð1954 Trinidad Ð Cock 1985<br />
Plaesius javanus 1950Ð1954 Trinidad Ð Cock 1985<br />
ST VINCENT<br />
Dactylosternum subdepressum 1950Ð1954 Trinidad Ð Cock 1985<br />
Hololepta quadridentata 1942 Trinidad + Bennett et al. 1976<br />
Plaesius javanus 1981 Trinidad Ð Cock 1985<br />
98 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong>
Table 4.5.2 (contÕd) Introductions for the biological control <strong>of</strong> Cosmopolites sordidus<br />
Country and species<br />
TAIWAN<br />
Liberated From Result Reference<br />
Plaesius javanus 1938 ? Ð Greathead 1971<br />
TANZANIA<br />
Plaesius javanus 1948 ? Ð Greathead 1971<br />
TONGA<br />
Plaesius javanus 1952 Fiji Ð Dumbleton 1957<br />
TRINIDAD<br />
+ O. Fakalata pers. comm.<br />
Plaesius javanus 1942 Jamaica + Bennett et al. 1976<br />
UGANDA<br />
Ð Sirjusingh et al. 1992<br />
Plaesius javanus<br />
VANUATU<br />
1934Ð35 Java Ð Greathead 1971<br />
Plaesius javanus ? ? ? Anon. 1979a<br />
WALLIS IS<br />
Plaesius javanus 1947 Fiji ? Cohic 1959<br />
4.5 Cosmopolites sordidus 99
100 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
The predatory hydrophilid beetle Dactylosternum hydrophiloides from<br />
Malaysia was liberated in 1939 and has become established but has not had a<br />
major effect on weevil abundance (Wilson 1960).<br />
Braithwaite (1958) reports an unusual native predator <strong>of</strong> C. sordidus, a<br />
blue planarian worm Kontikia (= Geoplana) caerulea, which lives in moist<br />
sheltered situations. It sucks out the body fluids <strong>of</strong> its prey, leaving the<br />
cuticle intact.<br />
CAMEROON, MAURITIUS, SEYCHELLES, UGANDA<br />
Both P. javanus and Hololepta quadridentata were supplied from Trinidad<br />
to Cameroon (1952), Mauritius (1959) and the Seychelles (1950Ð54) and<br />
P. javanus from Java to Uganda in 1934Ð35. Neither species became<br />
established (Greathead 1971).<br />
COOK IS<br />
CUBA<br />
INDIA<br />
JAMAICA<br />
Although only Plaesius javanus is recorded as having been introduced from<br />
Fiji into the Cook Is during the period 1937 to 1940, voucher specimens<br />
(DSIR, NZ) show that Plaesius laevigatus was also present in the material<br />
liberated and both species still occur in the Cook Is, although the latter does<br />
not appear in voucher specimens in Fiji. Unfortunately the banana weevil<br />
borer is still an important problem (Walker and Deitz 1979; Waterhouse<br />
1995, 1997).<br />
In Cuba the ant Tetramorium bicarinatum (= T. guineense) was found to be<br />
capable <strong>of</strong> destroying up to 65% <strong>of</strong> C. sordidus in heavily infested<br />
plantations. With lower populations, 73.8% and 83.5% control was obtained<br />
in successive years. For colonisation <strong>of</strong> a plantation in 3 to 4 months, ants<br />
should be released over 25% to 30% <strong>of</strong> the area (Roche and Abreu 1983).<br />
The ant is a more effective predator during the dry than the wet season<br />
(Lopez and Ramos 1986). In countries where this widespread ant is already<br />
present (e.g. the Americas, Africa, Papua New Guinea, Australia, the<br />
oceanic Pacific) it might well be considered for manipulating C. sordidus<br />
abundance, but very careful consideration should be given to any proposal to<br />
introduce such an agressive broad-spectrum predator into a new country.<br />
The predatory beetle Dactylosternum hydrophiloides was introduced from<br />
Malaysia in 1948, but there is no record <strong>of</strong> the outcome (Whilshaw 1949).<br />
Although an initial release in 1918Ð19 <strong>of</strong> Plaesius javanus from Java was<br />
unsuccessful, this predator became established as a result <strong>of</strong> a further release<br />
<strong>of</strong> Fijian material in 1937Ð38 (Bennett et al. 1976). More recently Sirjusingh<br />
et al. (1992) recorded that it was no longer present.
4.5 Cosmopolites sordidus 101<br />
MYANMAR<br />
There do not appear to be any records in Myanmar <strong>of</strong> natural enemies <strong>of</strong><br />
C. sordidus which is very uncommon and apparently confined to aromatic<br />
and sweet-flavoured banana varieties. <strong>Control</strong> is achieved by cutting the<br />
pseudostems every three years, a practice readily accepted because the stems<br />
are used in Mohinga, a popular fish soup (H. Morris pers. comm. 1994).<br />
PAPUA NEW GUINEA<br />
Bananas are grown widely and are the staple food in some areas but<br />
C. sordidus is not a serious pest. The ant Tetramorium bicarinatum occurs<br />
there but there is no information on any possible interaction with C. sordidus<br />
(J.W. Ismay pers. comm. 1985).<br />
TRINIDAD<br />
P. javanus was established in Trinidad in 1942 and, from there, together with<br />
a native histerid Hololepta quadridentata, it was sent to other islands in the<br />
West Indies. Evidence <strong>of</strong> establishment was not available to Simmonds<br />
(1958) but, in 1972, H. quadridentata was recovered in St Vincent (Bennett<br />
et al. 1976).<br />
OTHER COUNTRIES<br />
Although details are not available, P. javanus has been widely distributed<br />
and is reported to be established in French Polynesia, Marianas, New<br />
Caledonia (where chemical control is still required) (Delobel 1977; Clausen<br />
1978; M. Kauma pers. comm. 1985) and Tonga (O. Fakalata pers. comm.<br />
1985). It is also widespread on Upolu Is, Samoa (T.V. Bourke pers. comm.<br />
1986).<br />
In addition to the records in table 4.5.2, Sirjusingh et al. (1992) list a<br />
number <strong>of</strong> predators for Central or South America (Table 4.5.3). However, it<br />
is seldom clear which <strong>of</strong> these may be introductions (intentional or<br />
otherwise) from elsewhere and which are native to the region, as some<br />
almost certainly are. Their biological control potential has not been assessed.
102 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Table 4.5.3 Additional natural enemies <strong>of</strong> C. sordidus in Central and<br />
South America<br />
<strong>Insect</strong><br />
DERMAPTERA<br />
LABIIDAE<br />
Country ?Native<br />
Carcinophora (= Psalis) americana Brazil probably<br />
Euborellia annulipes<br />
HEMIPTERA<br />
Brazil ?<br />
CYDNIDAE<br />
Geotomus pygmaeus Brazil probably not<br />
MIRIDAE<br />
Fulvius nigricornis Brazil probably not<br />
NABIDAE<br />
Phorticus pygmaeus Brazil probably not<br />
REDUVIIDAE<br />
Physoderes curculionis Brazil probably not<br />
COLEOPTERA<br />
CARABIDAE<br />
Galerita bicolor Florida probably<br />
Scarites spp.<br />
HISTERIDAE<br />
Florida probably<br />
Hololepta spp. St Vincent possibly<br />
Lioderma sp. Brazil probably<br />
Platysoma abrupta<br />
HYDROPHILIDAE<br />
Brazil probably not<br />
Dactylosternum abdominale Trinidad probably not<br />
D. hydrophiloides Trinidad probably not<br />
D. intermedium Trinidad probably not<br />
D. pr<strong>of</strong>undus Trinidad probably<br />
Omicrogiton insularis<br />
SILVANIDAE<br />
Brazil probably not<br />
Cathartus sp.<br />
STAPHYLINIDAE<br />
Brazil (not established)<br />
Belonuchus ferrugatus Brazil (not established)<br />
B. quadratus Brazil probably not<br />
Priochirus (= Leptochirus) unicolor<br />
DIPTERA<br />
RHAGIONIDAE<br />
Brazil (not established)<br />
Chrysopilus sp. Brazil (not established)
Biology <strong>of</strong> main natural enemies<br />
4.5 Cosmopolites sordidus 103<br />
Dactylosternum abdominale Col.: Hydrophilidae<br />
This is the most effective predator in Kenya. Its larvae are polyphagous<br />
predators and consume the contents <strong>of</strong> C. sordidus larvae and at high<br />
predator density may become cannibalistic. They also feed on the micr<strong>of</strong>auna<br />
and micro-flora <strong>of</strong> decomposing plant tissues. On the other hand, the<br />
adults cause significant mortality <strong>of</strong> C. sordidus eggs; although many are<br />
laid in inaccessible positions inside the pseudostem and the polyphagous<br />
adults do not specifically search for eggs. Adults cannot penetrate the<br />
narrow tunnels <strong>of</strong> young larvae, so can only capture newly hatched larvae:<br />
later instar larvae are not attacked (Koppenhšfer and Schmutterer 1993;<br />
Koppenhšfer et al. 1992, 1995). Development from egg to adult takes 17 to<br />
33 days, life span is 95 days and females lay an average <strong>of</strong> 1.7 egg cases per<br />
week, each case containing 4 eggs. The preoviposition period is 16.6 days<br />
(Koppenhšfer et al. 1995).<br />
Geotomus pygmaeus Hem.: Cydnidae<br />
This predator is recorded from India, Ceylon, Myanmar, Indonesia,<br />
Vietnam, China, Japan, New Caledonia, Fiji, Samoa, French Polynesia and<br />
Hawaii. Its extensive distribution is probably due to its ready transportation<br />
in the soil attached to the roots <strong>of</strong> cultivated plants. Although reported to<br />
attack C. sordidus eggs in Malaysia, China (1935) suggests that this species<br />
normally is unlikely to be a predator.<br />
Plaesius javanus Col.: Histeridae<br />
The predatory larvae and adults <strong>of</strong> this Indonesian beetle attack larvae and<br />
pupae <strong>of</strong> Cosmopolites sordidus and many other soil and litter-inhabiting<br />
insects. Eggs are laid singly in old banana stumps, at the base <strong>of</strong> the stem and<br />
on the rhizome below the soil surface. The eggs hatch in 8 to 9 days,<br />
producing active, voracious larvae which feed for 5 to 6 months, older larvae<br />
being capable <strong>of</strong> consuming up to 30 or more C. sordidus larvae per day. A<br />
prepupal period <strong>of</strong> 10 to 15 days is passed in a pupation cell constructed in<br />
the soil, followed by a pupal stage <strong>of</strong> about 14 days. The adult remains in the<br />
cell for 7 to 10 days before emerging and is then capable <strong>of</strong> consuming 7 or 8<br />
weevil larvae per day (Clausen 1978; Jepson 1914; Weddell 1932).<br />
Thyreocephalus interocularis Col.: Staphylinidae<br />
This is the second most effective predator on C. sordidus larvae in Kenya.<br />
Both adults and larvae are polyphagous. In the absence <strong>of</strong> other hosts adults<br />
may prey on their own larvae and larvae are also occasionally cannibalistic<br />
(Koppenhšfer and Schmutterer 1993). After a pre-oviposition period <strong>of</strong> 32<br />
days, females lay an average <strong>of</strong> 31 eggs in decomposing banana
104 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Comments<br />
pseudostems and in moist soil below banana mulch. Pupation occurs in the<br />
soil. Total development time averages 46 days and adults live an average <strong>of</strong><br />
142 days (Koppenhšfer 1994).<br />
Although weevils, as a group, seem to be poor candidates for biological<br />
control, the establishment <strong>of</strong> Plaesius javanus and P. laevigatus in Fiji<br />
appears to have reduced the pest status <strong>of</strong> Cosmopolites sordidus there.<br />
Introductions <strong>of</strong> P. javanus (Table 4.5.2) have resulted in successful<br />
establishment (but <strong>of</strong>ten not at the first attempt) in Cook Is, French<br />
Polynesia, Jamaica, Marianas, Mauritius, New Caledonia, Samoa and<br />
Trinidad, but no information is available on the effects it has produced.<br />
Introductions have been unsuccessful in Australia, Cameroon, Dominica,<br />
Honduras, Mauritius, Mexico, Puerto Rico, Seychelles, St Lucia, St Vincent,<br />
Taiwan, Tanzania, Tonga and Uganda (Bartlett 1937; Miwa 1938; Clausen<br />
1978). Based on his observation and that <strong>of</strong> others, Koppenhšfer (1993a,b,<br />
Koppenhšfer and Schmutterer 1993) considered that the biology <strong>of</strong><br />
P. javanus does not enable it to have any greater effect and that studies are<br />
necessary <strong>of</strong> the potential impact <strong>of</strong> proposed predator species before<br />
release. Two other predators have been established, one in Australia and one<br />
in Jamaica and St Vincent, but seemingly without much effect.<br />
If the beneficial effects <strong>of</strong> P. javanus (and P. laevigatus) in Fiji can be<br />
confirmed, it may be worth renewing efforts to establish them in other<br />
countries where C. sordidus is a major pest. Otherwise, resources available<br />
for biological control might be better deployed searching for, and<br />
evaluating, other natural enemies.<br />
It is possible that entomopathogenic nematodes (CSIRO 1993;<br />
Treverrow and Bedding 1993a) or fungi may hold some promise as<br />
biological pesticides. Nematodes generally have far less capability for selfperpetuation<br />
and dispersal in the environment than imported arthropod<br />
enemies, but they can be highly effective. However, in many tropical<br />
countries where bananas are a major staple food the distribution and<br />
application <strong>of</strong> mass-produced biological control agents, such as nematodes<br />
or fungi, is <strong>of</strong>ten impracticable because <strong>of</strong> storage and transport problems<br />
and lack <strong>of</strong> suitable equipment for application. In addition, like insecticides,<br />
they may be too costly for subsistence farmers who constitute the majority <strong>of</strong><br />
banana producers. Even if it only leads to a partial (but still significant)<br />
reduction in pest status, classical biological control is, under these<br />
circumstances, a particularly appropriate approach to reduce losses.
4.6 Deanolis sublimbalis<br />
India<br />
20°<br />
0°<br />
20°<br />
Myanmar<br />
Laos<br />
China<br />
++<br />
Thailand<br />
+<br />
Cambodia<br />
Malaysia<br />
Vietnam<br />
Singapore<br />
P<br />
Brunei<br />
P<br />
Indonesia<br />
+<br />
Taiwan<br />
++<br />
Philippines<br />
Australia<br />
+<br />
Papua<br />
New Guinea<br />
++<br />
105<br />
The red banded mango caterpillar, Deanolis sublimbalis tunnels in the flesh and seed <strong>of</strong> the<br />
fruit <strong>of</strong> mango, Mangifera indica,<br />
and also attacks the fruit <strong>of</strong> M. odorata,<br />
M. minor and<br />
Bouea burmanica.<br />
It is reported from India eastwards to <strong>Southeast</strong> Asia, southern China<br />
and Papua New Guinea. In this vast region there are scattered reports <strong>of</strong> damage ranging<br />
up to 50 per cent <strong>of</strong> fruit, but many areas within it from which there are no reports <strong>of</strong> damage<br />
or even <strong>of</strong> its presence.<br />
The only records <strong>of</strong> natural enemies are <strong>of</strong> two trichogrammatid egg parasitoids and a<br />
vespid larval predator, all in the Philippines. Further searches would be necessary before<br />
the potential <strong>of</strong> classical biological control could be evaluated, especially for infestation in<br />
new areas into which it has spread in recent years.<br />
20°<br />
0°<br />
20°
106 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Deanolis sublimbalis Snellen<br />
Synonyms<br />
Rating<br />
Origin<br />
Distribution<br />
Lepidoptera: Pyralidae: Odontinae<br />
red banded mango caterpillar, red banded borer<br />
Long known as Noorda albizonalis Hampson 1903 or Autocharis<br />
albizonalis (Hampson), this species should be referred to as Deanolis<br />
sublimbalis Snellen, because <strong>of</strong> the priority <strong>of</strong> its description (Snellen 1899)<br />
(M. Shaffer pers. comm. 1997) from specimens collected in Celebes (now<br />
Sulawesi). He also referred to two females from Batavia (now Jakarta).<br />
HampsonÕs type specimen (Hampson 1903), labelled Darjiling, is in the<br />
British Museum (Natural History) (BMNH); and the distribution <strong>of</strong> his<br />
species was given as Sikkim; Celebes, Palos B., Dongola (Doherty).<br />
Dongola is currently spelt Donggala and is at the southern headland <strong>of</strong> Palos<br />
bay at the head <strong>of</strong> which is Palu. DohertyÕs obituary (Hartent 1901) reveals<br />
that he collected there in August and September 1896.<br />
<strong>Southeast</strong> Asia China Pacific<br />
3 ++ Phil 2 ++ Yunnan Province 2 ++ PNG<br />
+ Thai<br />
P Brun, Indo<br />
The origin <strong>of</strong> mango ( Mangifera indica)<br />
is believed to be in the India-<br />
Myanmar region, from which it might be inferred that D. sublimbalis also<br />
evolved within this region, unless it has transferred to M. indica from a<br />
related plant.<br />
India, Myanmar, Thailand, China (Yunnan Province: Li et al. 1997), Brunei,<br />
Philippines, Indonesia (Java, Sulawesi, Irian Jaya), Papua New Guinea,<br />
Torres Strait (Dauan Is, Saibai Is: AQIS 1991; NAQS 1993) (Fenner 1987;<br />
Singh 1987).<br />
It is apparently not known in Pakistan (M.A. Poswal pers. comm. 1997),<br />
Sri Lanka (J. Edirisinghe pers. comm. 1997), Nepal (Neupane 1995) or<br />
Peninsular Malaysia (Yunus and Ho 1980; Tan Chai-Lin pers. comm. 1997)<br />
and does not occur on the Australian mainland or in the oceanic Pacific. It is
Biology<br />
4.6<br />
Deonalis sublimbalis<br />
107<br />
widely distributed throughout Papua New Guinea coastal mainland and<br />
islands (F. Dori pers. comm. 1997).<br />
Specimens in the BMNH carry the following labels<br />
India: Darjiling (now Darjeeling)<br />
Calcutta 22 March, 1945<br />
Orissa March, 1952<br />
Myanmar: Rangoon March, 1923<br />
Thailand and Philippines: no dates<br />
Brunei: 12 April, 1973, 5 September 1992<br />
Indonesia: Java August, 1922 (Koepoedan) Sulawesi<br />
(Minahassa, Tomohon).<br />
Irian Jaya July, 1936 (Cyclops Mts,<br />
Sabron 2000ft)<br />
Papua New Guinea: Kokoda August, 1933;<br />
and in the Australian National <strong>Insect</strong> Collection the following labels:<br />
Papua New Guinea: Finisterre Range 23 JuneÐ21 July, 1958<br />
(Gabumi, 2000ft)<br />
Telefomin 2 May and 18 June, 1959<br />
(Feramin 4700ft) 2 MayÐ18 June<br />
Port Moresby 5 MarchÐ12 May, 1963<br />
(Mt Lawes 1300ft)<br />
Referring presumably to the major (summer) crop <strong>of</strong> mangos, Fenner (1987)<br />
and Golez (1991a) comment that the eggs are oval, waxy white and<br />
generally laid in masses near the apex <strong>of</strong> the developing fruit. However<br />
F. Dori (pers. comm. 1997) reports that, in the winter (July) crop near Port<br />
Moresby (PNG), eggs were white to crimson. They were laid in groups <strong>of</strong> 1<br />
to 4 near or on the peduncle at the base <strong>of</strong> the fruit and sometimes covered by<br />
the sepals or deposited in small crevices in the fruit. On hatching, larvae<br />
travel to the apex to enter the fruit. Oviposition occurs as early as 45 to 55<br />
days after flower induction and continues up to fruit maturity. After an egg<br />
incubation period <strong>of</strong> 3 to 4 days, larvae hatch and pass through 5 instars in<br />
the next 14 to 20 days. The larva has a brown or black head and white body<br />
with red segmental bands. It feeds first in the pulp <strong>of</strong> the fruit (1st and 2nd<br />
instars) and later in the seeds. The tunnels formed gradually broaden as the<br />
larvae grow to about 2 cm in length. Fruit in all stages <strong>of</strong> development from<br />
marble size upwards are attacked. As many as 11 larvae may be found in a<br />
fruit, although there is commonly only one. A pre-pupal stage lasts about 2<br />
to 3 days and pupation occurs in a silk-lined earthern cocoon. In wooden
108 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Host plants<br />
cages in Papua New Guinea, larvae pupated in strongly spun cocoons<br />
covered with particles <strong>of</strong> chewed wood, suggesting that pupation on bark<br />
may occur in the field. In India, Sengupta and Behura (1955, 1957) record<br />
pupation generally inside the fruit, the moth emerging through an exit hole.<br />
The pupal period lasts from 9 to 14 days, so that the total period from egg to<br />
adult takes from 28 to 41 days. Adult longevity is 8 to 9 days. Adult males<br />
can be distinguished from females by having expanded dark brown, hairy,<br />
mesothoracic tibiae (Leefmans and Van der Vecht 1930; Vožte<br />
1936;<br />
Kalshoven 1981; Fenner 1987, 1997; Golez 1991a).<br />
Adults are generally nocturnal and, during the day, spend most <strong>of</strong> their<br />
time resting under leaves on the tree. They are seldom attracted to light.<br />
Females prefer to oviposit on fruit protected from direct light. Newly<br />
hatched larvae stay together and tunnel into the fruit near where the eggs<br />
were laid. If later instar larvae are crowded, some may leave by suspending<br />
themselves on silken threads, which also facilitate transfer to other fruits. A<br />
shorter developmental period was observed for both males and females<br />
reared on the pulp than on the seed <strong>of</strong> carabao mangoes, although those<br />
reared on the seeds were larger and lived longer, females producing more<br />
eggs. Development differed slightly on different mango varieties (Golez<br />
1991a).<br />
In cages in Papua New Guinea only a small proportion <strong>of</strong> pupae yielded<br />
adults in the several months after pupation, suggesting a pupal diapause<br />
which may synchronise the life cycle with the seasonal fruiting <strong>of</strong> its host<br />
(Fenner 1987, 1997).<br />
The commonest host, wherever D. sublimbalis occurs, is Mangifera indica,<br />
but there are records also from M. odorata in Papua New Guinea and<br />
Indonesia from M. minor in Papua New Guinea (F. Dori pers. comm. 1997).<br />
and from Bouea burmanica in Thailand (Beller and Bhenchitr 1936). All<br />
four belong to the family Anacardiaceae (Sengupta and Behura 1955;<br />
Kalshoven 1981; M. Schaffer pers. comm. 1997). Larvae are unable to<br />
develop on parts <strong>of</strong> the mango tree other than the fruit, or in the fruit <strong>of</strong><br />
avocado, chico, guava, jackfruit, papaya, santol, sineguelas or star apple<br />
(Golez 1991a). However, as the genus Mangifera contains many species it is<br />
quite possible that further wild hosts will be found (Fenner 1997). Indeed the<br />
label data, quoted earlier, on specimens collected well outside the major<br />
fruiting season <strong>of</strong> M. indica suggests that this may well be so. The genus<br />
Mangifera contains about 62 species <strong>of</strong> tall evergreen trees which are native<br />
to the area stretching from India to Papua New Guinea, with the greatest
Damage<br />
4.6<br />
Deonalis sublimbalis<br />
109<br />
number in the Malay Peninsula. Fifteen species bear edible fruit, but only<br />
M. indica is widely planted <strong>of</strong> the 6 species sometimes cultivated. M. indica<br />
probably originated in the Indo-Myanmar region and grows wild in the<br />
forests <strong>of</strong> India, particularly in hilly areas in the northeast. It has been grown<br />
throughout the Indian sub-continent for at least 4000 years. It was probably<br />
taken to Malaysia and eastwards further into <strong>Southeast</strong> Asia between 300<br />
and 400 AD and there are now many commercial varieties (Purseglove<br />
1968).<br />
Mango fruit in all stages <strong>of</strong> development are attacked, <strong>of</strong>ten leading to<br />
premature drop. First and second instar larvae feed on the tissues beneath the<br />
skin, making tunnels towards the seed. Larger larvae destroy the seed. Soon<br />
after boring starts, secondary infestations <strong>of</strong> bacteria, fungi, fruit flies (e.g.<br />
Bactrocera ferrugineus,<br />
B. frauenfeldi),<br />
and other pests occur. Liquid<br />
exudes from the skin <strong>of</strong> attacked fruit at the opening <strong>of</strong> the entry tunnel and<br />
trickles down to the drip point where it accumulates. It rapidly darkens to<br />
form a characteristic black spot, <strong>of</strong>ten about 1cm in diameter at the tip <strong>of</strong> the<br />
fruit (Fenner 1987). Another common sign <strong>of</strong> borer damage is the bursting <strong>of</strong><br />
the apex and longitudinal cracking <strong>of</strong> the fruit. In Guimaras Province<br />
(Philippines), up to 12.5% fruit infestation was recorded by Golez (1991a),<br />
with up to 14.5 larvae occurring per kg fruit. In years <strong>of</strong> serious infestation,<br />
yield could be reduced by as much as 40 to 50 per cent (Tipon 1979). In<br />
Papua New Guinea, fruit infestation levels <strong>of</strong> greater than 20% are<br />
encountered in the Port Moresby area (F. Dori pers. comm. 1997; T.L.<br />
Fenner pers. comm. 1997). In India the seeds are used as human food in<br />
times <strong>of</strong> famine and a flour is made from them (Purseglove 1968).<br />
However, for an insect that can be significantly damaging to mango fruit,<br />
it is remarkable that there are so very few references to it in the literature. It<br />
is, perhaps, instructive to list those that are directly relevant, so as to<br />
contribute to determining (i) whether it has, for many years, frequently been<br />
overlooked as a pest, (ii) whether it has spread to new areas in recent times,<br />
(iii) whether it has only become a pest <strong>of</strong> edible mangoes in recent times, (iv)<br />
whether suppression by natural enemies is no longer as effective as it once<br />
was and/or (v) whether there are other reasons.<br />
D. sublimbalis must have been present in northern India before it was<br />
described in 1903 (Hampson 1903), although it was not mentioned in the<br />
books by Maxwell-Lefroy and Howlett (1909), Fletcher (1914) or Ayyar<br />
(1963) all dealing with insects <strong>of</strong> agricultural importance in India. It was,<br />
however, reported a little later by Wadhi and Batra (1964) who referred to
110 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
papers by Sengupta and Behura (1955, 1957) and Sengupta and Misra<br />
(1956). It was also reported, briefly, by Nair (1975) and, in more detail, by<br />
Butani (1979). Strangely, the above Sengupta and Behura (1955) reference<br />
lists D. albizonalis among new records <strong>of</strong> crop pests in Orissa and then only<br />
<strong>of</strong> grafted mangoes in Puri District, implying that it was not known much<br />
earlier there as a damaging species. Furthermore, only recently<br />
(Zaheruddeen and Sujatha 1993) was D. sublimbalis recorded as having<br />
caused serious losses to mango fruits from marble size to maturity in<br />
Godavari Districts <strong>of</strong> Andhra Pradesh.<br />
Although a specimen was collected in Rangoon in 1923 (BMNH),<br />
D. sublimbalis was not listed by Ghosh (1940) in his major work Ô<strong>Insect</strong><br />
<strong>Pests</strong> <strong>of</strong> BurmaÕ or by Yunus and Ho (1980) in Malaysia when dealing with<br />
economic pests from 1920 to 1978. This striking absence <strong>of</strong> records from<br />
peninsular Malaysia continues to this day (Tan Chai-lin pers. comm. 1997).<br />
Nevertheless, D. sublimbalis has been well known in Thailand since 1936<br />
(Beller and Bhenchitr 1936; Cantelo and Pholboon 1965; Wongsiri 1991;<br />
Kuroko and Lewvanich 1993).<br />
In the Philippines it was not recorded by Cendana et al. (1984) in Ô<strong>Insect</strong><br />
<strong>Pests</strong> <strong>of</strong> Fruit Plants in the PhilippinesÕ, so it was evidently not generally<br />
regarded as a pest at that time, although a paper recording 40 to 50% damage<br />
in bad years had been delivered 5 years earlier (Tipon 1979). A<br />
comprehensive account <strong>of</strong> up to 12.5% infestation <strong>of</strong> fruit in Guimaras<br />
Province was published in 1991 by Golez (1991a,b).<br />
In contrast, in Indonesia it was present prior to 1899 (Snellen 1899) and<br />
has been well known as a mango pest since 1930 (Leefmans and van der<br />
Vecht 1930). Its damaging presence there is also documented by Vote<br />
(1936) and Kalshoven (1981).<br />
D. sublimbalis has been known in Irian Jaya since 1936 (BMNH<br />
specimen) and was common in mangoes in Jayapura in the early nineties<br />
(T.L. Fenner pers. comm. 1997). It was collected in Papua New Guinea<br />
(Kokoda) in 1933 (BMNH specimen) and was recorded again in 1958, 1959<br />
and 1963 (ANIC specimens) and is common nowadays in Port Moresby. It<br />
was first recorded on Australian islands in Torres Strait (Saibai I) in 1990<br />
and again in October 1996 (at a level <strong>of</strong> about 1% infestation on Dauan I)<br />
(Australian Quarantine Inspection Service).
Natural enemies<br />
Table 4.6.1<br />
4.6<br />
Deonalis sublimbalis<br />
111<br />
Leefmans and van der Vecht (1930) commented that no parasites had been<br />
bred in their studies on D. albizonalis in Java.<br />
In Luzon (Philippines) the egg parasitoids Trichogramma chilonis and<br />
T. chilotraeae (Table 4.6.1) were recorded by Golez (1991a) who reported,<br />
however, that no parasitoids were encountered at that time in the three<br />
municipalities <strong>of</strong> Guimaras, all <strong>of</strong> which had dry, dusty and windy<br />
conditions.<br />
Golez (1991a) reported that predation in Guimaras occurs as larvae leave<br />
the fruit, either to migrate to another fruit or to pupate in the soil. The most<br />
important predator was the vespid Rhychium attrisium which appeared to be<br />
the main cause <strong>of</strong> the high larval disappearance that occurs. R. attrisium is<br />
abundant in summer, especially during warm sunny days.<br />
Larvae were attacked by a fungus in the laboratory in Indonesia<br />
(Leefmans and van der Vecht 1930). The wasp Evania appendigaster is<br />
reported as a larval/pupal parasite (Golez 1991b), but Fenner (1997) points<br />
out that this record needs confirmation since the Evaniidae are reportedly all<br />
parasites <strong>of</strong> cockroach eggs.<br />
A tachinid, Carcelia ( Senometopia)<br />
sp., was reared from mango fruit<br />
possibly infested with D. sublimbalis near Nodup in September 1982<br />
(J. Ismay pers. comm. 1997) and also from a D. sublimbalis larva near<br />
Rabaul (both Papua New Guinea) in 1984 (F. Dori pers. comm. 1997).<br />
Natural enemies <strong>of</strong> Deanolis sublimbalis<br />
Species<br />
DIPTERA<br />
TACHINIDAE<br />
Location Reference<br />
Carcelia sp.<br />
HYMENOPTERA<br />
TRICHOGRAMMATIDAE<br />
Rabaul (PNG) F. Dori pers. comm. 1997<br />
Trichogramma chilonis<br />
Philippines Golez 1991a<br />
Trichogramma chilotraeae<br />
EVANIIDAE<br />
Philippines Golez 1991a<br />
Evania appendigaster<br />
VESPIDAE<br />
Golez 1991b<br />
Rhychium attrisium<br />
Philippines Golez 1991a
112 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Comment<br />
It is tempting to postulate that the damage that is actually due to the red<br />
banded mango caterpillar is commonly attributed to other causes. Perhaps<br />
this is due, in part, to the fact that larvae have <strong>of</strong>ten left the fruit before the<br />
cause <strong>of</strong> damage is investigated although, with a larval duration <strong>of</strong> 2 to 3<br />
weeks, this would be surprising, particularly when there is a characteristic<br />
dark spot for much <strong>of</strong> this time at the drip point <strong>of</strong> the mango fruit.<br />
The absence <strong>of</strong> records <strong>of</strong> its presence over vast areas within its<br />
distribution range suggests that its abundance must be very low (perhaps due<br />
to inhospitable host varieties or effective biological control) or, perhaps, that<br />
it does not occur there.<br />
The only record <strong>of</strong> effective chemical control is <strong>of</strong> 4 applications <strong>of</strong><br />
cyfluthrin or deltamethrin at 60, 75, 90 and 105 days after fruit induction<br />
(Golez 1991a).<br />
Further research for natural enemies attacking eggs, larvae and pupae<br />
within its long established range would be necessary to determine whether<br />
any are likely to be promising for biological control.<br />
If, as is very probable, D. sublimbalis produces a sex pheromone, its<br />
availability as a lure would be <strong>of</strong> great value as a means <strong>of</strong> monitoring the<br />
presence and distribution <strong>of</strong> the red banded mango caterpillar in mango and<br />
other hosts. Its identification, synthesis and availability should have high<br />
priority.
4.7 Diaphorina citri<br />
India<br />
20°<br />
Myanmar<br />
P Laos<br />
0°<br />
20°<br />
China<br />
++<br />
Thailand<br />
P<br />
Cambodia<br />
Vietnam<br />
++<br />
+ Brunei<br />
Malaysia<br />
+<br />
Singapore<br />
++<br />
Indonesia<br />
Taiwan<br />
+++<br />
++<br />
Philippines<br />
Australia<br />
+<br />
Papua<br />
New Guinea<br />
113<br />
The citrus psyllid Diaphorina citri is native to the Indo-Malaysian region, but has spread<br />
outside it to RŽunion, Mauritius, Saudi Arabia, Honduras, and Brazil. The sap it removes<br />
from new flushes <strong>of</strong> citrus growth is <strong>of</strong> minor consequence, but it is the vector <strong>of</strong> a<br />
devastating bacterial disease, citrus greening.<br />
Its major controlling factors are high rainfall (washing <strong>of</strong>f eggs and young nymphs) and<br />
two parasitoids, Tamarixia radiata and Diaphorencyrtus aligarhensis.<br />
Where these<br />
parasitoids are native they are very heavily attacked by hyperparasitoids which diminish<br />
their effectiveness. Freed <strong>of</strong> these hyperparasitoids T. radiata has been established in 3<br />
countries where it was not present: RŽunion, Madagascar and Taiwan, resulting in<br />
excellent biological control. Although T. radiata appears to be widespread in <strong>Southeast</strong><br />
Asia, observations might well disclose regions where it is not present and could be<br />
introduced with advantage. The prospects for successful biological control <strong>of</strong> D. citri are<br />
good when it invades regions where hyperparasitoids <strong>of</strong> T. radiata are absent or deficient.<br />
20°<br />
0°<br />
20°
114 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Diaphorina citri Kuwayama<br />
Rating<br />
Origin<br />
Distribution<br />
Hemiptera, Psyllidae<br />
citrus psyllid, <strong>Asian</strong> citrus psyllid<br />
<strong>Southeast</strong> Asia China Southern and Western Pacific<br />
8 ++ Viet, Indo, Phil 3+++ absent<br />
+ Msia, Sing<br />
P Myan, Thai<br />
These <strong>Southeast</strong> <strong>Asian</strong> ratings arose from an earlier survey <strong>of</strong> country<br />
opinions (Waterhouse 1993b) and may not reflect current assessments.<br />
The Indo-Malaysian region. D. citri was described from Punjab, India<br />
(Waterston 1922). There is evidence <strong>of</strong> recent spread into the southeastern<br />
and eastern portions <strong>of</strong> <strong>Southeast</strong> Asia.<br />
D. citri is widespread from Afganistan eastwards through Pakistan, India,<br />
Nepal and Bhutan to <strong>Southeast</strong> Asia, southern China (up to about 30¡N, Xie<br />
et al. 1988), Taiwan (Catling 1970; Tsai et al. 1984; Aubert 1990) and the<br />
Ryuku Is (Japan) (Miyakawa and Tsuno 1989). It has recently become<br />
established in Ende (Flores) and Timor and in Irian Jaya (Aubert 1989b,<br />
1990). D. citri was collected in June 1993 in the Jayapura area <strong>of</strong> Irian Jaya<br />
and citrus there showed symptoms <strong>of</strong> greening (Northern Australia<br />
Quarantine Strategy 1993). It has been introduced to RŽunion, Mauritius,<br />
Comoro Is (Hollis 1987), Saudi Arabia (Wooler et al. 1974), Yemen (BovŽ<br />
1986), Brazil (Silva et al. 1968; Bergmann et al. 1994) and Honduras<br />
(Burckhardt and Martinez 1989). In 1990 there were still limited areas free<br />
<strong>of</strong> D. citri in east Mindoro (Philippines) and Palau and Tambun (Malaysia).<br />
It is not yet recorded from Papua New Guinea and does not occur in<br />
Australia, the Oceanic Pacific or North America.<br />
In RŽunion it has not colonised citrus plantings above 800 m, where the<br />
lowest temperature is 7¡C, whereas in Malaysia the height limit is 1200 m<br />
with a minimum temperature <strong>of</strong> 14¡C.
Biology<br />
Table 4.7.1<br />
4.7<br />
Diaphorina citri<br />
115<br />
D. citri survives a wide range <strong>of</strong> temperature extremes from 45¡C in Saudi<br />
Arabia to Ð7¡ to Ð8¡C in China, thereby tolerating cold that will kill citrus<br />
(Xie et al. 1989a). Far more than temperature, high humidity and rainfall are<br />
important limiting factors, rain by washing <strong>of</strong>f eggs and early instar nymphs<br />
and humidity by favouring fungal attack. These two factors are mainly<br />
responsible for the low D. citri populations on the windward (rainy) side <strong>of</strong><br />
Mindoro (Philippines) and RŽunion (Aubert 1989a).<br />
There have been several studies on the life cycle <strong>of</strong> D. citri,<br />
which<br />
conform generally with the results in Table 4.7.1, leading to up to 11<br />
generations a year in Fujian Province, China (Xu et al. 1988b, 1994). D. citri<br />
has a short life cycle and high fecundity and is commonest in hot coastal<br />
areas. Mating commences soon after the insects become adult and, after a<br />
pre-oviposition period <strong>of</strong> about 12 days, eggs are laid singly inside halffolded<br />
leaves <strong>of</strong> buds, in leaf axils and other places on the young tender<br />
shoots. Average adult lifespan is 30 to 40 days, although overwintering<br />
adults had a lifespan <strong>of</strong> 260 days (Xu et al. 1994).<br />
Bionomics <strong>of</strong> D. citri (average in days) in Fujian Province,<br />
China (Xu et al. 1988b)<br />
Adult life-span<br />
Max Min<br />
Eggs per<br />
female<br />
Incubation Nymphal<br />
development<br />
EggÐadult<br />
Spring 96 28.1 17.7 10 31.8 42<br />
Summer 46 19.7 43.8 2 10.3 13<br />
Autumn 59 31.6 22.6 4 16.8 21<br />
Winter 131Ð165<br />
The abundance <strong>of</strong> both eggs and nymphs is correlated with the availability <strong>of</strong><br />
new growth flushes and breeding is largely suspended when trees become<br />
dormant. On its favoured host plant Murraya paniculata in Fujian,<br />
populations may average 51 adults per young shoot and a 4-year-old plant<br />
produces 900Ð1000 shoots. On mandarin ( Citrus reticulata)<br />
the average<br />
colony size is 20 per shoot, with 600 to 650 shoots, and peak abundance<br />
occurs about 6 weeks later than on M. paniculata (Aubert 1990). D. citri<br />
nymphs develop well under cool, humid spring conditions, but are seriously<br />
affected by fungal infections under hot, humid conditions. On<br />
M. paniculata,<br />
adult numbers were highest on leaf midveins (43%),<br />
followed by petioles (30.7%), leaf blades (23.7%) and stems (2.6%) (Tsai et<br />
al. 1984).
116 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Host plants<br />
Damage<br />
D. citri nymphs normally lead a sedentary existence clustered in groups,<br />
but will move away when disturbed. Adults are 2.5 mm long and jump when<br />
disturbed, whereupon they may fly up to 5 m before settling again. Seasonal<br />
migratory flights occur when adults fly up to about 7 m above ground level,<br />
entering mild winds which may carry them up to 4 km distant (Aubert 1990).<br />
Flying adults are attracted to yellow traps, which have been used for<br />
sampling (Aubert and Xie 1990). Adult D. citri have yellowish-brown<br />
bodies, greyish-brown legs and transparent wings. They have white spots or<br />
are light brown with a broad, beige, longitudinal, central band.<br />
D. citri feeds and breeds on the entire group <strong>of</strong> horticultural Citrus,<br />
with<br />
additional hosts in eight different genera belonging to the Aurantoidea<br />
(Aubert 1990). D. citri thus has a wider host range than the greening<br />
organism it transmits to citrus (see Damage). An indication <strong>of</strong> the relative<br />
suitability <strong>of</strong> its various host plants is shown in Table 4.7.2, although there<br />
may be local modifications <strong>of</strong> the groupings. This is probably due to<br />
different D. citri biotypes. For example, unlike RŽunion populations,<br />
Malaysian populations breed well on Bergera koenigii and, in the<br />
Philippines, adults are more attracted by Clausena anisumolens than by<br />
Murraya paniculata (Aubert 1990). Overall, jasmin orange, Murraya<br />
paniculata, is the preferred host and this plant is widely grown in Southern<br />
and <strong>Southeast</strong> Asia as an ornamental shrub and hedge plant.<br />
Although sap removal by large populations <strong>of</strong> D. citri can cause young<br />
foliage on flushes <strong>of</strong> growth to wilt, by far the most damaging effect <strong>of</strong><br />
feeding is due to the transmission <strong>of</strong> a gram-negative bacterium which is the<br />
cause <strong>of</strong> citrus greening, known as huanglungbin in China (Xu et al. 1988a).<br />
Citrus greening is known to affect 3 genera <strong>of</strong> the subtribe Citrinae, namely<br />
Citrus,<br />
Poncirus and Fortunella (Aubert 1990). It has also been<br />
experimentally transferred from Citrus to Madagascar periwinkle<br />
( Catharanthus roseus (Ke 1987). Once infected with the bacterium, D. citri<br />
remains infective for its lifetime, but does not pass on the infection<br />
transovarially. Amongst citrus, pummelo and lemon are less affected by<br />
greening than most other species. D. citri is the only known vector <strong>of</strong> citrus<br />
greening in Asia, although several other psyllids attacking citrus have been<br />
described: D. auberti (Comoro Is: Hollis 1987), Psylla citricola,<br />
P. citrisuga and Trioza citroimpura (China: Yang and Li 1984) and Psylla<br />
murrayii (Malaysia: Osman and Lim 1990).
Table 4.7.2<br />
Preferred host plant<br />
Good host plants<br />
Common host plants<br />
Occasional host plants<br />
Diaphorina citri host plants (after Aubert 1990)<br />
Murraya paniculata ( jasmin orange)<br />
Citrus aurantifolia (lime)<br />
Bergera (Murraya) koenigii (curry bush)<br />
Leaf sucking Egg laying Nymphal<br />
development<br />
+++ +++ +++<br />
+++ +++ +++<br />
Citrus limon (lemon)<br />
Citrus sinensis (sweet orange)<br />
Citrus medica (citron)<br />
Citrus reticulata (mandarin)<br />
Microcitrus australisiaca*<br />
Citrus maxima var. racemosa (pummelo)<br />
Citrus hystrix ( Mauritius papeda)<br />
Citrus madurensis<br />
Clausena excavata<br />
Clausena lansium<br />
++ ++ ++<br />
Citrus maxima (pummelo) + + +<br />
Triphasia trifoliata*<br />
+ + +<br />
Fortunella sp.* (kumquat) + + +<br />
Poncirus trifoliata*<br />
+ + Ð<br />
Clausena anisumolens (anise) + + +<br />
Merrillia caloxylon*<br />
+ Ð Ð<br />
Toddalia asiatica*<br />
+ Ð Ð<br />
4.7<br />
Diaphorina citri<br />
117
Table 4.7.2 (contÕd) Diaphorina citri host plants (after Aubert 1990)<br />
Occasional host plants<br />
Leaf sucking Egg laying Nymphal<br />
development<br />
Vepris lanceolata*<br />
+ Ð Ð<br />
Swinglea glutinesa*<br />
+ unknown unknown<br />
Atalantia sp. + unknown unknown<br />
Clausena indica*<br />
+ unknown unknown<br />
Murraya exotica*<br />
+ unknown unknown<br />
Citrus species hosts are, according to the classification <strong>of</strong> Jones (1990):<br />
+++ very common;<br />
++ usual<br />
+ occasional;<br />
Ð complete life cycle not observed<br />
*observations on caged insects<br />
118 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong>
4.7 Diaphorina citri 119<br />
In Africa, RŽunion, Madagascar and Saudi Arabia another psyllid Trioza<br />
erytreae transmits a slightly different citrus greening organism (see later<br />
under RŽunion).<br />
Citrus greening is believed to have originated in northeastern<br />
Guangdong Province (Lin and Lin 1990). Amongst other symptoms, the<br />
leaves <strong>of</strong> new green shoots first turn yellow at their base, then <strong>of</strong>ten become<br />
mottled yellow and drop. The branches remain small, upright and stiff.<br />
Diseased trees flower abundantly in the <strong>of</strong>f-season and flowers drop readily<br />
or result in small, irregular fruit whose base turns red before the remainder<br />
changes from green (Ke 1987). Citrus greening is widespread throughout<br />
South and <strong>Southeast</strong> Asia, where it is almost always the most serious disease<br />
<strong>of</strong> citrus. It is spread to new areas by infected nursery plants or infected<br />
budwood and within orchards by D. citri (Capoor et al. 1967; Whittle 1992).<br />
However, D. citri has been intercepted by quarantine in France on citrus<br />
imported from Honduras (Burckhardt and Martinez 1989). The tonnage <strong>of</strong><br />
citrus produced worldwide is second as a fruit crop only to that <strong>of</strong> grapes<br />
(Aubert 1987b). An extremely serious citrus disease which already affects<br />
nearly 50 countries in Asia and Africa must, therefore, be regarded as <strong>of</strong><br />
major importance. It is reported that a total <strong>of</strong> over a million trees are<br />
destroyed each year in China, Thailand, Malaysia, Indonesia and Philippines<br />
alone (Aubert 1987a). In Indonesia citrus greening has caused the loss <strong>of</strong><br />
many millions <strong>of</strong> trees. Small farmers are frequently reluctant to remove<br />
declining trees before they almost cease bearing. This tends to increase<br />
D. citri populations, which breed on young flush since a symptom <strong>of</strong><br />
greening is unseasonal flushing (Whittle 1992). The recent history <strong>of</strong><br />
production in northern Vietnam, where citrus is grown mainly in larger<br />
orchards or state farms, is typically cyclical, with the gradual destruction <strong>of</strong><br />
trees by greening and then wholesale removal and replanting. A new cycle <strong>of</strong><br />
planting commenced in the late 1980s, but greening is already to be seen in<br />
many young orchards, although populations <strong>of</strong> D. citri are still low (Whittle<br />
1992). Only by keeping populations at very low levels by biological control<br />
and/or insecticides will the rate <strong>of</strong> spread <strong>of</strong> greening be diminished.<br />
<strong>Insect</strong>icides are said to be highly cost effective if used only during a<br />
restricted flushing period, but if needed frequently they are very costly and<br />
environmentally undesirable. Recent developments with carefully specified,<br />
highly refined petroleum oils has given high levels <strong>of</strong> control <strong>of</strong> D. citri<br />
(A. Beattie pers. comm. 1995), with presumably little direct effect on its<br />
parasitoids. Whittle (1992) reported that he was unable to find D. citri in the<br />
vicinity <strong>of</strong> Ho Chi Minh City (southern Vietnam), a very unusual situation<br />
for an area with a fairly long history <strong>of</strong> citrus cultivation.
120 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
The <strong>Asian</strong> citrus greening bacterium can withstand high temperatures<br />
and occurs in China, <strong>Southeast</strong> Asia, India and Saudi Arabia. On the other<br />
hand, Southern African greening, which is transmitted by the psyllid Trioza<br />
erytreae, is heat-sensitive and symptoms do not develop in climates where<br />
temperatures above 30¡C are recorded for several hours a day. In addition to<br />
Southern Africa, this greening occurs also in North Yemen (Garnier et al.<br />
1988).<br />
Natural enemies<br />
Identified natural enemies are listed in Table 4.7.3. There are also reports <strong>of</strong><br />
a number <strong>of</strong> unidentified predators (coccinellids, chrysopids, mantids,<br />
spiders). It is noteworthy that only 2 primary parasitoidsÑboth attacking<br />
D. citri nymphsÑhave so far been recorded, the widespread endoparasitic<br />
encyrtid Diaphorencyrtus aligarhensis and the more restricted ectoparasitic<br />
eulophid Tamarixia radiata, which has been introduced to several countries<br />
for biological control. Both feed on the haemolymph <strong>of</strong> some hosts, resulting<br />
in their death, as well as using other hosts for oviposition.<br />
Where they occur naturally, both D. aligarhensis and T. radiata are<br />
heavily attacked by a wide range <strong>of</strong> hyperparasitoids (Table 4.7.4). Of these,<br />
Tetrastichus sp. is the most important for T. radiata, causing an average <strong>of</strong><br />
21.8% parasitisation in 1988 (rising to a maximum <strong>of</strong> 87.9%) and 28.7% in<br />
1989 in Fujian Province, China. Chartocerus walkeri (9.3% in 1988 and<br />
13.2% in 1989) is the most important for D. aligarhensis (Table 4.7.5).<br />
A valuable illustrated guide to the hyperparasitoids associated with<br />
D. citri is provided by Qing and Aubert (1990).
Table 4.7.3 Natural enemies <strong>of</strong> Diaphorina citri (* indicates introduced to this country)<br />
Species Region Reference<br />
HYMENOPTERA<br />
ENCYRTIDAE<br />
Diaphorencyrtus aligarhensis<br />
(= Aphidencyrtus diaphorinae<br />
= Diaphorencyrtus diaphorinae<br />
= Psyllaephagus diaphorinae<br />
= Aphidencyrtus aligarhensis)<br />
EULOPHIDAE<br />
Tamarixia radiata<br />
(= Tetrastichus radiatus)<br />
India<br />
Vietnam<br />
Taiwan<br />
Shafee et al. 1975; Hayat 1981<br />
Myartseva & Tryapitzyn 1978;<br />
van Lam 1996<br />
Prinsloo 1985<br />
Lin & Tao 1979<br />
Comores Is Aubert 1984b<br />
RŽunion Aubert & Quilici 1984, Quilici 1989<br />
Philippines Prinsloo 1985; Gavarra & Mercado 1989;<br />
Gavarra et al. 1990<br />
China Tang 1989<br />
Indonesia<br />
Malaysia<br />
India<br />
RŽunion*<br />
Nurhadi 1989; Nurhadi & Crih 1987<br />
Lim et al. 1990<br />
Waterston 1922; Husain & Nath 1924; Quilici 1989,<br />
Etienne and Aubert 1980<br />
Saudi Arabia Aubert 1984a<br />
Mauritius* Aubert 1984c<br />
Nepal Lama et al. 1988; Otake 1990<br />
Taiwan* Chiu et al. 1988<br />
China Liu 1989; Tang 1989; Qing & Aubert 1990<br />
Indonesia<br />
Malaysia<br />
Nurhadi & Crih 1987; Nurhadi 1989<br />
Lim et al. 1990<br />
Thailand Qing & Aubert 1990<br />
Vietnam Myartseva & Trijapitzyin 1978;<br />
van Lam 1996<br />
4.7 Diaphorina citri 121
Table 4.7.3 (contÕd) Natural enemies <strong>of</strong> Diaphorina citri (* indicates introduced to this country)<br />
Species<br />
COLEOPTERA<br />
Region Reference<br />
COCCINELLIDAE<br />
Cheilomenes sexmaculata China Xia et al. 1987<br />
NEUROPTERA<br />
CHRYSOPIDAE<br />
Chrysopa boninensis China Liu 1989<br />
ARACHNIDA<br />
SALTICIDAE<br />
Marpissa tigrina India Sanda 1991<br />
FUNGI<br />
Beauveria bassiana China Chen et al. 1990<br />
Cephalosporium (= Verticillium) lecanii China Xie et al. 1988<br />
Fusarium lateritium China Xie et al. 1988<br />
Paecilomyces sp. China Xie et al. 1988<br />
122 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong>
Table 4.7.4 Hyperparasitoids <strong>of</strong> Diaphorina citri (mostly after Tang 1989)<br />
Hyperparasitoid<br />
EULOPHIDAE<br />
Attacks Region Reference<br />
Tetrastichus sp. T.r. & D.a. China Tang 1989<br />
D.a. Taiwan Hayat & Lin 1988; Chien et al. 1989<br />
ENCYRTIDAE<br />
Philippines Balthazar 1966, unpublished<br />
Syrphophagus taiwanus T.r. & D.a. Taiwan Hayat & Lin 1988; Chien et al. 1989<br />
T.r. & D.a. China Tang 1989<br />
Ageniaspis sp. D.a. Taiwan Hayat & Lin 1988; Chien et al. 1989<br />
Cheiloneurus sp.<br />
D.a.<br />
Taiwan Hayat & Lin 1988; Chien et al. 1989,<br />
?<br />
Philippines Baltazar 1966, unpublished<br />
?Psyllaephagus sp.<br />
T.r. & D.a. China<br />
Tang 1989<br />
Philippines Balthazar 1966, unpublished<br />
Tang 1989<br />
Several unidentified<br />
SIGNIPHORIDAE<br />
D.a. China Tang 1989<br />
Chartocerus walkeri T.r. & D.a. Taiwan Hayat & Lin 1988; Chien et al. 1989<br />
T.r. & D.a. China<br />
Tang 1989<br />
Signiphora sp.<br />
PTEROMALIDAE<br />
D.a. Gavarra et al. 1990<br />
Pachyneuron concolor<br />
APHELINIDAE<br />
T.r. & D.a. Taiwan Hayat & Lin 1988; Chien et al. 1989<br />
Coccophagus ceroplastae D.a. Taiwan Hayat & Lin 1988; Chien et al. 1989<br />
Coccophagus sp. D.a. Taiwan Hayat & Lin 1988; Chien et al. 1989<br />
4.7 Diaphorina citri 123
Table 4.7.4 (contÕd) Hyperparasitoids <strong>of</strong> Diaphorina citri (mostly after Tang 1989)<br />
Hyperparasitoid Attacks Region Reference<br />
APHELINIDAE (contÕd)<br />
Marietta leopardina<br />
(= Marietta javensis)<br />
T.r. = Tamarixia radiata D.a. = Diaphorencyrtus aligarhensis<br />
T.r. & D.a.<br />
D.a.<br />
Taiwan<br />
Philippines<br />
Encarsia spp. T.r. & D.a. Taiwan<br />
China<br />
Unidentified sp. T.r. & D.a. Taiwan Chien et al. 1989<br />
Hayat & Lin 1988; Chien et al. 1989<br />
Balthazar 1966, unpublished<br />
Hayat & Lin 1988; Chien et al. 1989<br />
Tang 1989<br />
124 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong>
Table 4.7.5 Hyperparsitoids <strong>of</strong> Tamarixia radiata and Diaphorencyrtus aligarhensis in Fujian and Taiwan (after Qing<br />
1990)<br />
T. radiata<br />
Percentage <strong>of</strong> hyperparasitisation<br />
D. aligarhensis<br />
Hyperparasitoid<br />
EULOPHIDAE<br />
Fujian Fujian Taiwan Fujian Fujian Taiwan<br />
Tetrastichus sp.<br />
PTEROMALIDAE<br />
21.82 28.65 0.01 2.90 3.68<br />
Pachyneuron concolor<br />
SIGNIPHORIDAE<br />
0.45 18.50<br />
Chartocerus walkeri<br />
ENCYRTIDAE<br />
0.08 1.09 0.03 9.26 13.16 13.50<br />
Syrphophagus taiwanus 0.05 1.09 4.21 6.80<br />
?Psyllaephagus sp. 0.04 0.10 10.35 6.58<br />
Cheiloneurus sp.<br />
Ageniaspis sp.<br />
0.01<br />
unidentified sp.A<br />
Sp.B<br />
Sp.C<br />
Sp.D<br />
APHELINIDAE<br />
Encarsia sp. near transvena<br />
(= E. shafeei)<br />
3.45<br />
0.91<br />
0.18<br />
0.18<br />
0.26<br />
0.11 0.80<br />
Encarsia sp. A 0.08 0.10 0.91 1.05<br />
Encarsia sp. B 0.22 0.20 0.91<br />
4.7 Diaphorina citri 125
Table 4.7.5 (contÕd) Hyperparsitoids <strong>of</strong> Tamarixia radiata and Diaphorencyrtus aligarhensis in Fujian and Taiwan (after Qing<br />
1990)<br />
Percentage <strong>of</strong> hyperparasitisation<br />
T. radiata D. aligarhensis<br />
Hyperparasitoid Fujian Fujian Taiwan Fujian Fujian Taiwan<br />
APHELINIDAE (contÕd)<br />
Marietta leopardina 0.25 2.50<br />
Coccophagus ceroplastae 0.01<br />
Coccophagus sp. 0.10<br />
Unidentified sp. 0.05 0.01<br />
Totals 22.24 30.14 0.90 30.14 28.94 39.72<br />
126 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong>
4.7 Diaphorina citri 127<br />
It is noteworthy that T. radiata, which has fairly recently (1984Ð1988)<br />
been introduced into Taiwan, was hyperparasitised to the extent only <strong>of</strong><br />
0.95% in 1989, whereas 42.2% <strong>of</strong> the native D. aligarhensis was attacked<br />
(Qing 1990).<br />
The levels <strong>of</strong> hyperparasitisation <strong>of</strong> both primary parasitoids seriously<br />
affects their capacity to develop high populations and hence to produce<br />
maximum reduction <strong>of</strong> host populations. Nevertheless, each primary<br />
parasitoid killed is also a D. citri killed, so the overall mortality <strong>of</strong> D. citri is<br />
the sum <strong>of</strong> the mortalities produced by both primary parasitoids and their<br />
hyperparasitoids. It is abundantly clear that all hyperparasitoids must be<br />
rigorously excluded when transferring primary parasitoids from one region<br />
to another.<br />
Attempts at biological control<br />
CHINA<br />
The parasitoid Tamarixia radiata, obtained originally from India, has been<br />
used in successful biological control projects in RŽunion, Mauritius and<br />
Taiwan and in an attempt in the Philippines (Table 4.7.6). These projects and<br />
comments on the situation in several other countries follow.<br />
Table 4.7.6 Introductions for the biological control <strong>of</strong> Diaphorina citri<br />
Species<br />
EULOPHIDAE<br />
From To Year Result Reference<br />
Tamarixia radiata India RŽunion 1978 + Aubert & Quilici 1984;<br />
Quilici 1989<br />
RŽunion Mauritius after 1978 + Quilici 1989<br />
RŽunion Taiwan 1983Ð86 + Chiu et al. 1988;<br />
Chien et al. 1988<br />
RŽunion Philippines 1989 +<br />
?<br />
Gavarra et al. 1990<br />
Mercado et al. 1991<br />
In Guangdong, predators (lacewings, ladybird beetles, thrips, spiders)<br />
caused about 80% mortality <strong>of</strong> D. citri. Duration <strong>of</strong> daylight (short days<br />
reducing oviposition), quality <strong>of</strong> the flushes, and pesticide usage were other<br />
important factors influencing D. citri populations (Chen 1988). It appears<br />
that some Chinese farmers may spray citrus up to 50 times a year.<br />
In Fujian there are 8 generations a year <strong>of</strong> D. citri on jasmin orange,<br />
Murraya paniculata and populations reach their peak in summer and early<br />
autumn during hot, dry weather when fresh shoots appear regularly.<br />
Populations are lowest in cold, wet weather with average temperatures <strong>of</strong>
128 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
9.1¡ to 12.2¡C. Rainfall affects populations since eggs are laid on very young<br />
twigs and are easily washed <strong>of</strong>f. A Tamarixia sp. was recorded in September<br />
1987 and caused 83.3% parasitisation <strong>of</strong> nymphs in late autumn. In spring<br />
1988 its population was low, but D. citri mainly overwinters as the adult and<br />
Tamarixia only attacks nymphs. Predators included coccinellids (especially<br />
Cheilomenes sexmaculata and Harmonia axyridis), lacewings, spiders and<br />
praying mantids (Xia et al. 1987; Ke 1991).<br />
In Guangdong a maximum <strong>of</strong> 75% mortality <strong>of</strong> D. citri was recorded as<br />
being due to the hyperparasitoid Tetrastichus sp. (Liu 1989).<br />
Beauveria bassiana (Chen et al. 1990a), Cephalosporium (Verticillium)<br />
lecanii and two other fungi (Fusarium lateritium and Paecilomyces sp.)<br />
were found attacking D. citri. Suspensions <strong>of</strong> C. lecanii sprayed on to<br />
D. citri displayed a very high pathogenicity (Xie et al. 1988, 1989b).<br />
INDONESIA<br />
Citrus greening is also known as citrus vein phloem degeneration. In East<br />
Java, both T. radiata (the commoner) and D. aligarhensis (the more<br />
widespread) were found in 1987 attacking D. citri on Murraya paniculata<br />
(Nurhadi and Crih 1987). D. citri is known to occur in Irian Jaya and may<br />
have been introduced in recent times, but it is not known if it is parasitised<br />
there.<br />
MALAYSIA<br />
Both Tamarixia radiata and Diaphorencyrtus aligarhensis are present with<br />
parasitisation rates ranging up to 28% in 4th and 5th instar nymphs (Osman<br />
and Quilici 1991) or up to 36% parasitisation (Lim et al. 1990). T. radiata is<br />
also present in Sarawak (S. Leong, pers. comm. 1995).<br />
NEPAL AND BHUTAN<br />
Both T. radiata and D. aligarhensis are present in some parts <strong>of</strong> both<br />
counties and may cause parasitisation <strong>of</strong> D. citri in excess <strong>of</strong> 90% (Lama and<br />
Amtya 1991; Lama et al. 1987).<br />
PHILIPPINES<br />
Citrus greening, also known as citrus leaf mottle, was already causing<br />
serious damage in the early 1960s. However, as late as 1988, the windward<br />
side <strong>of</strong> Mindoro island with an average rainfall <strong>of</strong> 3000 mm was virtually<br />
free <strong>of</strong> D. citri and citrus greening, presumably due to the adverse effects <strong>of</strong><br />
high rainfall (Aubert 1989a). D. aligarhensis was reared from D. citri<br />
(25.7% parasitisation) and also 4 hyperparasitoids (Marietta sp. and 3<br />
unidentified species), resulting in an overall mortality <strong>of</strong> D. citri <strong>of</strong> 48.3%. A<br />
Beauveria sp. attacked many psyllids and in turn was parasitised by another<br />
ascomycete, probably a Melanospora sp. (Gavarra and Mercado 1989).<br />
Later (Mercado et al. 1991), up to 62.2% parasitisation by D. aligarhensis
RƒUNION<br />
4.7 Diaphorina citri 129<br />
was reported in Mindoro. In another study, Gavarra et al. (1990) recorded<br />
that, in addition to the primary parasitoid D. aligarhensis (17.6 to 36.1%<br />
parasitisation), 5 hyperparasitoids were reared from D. citri: Marietta<br />
leopardina (= M. javensis), Tetrastichus sp., Psyllaephagus sp., Chiloneurus<br />
sp. and Signiphora sp.<br />
Because it was apparently absent from the Philippines (Baltazar 1966),<br />
Tamarixia radiata was introduced from RŽunion in 1988, but attempts to<br />
rear it failed. A second consignment late that year was soon followed by the<br />
discovery <strong>of</strong> it nearby in the field in April 1989, with recoveries continuing<br />
in 1990 (Gavarra et al. 1990). However, Mercado et al. (1991) expressed<br />
some doubts that it had become established. It is thus not clear whether<br />
T. radiata ever occurred naturally in the Philippines.<br />
The rainy, windward, east side <strong>of</strong> RŽunion has much lower D. citri<br />
populations and citrus trees there are much less exposed to transmission <strong>of</strong><br />
greening (Aubert 1989a). Quilici (1989) has provided a valuable overview<br />
<strong>of</strong> the biological control <strong>of</strong> citrus psyllids in RŽunion. In the early 1970s,<br />
RŽunion and Mauritius were the only places known where Diaphorina citri<br />
and Trioza erytreae, the two psyllid vectors <strong>of</strong> citrus greening disease,<br />
occurred (Aubert 1987c). (Both are now known also from Saudi Arabia and<br />
Yemen: BovŽ 1986). Both psyllids were abundant in RŽunion and Mauritius<br />
and citrus greening was seriously affecting citrus production in both islands.<br />
The <strong>Asian</strong> citrus psyllid D. citri was most abundant below 500 m in the<br />
hotter and drier leeward side <strong>of</strong> RŽunion, where the average rainfall is below<br />
1000 m. On the other hand, the drought-sensitive African psyllid T. erytreae<br />
was particularly abundant in the cooler, moister regions above 600 m. The<br />
only nymphal parasitoid <strong>of</strong> both species was the relatively ineffective<br />
D. aligarhensis. Several predators exerted little control.<br />
Tamarixia dryi was introduced in 1974 from South Africa and, after<br />
elimination <strong>of</strong> hyperparasitoids, was mass produced and released in<br />
neglected, unsprayed citrus orchards colonised by D. citri. Populations <strong>of</strong><br />
Trioza erytreae diminished progressively from 1979 to 1982, since when<br />
T. erytreae has not been recorded, although Tamarixia dryi is still abundant<br />
on another psyllid, Trioza litseae (= T. eastopi).<br />
In 1978 Tamarixia radiata was introduced from India and released on<br />
the leeward (west) side <strong>of</strong> RŽunion. From 1982 onwards D. citri has virtually<br />
disappeared from commercial citrus orchards, although on Murraya<br />
paniculata hedges there persist low populations <strong>of</strong> D. citri which are<br />
parasitised by T. radiata and occasionally, especially at higher altitudes, by<br />
D. aligarhensis.
130 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
The excellent success <strong>of</strong> these two biological control projects is ascribed<br />
to 3 factors:<br />
(i) the absence <strong>of</strong> hyperparasitoids <strong>of</strong> the primary parasitoids <strong>of</strong> Tamarixia<br />
dryi and T. radiata.<br />
(ii) the presence <strong>of</strong> an alternative host for Tamarixia dryi, which enabled it<br />
to maintain itself as Trioza erytreae populations diminished.<br />
(iii) the maintenance on Murraya paniculata hedges <strong>of</strong> low populations <strong>of</strong><br />
D. citri, heavily parasitised by both T. radiata and D. aligarhensis<br />
(Aubert 1987c; Etienne and Aubert 1980; Quilici 1989).<br />
SAUDI ARABIA AND YEMEN<br />
In Saudi Arabia, both D. citri and Trioza erytreae are present; the former is<br />
the main vector <strong>of</strong> citrus greening. Both vectors are also present in Yemen<br />
where citrus greening at high elevations is probably the African form<br />
transmitted by T. erytreae (BovŽ 1986). In Saudi Arabia lime and lemon<br />
trees are favoured hosts <strong>of</strong> D. citri (Wooler et al. 1974).<br />
TAIWAN<br />
The nymphal ectoparasitoid Tamarixia radiata was introduced from<br />
RŽunion and, after mass rearing, released widely in citrus orchards and on<br />
Murraya paniculata hedges from 1984 to 1988. It became established,<br />
attaining parasitisation rates <strong>of</strong> up to 100%. Hyperparasitisation was<br />
initially, in 1988, below 1% (Chien 1989; Chien et al. 1988; Su and Chen<br />
1991), but by 1991 had risen gradually to 5.6% (Chien et al. 1991a). This is<br />
in contrast with levels <strong>of</strong> 72% by some 10 species attacking the native<br />
Diaphorencyrtus aligarhensis. High levels <strong>of</strong> attack on D. aligarhensis is<br />
one reason why this species is far less effective against the citrus psyllid than<br />
the introduced T. radiata (Chien et al. 1988). T. radiata was capable <strong>of</strong><br />
maintaining D. citri at low densities in relatively stable habitats where<br />
Murraya paniculata was occasionally present, whereas D. aligarhensis has<br />
adapted to unstable habitats. However, it only provides partial control due to<br />
25.5 to 51.1% hyperparasitisation throughout the island. In the Taichung<br />
area, T. radiata was more abundant than D. aligarhensis, but the peak<br />
abundance <strong>of</strong> the two did not overlap and the total parasitisation varied from<br />
80 to 100% from February to April and 32 to 80% for the remainder <strong>of</strong> the<br />
year. Application <strong>of</strong> methomyl gave good control <strong>of</strong> D. citri, but it reduced<br />
parasitisation to a level <strong>of</strong> 0 to 4%. In an untreated citrus orchard with only<br />
0.1 to 0.4 D. citri adults per 10 cm length branch, the parasitoids caused 15.5<br />
to 46.7% parasitisation (Chien et al. 1991a). Citrus greening in Taiwan is<br />
known as likubin or leaf mottle disease.
VIETNAM<br />
4.7 Diaphorina citri 131<br />
Tamarixia radiata was found parasitising 3 to 10% <strong>of</strong> 4th and 5th instar<br />
nymphs <strong>of</strong> D. citri and Diaphorencyrtus aligarhensis was also present<br />
(Myartzeva and Trijapitzyin 1978; Trung 1991; van Lam 1996).<br />
Major natural enemies<br />
HYMENOPTERA<br />
Diaphorencyrtus aligarhensis Hym.: Encyrtidae<br />
This primary endoparasitoid was described by Shafee et al. (1975), from<br />
India as Aphidencyrtus aligarhensis. Its hosts include Diaphorina citri,<br />
D. auberti, D. cardiae and Psylla sp. (Qing and Aubert 1990).<br />
The D. citri mummy parasitised by D. aligarhensis is brownish and<br />
hemi-spherical and encloses the parasitoid pupa. The parasitoid emerges<br />
from the side <strong>of</strong> the abdomen. Development from egg to adult takes 18 to 23<br />
days at 25 ± 1¡C and 80 to 85% relative humidity (Tang and Huang 1991).<br />
No males occur and unmated females produce females. On average, 4.5 eggs<br />
are laid per day with an average production <strong>of</strong> 144 per female. Third and 4th<br />
instar D. citri nymphs are preferred over 2nd instar, and 1st and 5th instars<br />
are not parasitised. Usually only one egg is inserted into each host, but the<br />
haemolymph <strong>of</strong> many young nymphs is consumed leading to their death<br />
(Tang and Huang 1991).<br />
Tamarixia radiata Hym.: Eulophidae<br />
This ectoparasitoid was described from India (Waterston 1922) where it is<br />
an important species (Husain and Nath 1924). It has been recorded in China<br />
(in 1982: Tang 1989), Indonesia (Nurhadi 1989), Malaysia (Lim et al. 1990),<br />
Nepal (Lama et al. 1988), Saudi Arabia (Aubert 1984a), Thailand (Qing and<br />
Aubert 1990) and Vietnam (Myartzeva and Trijapitzyin 1978). T. radiata<br />
has been introduced to, and established in, RŽunion (Aubert and Quilici<br />
1984), Mauritius (Quilici 1989) and Taiwan (Chiu et al. 1988).<br />
T. radiata was found to be the dominant parasitoid <strong>of</strong> D. citri on Murraya<br />
paniculata in Fujian, comprising 62.6% <strong>of</strong> all parasitoids and<br />
hyperparasitoids emerging. The second in abundance was the<br />
hyperparasitoid Tetrastichus sp., most <strong>of</strong> which were bred from T. radiata,<br />
an average <strong>of</strong> 21.8% hyperparasitisation, rising to a maximum <strong>of</strong> 87.9%,<br />
whereas the other primary parasitoid D. aligarhensis was hyperparasitised to<br />
an average <strong>of</strong> 34.1% (Tang 1989).<br />
The T. radiata female oviposits ventrally between the thorax and<br />
abdomen <strong>of</strong> the nymph, preferably <strong>of</strong> the 5th instar, and its fully grown larva<br />
spins silk to attach itself and its host to the plant substrate. The D. citri<br />
mummy parasitised by T. radiata has a dark brown, flattened body and the
132 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
parasitoid pupa remains external to, and on the ventral surface <strong>of</strong>, the host.<br />
The adult wasp emerges via a hole cut through the thorax <strong>of</strong> the host (Qing<br />
1990; Tang and Huang 1991). Under favourable conditions, parasitisation<br />
can exceed 90%, as in India (Husain and Nath 1927) and also in RŽunion,<br />
Nepal and Taiwan (Quilici and Fauvergue 1990).<br />
Male T. radiata are capable <strong>of</strong> multiple matings, but females usually<br />
mate only once. The egg to adult period was 11.4 days (egg 1.9, larva 4.0,<br />
prepupa 0.6, pupa 4.9 days), females lived 23.6 days and males lived 14.8<br />
days (Chien et al. 1991a,b; Chu and Chien 1991). Fauvergue and Quilici<br />
(1991) report reduction <strong>of</strong> the duration <strong>of</strong> immature stages with increasing<br />
temperature from 17 days at 20¡C to 8 days at 30¡C. Adult females lived 37<br />
days at 20¡C and 8 days at 35¡C. Females kill some 80% <strong>of</strong> D. citri hosts by<br />
parasitisation and 20% by host feeding. When 40 psyllids were presented per<br />
day a female killed 513 psyllids in a lifetime. At an optimum temperature <strong>of</strong><br />
25¡C, 24, 5th instar nymphs were killed per day (Chien et al. 1993). Adult<br />
parasitoids can be cold stored at 8¡C for between 46 and 60 days (Chien et al.<br />
1993). Oosorption occurred when hosts were unavailable. This extended the<br />
reproductive period, but diminished the total number <strong>of</strong> eggs laid (Chien et<br />
al. 1994b). Feeding by females on the honeydew produced by the host and on<br />
host haemolymph provides nutrients for egg production. The parasitoid fed<br />
on the exudate <strong>of</strong> 28% <strong>of</strong> host eggs parasitised (Chien et al. 1994a). The<br />
optimal host density over the entire T. radiata lifetime was found to be 2 to 8<br />
per day, <strong>of</strong> which 90 to 94% were utilised. For the peak oviposition period,<br />
optimal density was 2 to 20, <strong>of</strong> which 87 to 90% were utilised (Chien et al.<br />
1995).<br />
The sex ratio <strong>of</strong> T. radiata is 1:3 in favour <strong>of</strong> females. Unmated females<br />
give rise only to male <strong>of</strong>fspring. Oviposition occurs on 3rd, 4th and 5th instar<br />
nymphs and there is discrimination against ovipositing in nymphs<br />
containing older D. aligarhensis larvae. The average number <strong>of</strong> <strong>of</strong>fspring is<br />
reported as 134 with 6.5 eggs laid per day (Tang and Huang 1991).<br />
Observations in China indicate that T. radiata is more affected by low<br />
temperatures than D. citri. Thus T. radiata breeds more effectively in<br />
Xiamen, where the lowest winter temperature is 3.9¡C, than in Fuzhou,<br />
where overwintering is jeopardised by lowest minimum temperatures <strong>of</strong><br />
Ð2.5¡C (Aubert 1990).
4.7 Diaphorina citri 133<br />
Tetrastichus sp. Hym.: Eulophidae<br />
This undescribed species is an important hyperparasitoid <strong>of</strong> Tamarixia<br />
radiata in China. Average hyperparasitisation amounted to nearly 25%, with<br />
a maximum <strong>of</strong> 87.9%. The genus Tetrastichus contains more than 150<br />
species attacking a wide variety <strong>of</strong> hosts.<br />
ARACHNIDA<br />
Marpissa tigrina: Salticidae<br />
The number <strong>of</strong> D. citri consumed daily by an individual spider increased<br />
with an increase in available prey up to 40. Further increases in prey<br />
numbers reduced predation. The results suggest that M. tigrina is a highly<br />
efficient predator <strong>of</strong> D. citri (Sanda 1991).<br />
Comments<br />
Diaphorina citri and its two primary parasitoids, Diaphorencyrtus<br />
aligarhensis and Tamarixia radiata, are (especially the first two species)<br />
very widespread in <strong>Southeast</strong> Asia. In these countries the prospects for<br />
biological control are unpromising, although the careful timing <strong>of</strong> least<br />
harmful, but still effective, insecticides (or, preferably, special petroleum<br />
oils) would favour a build up <strong>of</strong> the parasitoids. The role played by hedges<br />
and other plantings <strong>of</strong> the common, favoured host, jasmin orange, Murraya<br />
paniculata in encouraging either D. citri or its parasitoids is worthy <strong>of</strong><br />
careful investigation, for this may differ widely according to the insecticidal<br />
treatments in the nearby citrus plantings. Overhead irrigation to reduce<br />
numbers <strong>of</strong> eggs and young nymphs during periods <strong>of</strong> growth flushes is<br />
probably uneconomical in most situations, but is possibly a factor to<br />
consider in a pest management approach.<br />
Although D. aligarhensis appears to be a less effective parasitoid than<br />
T. radiata, it still may contribute useful suppression <strong>of</strong> D. citri where it can<br />
be introduced without encountering hyperparasitoids.<br />
In contrast to much <strong>of</strong> <strong>Southeast</strong> Asia, the prospects for successful<br />
biological control <strong>of</strong> D. citri appear to be promising for countries that have<br />
been recently invaded, particularly if there are few or no hyperparasitoids<br />
already present that are capable <strong>of</strong> attacking T. radiata and/or<br />
D. aligarhensis. In this context it may be valuable to explore the situation in<br />
Irian Jaya and Timor where D. citri has been recorded only recently.<br />
Successful biological control there may slow the spread <strong>of</strong> D. citri into<br />
Papua New Guinea, Australia and the oceanic Pacific. Brazil has a range <strong>of</strong><br />
native psyllids that are attacked by Tamarixia spp. so it is possible that there<br />
are already hyperparasitoids present that would attack T. radiata were it to<br />
be introduced.
134 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Since Tamarixia leucaenae attacks both Heteropsylla cubana and<br />
H. spinulosa, it would be valuble to know whether Tamarixia radiata will<br />
also attack Heteropsylla spinulosa. This is the psyllid that has been<br />
successfully introduced to Papua New Guinea, Australia and some oceanic<br />
Pacific countries for the biological control <strong>of</strong> creeping, sensitive plant,<br />
Mimosa invisa. If it does attack H. spinulosa, there would clearly be a<br />
conflict <strong>of</strong> interest between biological control <strong>of</strong> D. citri and <strong>of</strong> M. invisa,<br />
were Heteropsylla spinulosa to be considered for the latter. However, at<br />
least some species <strong>of</strong> Tamarixia are satisfactorily host restricted and it is<br />
quite possible that T. radiata is one <strong>of</strong> them.
4.8 Dysdercus cingulatus<br />
India<br />
Myanmar<br />
+<br />
20°<br />
Laos<br />
+<br />
0°<br />
20°<br />
China<br />
+<br />
Thailand<br />
+<br />
Cambodia<br />
P<br />
Vietnam<br />
++<br />
P<br />
+ Brunei<br />
Malaysia<br />
+<br />
Singapore<br />
++<br />
Indonesia<br />
Taiwan<br />
++<br />
Philippines<br />
Australia<br />
Papua<br />
New Guinea<br />
P<br />
135<br />
Dysdercus cingulatus is native to the <strong>Southeast</strong> <strong>Asian</strong> region.<br />
No parasitoids <strong>of</strong> the cotton stainer are known and surprisingly few predators are<br />
reported. No effective parasitoids <strong>of</strong> the many Dysdercus species that occur in other parts<br />
<strong>of</strong> the world are known. On present knowledge, therefore, the prospects for classical<br />
biological control <strong>of</strong> this bug would appear to be very remote.<br />
20°<br />
0°<br />
20°
136 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Dysdercus cingulatus (Fabricius)<br />
Rating<br />
Origin<br />
Distribution<br />
Biology<br />
Hemiptera, Pyrrhocoridae<br />
cotton stainer, red cotton bug, red seed bug<br />
<strong>Southeast</strong> Asia China Southern and Western Pacific<br />
++ Viet, Indo, Phil<br />
11 + Myan, Thai, Laos,<br />
Msia, Sing<br />
+<br />
P Brun P PNG, Sol Is, Van, Marianas,<br />
Carolines, N Cal.<br />
<strong>Southeast</strong> Asia.<br />
Waterhouse (1993a) was in error in accepting Kalshoven (1981), who listed<br />
the distribution <strong>of</strong> D. cingulatus as Ôwidespread from the Mediterranean to<br />
AustraliaÕ. In fact, it occurs from north eastern India through Bangladesh to<br />
all <strong>Southeast</strong> <strong>Asian</strong> countries, southern China, southern Japan, Irian Jaya,<br />
Papua New Guinea, eastern Australia, Saipan, Palau, Pohnpei, Yap,<br />
Solomon Is., Vanuatu and New Caledonia (CIE 1985). There are some 50<br />
species <strong>of</strong> Dysdercus,<br />
some <strong>of</strong> which appear to be native to the African,<br />
Ethiopian, Southern <strong>Asian</strong> and American regions respectively (Freeman<br />
1947). D. cingulatus is not recorded from Africa, Europe, Western Asia or<br />
the Americas.<br />
Eggs are usually laid singly in batches <strong>of</strong> about 100 (range 25 to 112) in<br />
small depressions in the soil under the host plant. They are camouflaged with<br />
soil particles or other debris. Some 80% <strong>of</strong> the eggs hatch in about 6 days if<br />
there is the essential high humidity. The optimum hatch occurs at 30¡C and<br />
80% RH. There are 5 nymphal instars which take 25 to 27 days to complete<br />
and the oviposition to adult period is thus 31 to 33 days. The first instar<br />
nymphs do not feed. Later instars suck sap with a preference for pods and<br />
seeds. All stages are gregarious. The male:female ratio is 3:2 and mating<br />
takes place readily and repeatedly, pairs <strong>of</strong>ten remaining in copula for 2 to 4<br />
days, during which they move and feed (Srivastava and Bahadur 1958;<br />
Thomas 1966; Ahmad and Aziz 1982, 1983, Farine and Lobreau 1984,<br />
Siddiqi 1985, 1987; Khoo et al. 1991).
Host plants<br />
Damage<br />
4.8<br />
Dysdercus cingulatus<br />
137<br />
D. cingulatus is one <strong>of</strong> the commonest insects in cultivated land in<br />
Indonesia. Adults range from 11 to 17 mm in length and are orange and black<br />
in colour, with a characteristic white band on the pronotum and single large<br />
black spots on each orange forewing.<br />
Both adults and nymphs produce a complex mixture <strong>of</strong> compounds as a<br />
defensive secretion (Farine et al. 1992, 1993) and females produce a sex<br />
pheromone from glands in the thorax (Siddiqi 1988; Siddiqi and Khan<br />
1982).<br />
The principal host plants <strong>of</strong> D. cingulatus are in the families Malvaceae and<br />
Bombacaceae and include cotton, kapok, okra and rosella. It was the most<br />
important pest <strong>of</strong> cotton in the coastal districts <strong>of</strong> Malaysia in the early days<br />
and also fed on the seeds <strong>of</strong> kapok, rosella, hibiscus, hemp, okra and other<br />
malvaceous plants (Jack and Sands 1922; Dresner 1955). In a host<br />
preference test in Malaysia, the following order was recorded kapok > okra ><br />
urena > maize > sorghum (Chong 1975). It has also been recorded on wheat<br />
(Srivastava and Gupta 1971), pearl millet, Pennisetum glaucum<br />
(= P. americanum)<br />
(Ahmad 1979) and a range <strong>of</strong> weeds.<br />
Like other species <strong>of</strong> Dysdercus throughout the world D. cingulatus is most<br />
important as a pest <strong>of</strong> cotton. The sap removed and the fungus introduced<br />
into the punctures caused when feeding on the developing cotton bolls<br />
causes staining <strong>of</strong> the lint, giving rise to one <strong>of</strong> its common names.<br />
Natural enemies<br />
Very little has been published on the natural enemies <strong>of</strong> D. cingulatus and no<br />
parasitoids are known. In view <strong>of</strong> the large number <strong>of</strong> parasitoids <strong>of</strong> Nezara<br />
viridula eggs (Table 4.12.1), it seems strange that none are recorded from<br />
eggs <strong>of</strong> the cotton stainer. There appear to be no statements in the literature<br />
that they have been looked for unsuccessfully.<br />
The few records retrieved <strong>of</strong> natural enemies <strong>of</strong> D. cingulatus are shown<br />
in Table 4.8.1. The pyrrhocorid predator Antilochus coquebertii has been<br />
reported attacking the cotton stainer in India and Malaysia (Yunus & Ho<br />
1980; Zaidi 1985). It is believed to inject saliva containing proteolytic<br />
enzymes into eggs, nymphs or adults before sucking out the liquified<br />
contents. In the Philippines an ectoparasitic mite, Hemipterotarseius sp. was<br />
found on the pronotum <strong>of</strong> adult bugs, two species <strong>of</strong> spiders were observed<br />
attacking bugs and a nematode was found in the abdomen <strong>of</strong> a female. No
138 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Comment<br />
Table 4.8.1<br />
egg parasitoids were found (Encarnacion 1970). Singh and Bardhan (1974)<br />
showed that D. cingulatus was moderately susceptible to the DD 136 strain<br />
<strong>of</strong> the nematode Steinernema carcocapsae.<br />
A superficial review <strong>of</strong> the literature on the natural enemies <strong>of</strong> other<br />
species <strong>of</strong> Dysdercus revealed two tachinid parasitoids, one <strong>of</strong> each in South<br />
America and Africa and three asassin bug predators in Africa but no egg<br />
parasitoids (Table 4.8.2).<br />
Too little is known about the natural enemies <strong>of</strong> D. cingulatus to indicate<br />
whether there are any prospects for classical biological control or for<br />
manipulating them where they already occur. Since D. cingulatus now<br />
appears to have evolved in the <strong>Southeast</strong> <strong>Asian</strong> region and since effective<br />
natural enemies <strong>of</strong> related species elsewhere are not known, the prospects<br />
for its classical biological control would appear to be remote.<br />
Natural enemies <strong>of</strong> Dysdercus cingulatus<br />
Enemy<br />
INSECTA<br />
HEMIPTERA<br />
Country Reference<br />
PYRRHOCORIDAE<br />
Antilochus coquebertii<br />
India<br />
Malaysia<br />
Zaidi 1985<br />
Thomas 1966; Yunus & Ho 1980<br />
ACARINA<br />
Hemipterotarseius sp.<br />
ARACHNIDA<br />
Philippines Encarnacion 1970<br />
Spider 1 Philippines Encarnacion 1970<br />
Spider 2<br />
NEMATODA<br />
Philippines Encarnacion 1970<br />
Species 1<br />
FUNGI<br />
Philippines Encarnacion 1970<br />
Aspergillus flavus<br />
India Kshemkalyani et al. 1989
Table 4.8.2<br />
4.8<br />
Natural enemies <strong>of</strong> Dysdercus spp.<br />
Dysdercus cingulatus<br />
Enemy<br />
HEMIPTERA<br />
REDUVIIDAE<br />
Country Reference<br />
Phonoctonus nigr<strong>of</strong>asciatus Zimbabwe Sweeney 1960<br />
Phonoctonus subimpictus Ivory Coast Galichet 1956<br />
Phonoctonus sp. Mozambique Barbosa 1950<br />
DIPTERA<br />
TACHINIDAE<br />
Acaulona brasiliana<br />
Bogosia helva<br />
Argentina Blanchard 1966<br />
Ivory Coast Galichet 1956<br />
139
4.9 Dysmicoccus brevipes<br />
India<br />
20°<br />
0°<br />
20°<br />
Myanmar<br />
Laos<br />
China<br />
+<br />
Thailand<br />
P<br />
Cambodia<br />
+<br />
Vietnam<br />
+++<br />
P<br />
++ Brunei<br />
Malaysia<br />
Singapore<br />
++<br />
Indonesia<br />
Taiwan<br />
+<br />
++<br />
Philippines<br />
Australia<br />
Papua<br />
New Guinea<br />
+<br />
141<br />
The pineapple mealybug, Dysmicoccus brevipes,<br />
is <strong>of</strong> Central or South American<br />
origin. It occurs in both parthenogenetic and bisexual forms, the females <strong>of</strong> which are<br />
morphologically indistinguishable, although they may possibly prove to be distinct species.<br />
The closely-related, bisexual, D. neobrevipes also occurs on pineapple.<br />
Attempts have been made by several countries to establish natural enemies for<br />
biological control, but none has had any great success without the control <strong>of</strong> attendant<br />
ants. When ants were controlled the mealybug was no longer a problem. Ant control is less<br />
likely to be an answer to the problem in the absence <strong>of</strong> suitable natural enemies. It is<br />
desirable, therefore, to establish appropriate natural enemies in anticipation <strong>of</strong> effective<br />
ant control, an aspect which is now being actively investigated in Hawaii.<br />
20°<br />
0°<br />
20°
142 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Dysmicoccus brevipes (Cockerell)<br />
Rating<br />
Origin<br />
Distribution<br />
Taxonomy<br />
Hemiptera, Pseudococcidae<br />
pineapple mealybug<br />
<strong>Southeast</strong> Asia China Southern and Western Pacific<br />
+++ Viet +++ Cook Is, Guam<br />
10 ++ Msia, Indo, Phil 17 ++ FSM, Niue, Van<br />
+ Camb + + Kiri, N Cal, PNG, Sam,<br />
Sol Is<br />
P Brun, Thai P Fiji, Fr P, A Sam.<br />
Tok, Tong, Tuv<br />
Carter (1935) considered D. brevipes to be native to South America,<br />
although Ferris (1950) believed it to be <strong>of</strong> North American origin. The<br />
pineapple plant ( Ananas comosus)<br />
is thought to be native to South America<br />
and has been known in Central America since pre-Columbian times.<br />
Although the pineapple mealybug is widely polyphagous, if it evolved in<br />
association with pineapple plants, it would appear to be logical to assign its<br />
origin to Central and/or South America.<br />
D. brevipes is one <strong>of</strong> the most widespread mealybugs, occurring throughout<br />
the tropics and in many temperate areas, especially those where pineapples<br />
are grown (Williams and Watson 1988). These include tropical Africa,<br />
Mauritius, tropical Asia, <strong>Southeast</strong> Asia, Taiwan, Australia, Pacific islands<br />
(including Hawaii), southern USA (Florida, Louisiana) West Indies and<br />
Central and South America (Bartlett in Clausen 1978).<br />
Dysmicoccus brevipes was earlier known as Pseudococcus brevipes,<br />
but its<br />
genus was changed by Ferris (1950). Before 1959 it was confused with a<br />
similar mealybug (<strong>of</strong>ten on the same host plants), which was described by<br />
Beardsley (1959) as Dysmicoccus neobrevipes.<br />
Both species occur in<br />
Hawaii, where D. brevipes is parthenogenetic and the females are pink,<br />
whereas D. neobrevipes is bisexual and females are grey in colour.<br />
Parthenogenetic D. brevipes is known also from Jamaica and West Africa<br />
(Beardsley 1965).
Biology<br />
4.9<br />
Dysmicoccus brevipes<br />
143<br />
In some countries (Ivory Coast, Madagascar, Dominican Republic,<br />
Martinique, Malaysia) both sexes <strong>of</strong> D. brevipes occur (Beardsley 1965;<br />
Lim 1973). D. neobrevipes is known from many countries in Central and<br />
South America and probably also originated there (Williams and Watson<br />
1988). It is known from Mexico, Jamaica, American Samoa and Samoa,<br />
Cook Is, Kiribati and Guam. In <strong>Southeast</strong> Asia it occurs in Malaysia, the<br />
Philippines and Thailand (where there have been recent serious outbreaks<br />
(Beardsley 1965; Rohrback et al. 1988; Williams and Watson 1988).<br />
D. brevipes females are broadly oval to rotund in shape, pinkish in colour,<br />
and have a thick waxy covering with short conical waxy projections<br />
(Kalshoven 1981). In Hawaii, D. brevipes is parthenogenetic and<br />
ovoviviparous (i.e. it produces its young alive). About 250 crawlers are<br />
produced per female over a 3 to 4 week period and take some 34 days to<br />
mature. Females start producing young about 25 days after the third moult<br />
(Ito 1938).<br />
In peninsular Malaysia the bisexual form is widespread but the<br />
parthenogenetic form was not found (Lim 1973). In the male, there are two<br />
nymphal, one prepupal and one pupal instar <strong>of</strong> 10, 6, 3 and 4 days duration<br />
respectively. Adult males live for 1 to 3 days, whereas adult females live 17<br />
to 49 days. The female has 3 nymphal instars, lasting 10, 7 and 7 days<br />
respectively. A female produces 19 to 137 <strong>of</strong>fspring with a sex ratio <strong>of</strong> 1:1<br />
(Lim 1973). The bisexual form in Malaysia has a 10 day shorter life cycle<br />
than the parthenogenetic form in Hawaii and might have as many as 9<br />
generations a year (Lim 1973). The bisexual form <strong>of</strong> D. brevipes is capable<br />
<strong>of</strong> producing green spotting on pineapple leaves, whereas the<br />
parthenogenetic form is not (Beardsley 1965).<br />
In India, Ghose (1983) studied the parthenogenetic form <strong>of</strong> D. brevipes<br />
at 30¡C and 60Ð66% R.H. The nymphs completed their development in 19<br />
days and, after a pre-oviposition period <strong>of</strong> 16 days, produced an average <strong>of</strong><br />
240 young over a period <strong>of</strong> 40 days.<br />
D. brevipes occurs mainly on the underground parts <strong>of</strong> the pineapple<br />
stem (where the stem is covered by leaf bases) and on the roots. Leaves or<br />
fruit are less heavily infested, except on weak plants. However the bisexual<br />
form also infests the crown <strong>of</strong> the pineapple plant (Rohrbach et al. 1988).<br />
By comparison, D. neobrevipes,<br />
which is always bisexual, is found only<br />
on the aerial parts <strong>of</strong> the pineapple plant Ñ leaves, stems, aerial roots,<br />
flowers and fruit clusters. Unlike D. brevipes,<br />
it does not infest grasses . In<br />
Hawaii, only D. neobrevipes causes green spotting <strong>of</strong> pineapple leaves<br />
(Beardsley 1959).
144 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Hosts<br />
Damage<br />
The pineapple mealybug is generally attended by ants seeking<br />
honeydew. They not only protect it against natural enemies but also assist in<br />
dispersal by transporting it to new plants. Where appropriate natural enemies<br />
are present, but attendant ants are not, the mealybug is no longer a problem<br />
(Beardsley et al. 1982). The identity <strong>of</strong> the ants varies from place to place,<br />
although 3 very widespread species are <strong>of</strong>ten involved, the bigheaded ant,<br />
Pheidole megacephala,<br />
the Argentine ant, Iridomyrmex humilis,<br />
and the fire<br />
ant Solenopsis geminata.<br />
The gradual invasion <strong>of</strong> new pineapple plantings<br />
by ants is accompanied by progressive outbreaks <strong>of</strong> mealybugs, so ant<br />
control is essential. It is interesting that mated queens <strong>of</strong> P. megacephala<br />
must, following the nuptial flight, rejoin established colonies to survive.<br />
Thus, invasion into new territory is accomplished by extension <strong>of</strong> existing<br />
nests, a feature that is <strong>of</strong> importance in controlling the big-headed ant<br />
(Beardsley et al. 1982).<br />
D. brevipes occurs widely on its preferred host, pineapple, wherever this is<br />
grown in moist tropical or subtropical areas, but it can be found on almost<br />
any kind <strong>of</strong> plant and is sometimes a pest <strong>of</strong> sugarcane and bananas. It also<br />
occurs on areca palm, c<strong>of</strong>fee, groundnut, oil palm, rice, sisal, soybean,<br />
Pandanus palm and a range <strong>of</strong> grasses and weeds (Clausen 1978; Kalshoven<br />
1981; Khoo et al. 1991).<br />
In Hawaii, through their feeding, both D. brevipes and D. neobrevipes<br />
produce symptoms <strong>of</strong> toxicosis on pineapple, including stunting, reddening<br />
and wilting <strong>of</strong> young plants, due to what is now held to be a virus, and termed<br />
pineapple mealybug wilt. D. neobrevipes,<br />
but not the parthenogenetic form<br />
<strong>of</strong> D. brevipes, also produces a green spotting on the leaves. Where the<br />
bisexual form <strong>of</strong> D. brevipes occurs elsewhere it is capable <strong>of</strong> producing<br />
green spotting.<br />
Unless collected by ants, honeydew produced by the mealybugs, leads to<br />
the massive growth <strong>of</strong> sooty moulds which reduces photosynthesis, affects<br />
sales <strong>of</strong> fruit and attracts Carpophilus spp. beetles, which contaminate<br />
canned fruit. Large colonies <strong>of</strong> D. brevipes in leaf sheaths near the roots <strong>of</strong><br />
sugarcane result in poor growth and, when on groundnuts, cause the seed<br />
pods to become discoloured.
Natural enemies<br />
4.9<br />
Dysmicoccus brevipes<br />
145<br />
A number <strong>of</strong> coccinellid predators are recorded attacking D. brevipes (Table<br />
4.9.1) and it is possible that some <strong>of</strong> these have a sufficiently narrow host<br />
range to be considered for introduction. However, it is more likely that the<br />
dipterous and lepidopterous predators will be more specific. The encyrtid<br />
parasitoids would appear to be even more promising and it seems that the full<br />
range <strong>of</strong> species attacking D. brevipes in Central and South America has not<br />
yet been identified.<br />
Attempts at biological control<br />
HAWAII<br />
A number <strong>of</strong> natural enemies, mainly encyrtid parasitoids and coccinellid<br />
and dipterous predators have been successfully introduced, in particular into<br />
Hawaii, Puerto Rico and the Philippines (Table 4.9.2). Details are provided<br />
in the country accounts that follow. It will become clear that a substantial<br />
degree <strong>of</strong> control <strong>of</strong> D. brevipes can be achieved in the absence <strong>of</strong> ants,<br />
which clearly protect the mealybugs against parasitoids and predators.<br />
The pineapple mealybug has, for many years, constituted the most serious<br />
insect problem <strong>of</strong> the pineapple industry (Carter 1932; Beardsley 1959) and<br />
it is also a minor pest <strong>of</strong> sugarcane and bananas. An encyrtid wasp<br />
Euryrophalus schwarzi (= E. pretiosa)<br />
was reared from D. brevipes<br />
collected from sugarcane (Beardsley 1959).<br />
In the early 1920s several natural enemies were introduced from Mexico<br />
and Panama, but none became established (Rohrbach et al. 1988). Anagyrus<br />
ananatis and Hambletonia pseudococcina from Central America and Brazil<br />
were established in 1935Ð36 and were effective in Maui where the dominant<br />
ant was the crazy ant Paratrechina longicornis.<br />
Parasitisation was high and<br />
pineapple wilt quite severe (Carter 1945). Other introductions known to<br />
have become established are an encyrtid parasitoid ( Euryrhopalus<br />
propinquus),<br />
a cecidomyiid predator ( Vincentodiplosus pseudococci),<br />
and<br />
two less effective coccinellid predators ( Scymnus (= Nephus)<br />
bilucenarius<br />
and Scymnus uncinatus)<br />
(Lai and Funasaki 1986).<br />
Overall, although the biological control <strong>of</strong> the pineapple mealybug has<br />
not been completely successful, a considerable reduction in abundance has<br />
resulted from the combined action <strong>of</strong> the cecidomyiid predator<br />
Vincentodiplosis and the encyrtid parasitoids Anagyrus ananatis and<br />
Hambletonia pseudococcina.<br />
They are highly effective only where ants are<br />
adequately controlled (Clausen 1978). Coccinellids are important for short<br />
periods, particularly in the middle <strong>of</strong> large plantings, where the absence <strong>of</strong><br />
ants renders D. brevipes exposed to attack (Carter 1935, 1944).
Table 4.9.1<br />
Natural enemies <strong>of</strong> Dysmicoccus brevipes<br />
Country References<br />
HEMIPTERA<br />
DIASPIDIDAE<br />
Diaspis bromeliae<br />
ORTHOPTERA<br />
Mauritius Jepson 1939a<br />
Conocephalus saltator<br />
NEUROPTERA<br />
CHRYSOPIDAE<br />
Hawaii Carter 1935<br />
Chrysopa irregularis<br />
Fiji Lever 1940<br />
Chrysopa ramburi<br />
Fiji Lever 1940<br />
Chrysopa sp.<br />
COLEOPTERA<br />
COCCINELLIDAE<br />
Fiji Lever 1940<br />
Brachycantha sp. Guatemala Carter 1935<br />
Cryptolaemus montrouzieri<br />
Hawaii<br />
Kalshoven 1981<br />
Fiji, New Caledonia<br />
Williams & Watson 1988<br />
Cryptolaemus sp. Fiji, New Caledonia Williams & Watson 1988<br />
Rhizobius ventralis<br />
Hawaii, New Caledonia Williams & Watson 1988<br />
Scymnus apiciflavus<br />
Malaysia Yunus & Ho 1980<br />
Scymnus bilucenarius<br />
Guatemala Carter 1935<br />
Scymnus mauritiusi<br />
Mauritius Jepson 1939a<br />
Scymnus sp. Fiji, New Caledonia Cohic 1958<br />
Sticholotis quatrosignata<br />
Malaysia Yunus & Ho 1980<br />
Coccinellids<br />
Other species in introductions listed in table 4.9.2<br />
Taiwan Takahashi 1939<br />
146 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong>
Table 4.9.1 (contÕd) Natural enemies <strong>of</strong> Dysmicoccus brevipes<br />
DIPTERA<br />
Country References<br />
CECIDOMYIIDAE<br />
Schizobremia formosana Taiwan Takahashi 1939<br />
Cecidomyiid sp. 1 Guatemala Carter 1935<br />
Cecidomyiid sp. 2 Mauritius Jepson 1939a<br />
Cecidomyiid sp. 3 Puerto Rico Plank & Smith 1940<br />
DROSOPHILIDAE<br />
Gitonides perspicax Mauritius Jepson 1939a<br />
Pseudiastata nebulosa Guatemala Carter 1935<br />
LEPIDOPTERA<br />
PYRALIDAE<br />
Species 1<br />
TINEIDAE<br />
Puerto Rico Plank & Smith 1940<br />
Drosica abjectella South Africa BŸttiker 1957<br />
Species 1<br />
HYMENOPTERA<br />
ENCYRTIDAE<br />
Puerto Rico Plank & Smith 1940<br />
Aenasius acuminatus Trinidad Kerrich 1967<br />
Aenasius theobromae Trinidad Kerrich 1953<br />
Anagyrus ananatis Brazil Carter 1937;<br />
Gabriel et al. 1982<br />
Anagyrus sp. Brazil Compere 1936<br />
Encyrtid sp. 1 Brazil Compere 1936<br />
Euryrhopalus propinquus Brazil, Guyana, Hawaii Kerrich 1967<br />
4.9<br />
Dysmicoccus brevipes<br />
147
Table 4.9.1 (contÕd) Natural enemies <strong>of</strong> Dysmicoccus brevipes<br />
HYMENOPTERA<br />
ENCYRTIDAE (contÕd)<br />
Euryrhopalus schwarzi (= E. pretiosa) Guatemala<br />
Hawaii<br />
Country References<br />
Clausen 1978<br />
Beardsley 1959<br />
Hambletonia pseudococcina Brazil, Colombia, Venezuela Carter 1937<br />
Leptomastix dactylopii California Clausen 1978<br />
Pseudaphycus angustifrons Cuba Gahan 1946<br />
Pseudaphycus dysmicocci Trinidad Clausen 1978<br />
Pseusaphycus sp. Brazil Clausen 1978<br />
Thysanus niger<br />
CHALCIDIDAE<br />
Puerto Rico Bartlett 1945<br />
Species 1 Guatemala Carter 1935<br />
Species 2<br />
UNIDENTIFIED FAMILY<br />
Guatemala Carter 1935<br />
7 species<br />
ARACHNIDA<br />
Malaysia Yunus & Ho 1980<br />
spiders Hawaii Carter 1944<br />
148 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong>
Table 4.9.2 Introductions for the biological control <strong>of</strong> Dysmicoccus brevipes<br />
Country From Year Result Reference<br />
HYMENOPTERA<br />
ENCYRTIDAE<br />
Aenasius colombiensis Colombia 1935 Ð Lai & Funasaki 1986<br />
Aenasius sp. Panama 1931 Ð Lai & Funasaki 1986<br />
Anagyrus ananatis<br />
Hawaii Brazil 1934Ð35 + Carter 1937<br />
(= Anagyrus coccidivorus)<br />
Peurto Rico Brazil 1937Ð38 Ð Bartlett 1939, 1943<br />
via Hawaii<br />
Clausen 1978<br />
Anagyrus kivuensis California 1946 Ð Lai & Funasaki 1986<br />
Euryrhopalus propinquus Hawaii British<br />
Guiana<br />
1935 + Lai & Funasaki 1986<br />
Euryrhopalus schwarzi Hawaii Guatemala 1935 + Clausen 1978<br />
Hambletonia pseudococcina Hawaii Brazil 1935Ð36 Ð Carter 1937; Clausen 1978<br />
Colombia 1935Ð36 + Carter 1937; Clausen 1978<br />
Venezuela 1935Ð36 + Carter 1937<br />
Jamaica Hawaii<br />
1936 Ð Clausen 1978<br />
Puerto Rico Brazil via<br />
Hawaii<br />
1937Ð38 + Bartlett 1939<br />
Florida Puerto Rico 1944 + Annand 1945; Clausen 1956<br />
Leptomastix dactylopii Hawaii California ? Clausen 1978<br />
Pseudaphycus dysmicocci Hawaii Trinidad 1958 ? Clausen 1978<br />
Pseudaphycus sp. Hawaii Brazil 1946 ? Clausen 1978<br />
Zaplatycerus fullawayi<br />
PLATYGASTERIDAE<br />
Mexico 1930 Ð Lai & Funasaki 1986<br />
Allotropa sp. Panama 1931 Ð Lai & Funasaki 1986<br />
4.9 Dysmicoccus brevipes 149
Table 4.9.2 (contÕd) Introductions for the biological control <strong>of</strong> Dysmicoccus brevipes<br />
COLEOPTERA<br />
COCCINELLIDAE<br />
Cleothera sp. Panama 1931 Ð Lai & Funasaki 1986<br />
Cryptolaemus montrouzieri Mauritius<br />
Easter Is<br />
(Chile)<br />
S. Africa 1938Ð39 Ð<br />
+<br />
Jepson 1939b,<br />
Moutia & Mamet 1946<br />
Ripa et al. 1995<br />
Cryptolaemus sp. Taiwan ? Sakimura 1935<br />
Diomus sp. Jamaica<br />
Panama<br />
Country From Year Result Reference<br />
Hawaii 1939<br />
1931<br />
Ð<br />
Ð<br />
Lai & Funasaki 1986<br />
Lai & Funasaki 1986<br />
Hyperaspis albicollis Panama 1924 Ð Lai & Funasaki 1986<br />
Hyperaspis c-nigrum Brazil 1935 Ð Lai & Funasaki 1986<br />
Hyperaspis silvestri Philippines Hawaii 1931 + Clausen 1978<br />
Hyperaspis 12 ´ spp. Hawaii various ? Clausen 1978<br />
Hyperaspis sp. Jamaica Hawaii 1939 Ð Clausen 1978<br />
Scymnus (=Diomus) margipallens Philippines Hawaii 1931 + Clausen 1978<br />
Scymnus (=Nephus) bilucernarius Hawaii<br />
Mexico 1930 +<br />
Lai & Funasaki 1986<br />
Lai & Funasaki 1986<br />
Scymnus pictus Panama 1924 Ð Lai & Funasaki 1986<br />
Scymnus quadrivittatus California 1948 Ð Lai & Funasaki 1986<br />
Scymnus uncinatus Hawaii Mexico,<br />
Panama<br />
1922 + Lai & Funasaki 1986<br />
Scymnus 6 ´ spp. Hawaii various ? Clausen 1978<br />
Scymnus sp. Taiwan Saipan ? Sakimura 1935<br />
150 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong>
Table 4.9.2 (contÕd) Introductions for the biological control <strong>of</strong> Dysmicoccus brevipes<br />
DIPTERA<br />
Country From Year Result Reference<br />
CECIDOMYIIDAE<br />
Cecidomyiid sp. Panama 1931 Ð Lai & Funasaki 1986<br />
Cleodiplosis koebelei Philippines Hawaii 1931 + Clausen 1978<br />
Dicrodiplosis guatemalensis Hawaii Guatemala 1935 + Clausen 1978<br />
Vincentodiplosis (= Lobodiplosis) pseudococci Hawaii Mexico 1930 + Clausen 1978<br />
DROSOPHILIDAE<br />
Pseudiastata nebulosa Hawaii Guatemala 1924<br />
1932<br />
Ð<br />
Ð<br />
Carter 1935<br />
Carter 1935<br />
Pseudiastata pseudococcivora Panama 1931, 1951 Ð Lai & Funasaki 1986<br />
4.9 Dysmicoccus brevipes 151
152 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
BRAZIL<br />
Bisexual D. brevipes is present wherever pineapples are grown, but it is not<br />
recorded whether the parthenogenetic strain also occurs. Mealybug wilt was<br />
rare in 1946 and large mealybug colonies uncommon and always covered<br />
with soil mounds built by Solenopsis sp. ants. Sparse green spotting was<br />
general on the leaves <strong>of</strong> these plants. Natural enemies were numerous and<br />
included Anagyrus sp., Hambletonia pseudococcina and Pseudaphycus sp.<br />
The latter parasitised mealybugs on the aerial parts <strong>of</strong> the pineapple plant, so<br />
that large colonies were rare. It was never found on colonies under the soil<br />
surface. Its life cycle lasts 14 to 20 days and up to 6 individuals may emerge<br />
from a single host. Predators, mainly coccinellids, were generally present<br />
and attacked the underground mealybug colonies (Carter 1949).<br />
COOK IS<br />
Carter (1973) reported that mealybug wilt <strong>of</strong> pineapple was a serious threat<br />
to the newly-developing pineapple industry on Atui and Mangaia.<br />
D. brevipes was present, but not D. neobrevipes, and the mealybug was<br />
attended by Pheidole megacephala.<br />
FIJI<br />
D. brevipes is the main pest on pineapple and causes pineapple wiltÑthe<br />
only record in the Pacific outside Hawaii <strong>of</strong> this condition. It is also a minor<br />
pest <strong>of</strong> sugarcane. It is controlled to some degree by the coccinellids<br />
Cryptolaemus sp., C. montrouzieri and Scymnus sp. (Lever 1945). Three<br />
chrysopids were predators, including Chrysopa ramburi and C. irregularis<br />
(Lever 1940).<br />
GUATEMALA<br />
Two coccinellid predators <strong>of</strong> D. brevipes were reported by Carter (1935),<br />
Scymnus bilucenarius was widespread except in highest elevations, but it<br />
apparently exerted little control; and Brachycantha sp. which was<br />
uncommon. A drosophilid fly, Pseudiastata nebulosa, which was heavily<br />
parasitised by two chalcidid wasps, was found and frequently in large<br />
numbers. It was regarded as a promising species for biological control by<br />
Carter (1935), who introduced it to Hawaii in 1932, but the colony died out.<br />
Cecidomyiid predators were very common in the lowlands and occurred<br />
occasionally in the highlands. They attacked large mealybug colonies on<br />
fruit, but were apparently a minor control factor. No hymenopterous<br />
parasitoids were discovered.<br />
GOLD COAST<br />
The parasitoid Pseudaphycus angelicus was reared on D. brevipes in the<br />
laboratory (Anon. 1953) as also was Anagyrus ananatis from California<br />
(Anon. 1957). Both were released in 1953Ð54 and the former became<br />
established. Scymnus sordidus was also introduced from California, reared<br />
on D. brevipes and released, but establishment is not reported (Anon. 1957).
4.9 Dysmicoccus brevipes 153<br />
INDONESIA<br />
Kalshoven (1981) reported that D. brevipes is attended by the ant,<br />
Monomorium sp.<br />
IVORY COAST<br />
D. brevipes is bisexual with a sex ratio usually <strong>of</strong> 2 males to 1 female. Ants<br />
attending the pineapple mealybug were species <strong>of</strong> Camponotus,<br />
Crematogaster and Pheidole (Re‡l 1959).<br />
JAMAICA<br />
Here and throughout Central America D. brevipes colonies <strong>of</strong> any size were<br />
invariably attended by Solenopsis ants. Where Solenopsis was not present<br />
mealybug colonies were rare and small (Carter 1935).<br />
MALAYSIA<br />
The pineapple mealybug D. brevipes is the most serious insect pest <strong>of</strong><br />
pineapple in peninsular Malaysia. Infected plants become stunted and<br />
reddish and eventually wilt. Fruit are small and unsuitable for canning. In<br />
addition to wilting, the mealybug causes green spotting <strong>of</strong> the leaves, which<br />
is not <strong>of</strong> economic importance (Khoo et al. 1991).<br />
The bisexual form <strong>of</strong> D. brevipes has been studied in some detail by Lim<br />
(1972). It was the only form found in 14 pineapple areas visited in Johore<br />
and Selangor. The bisexual form had a life cycle 10 day shorter than the<br />
parthenogenetic form in Hawaii, although it was less prolific.<br />
MAURITIUS<br />
D. brevipes was first reported as a major pest <strong>of</strong> pineapples in 1933, probably<br />
having been introduced on pineapple suckers from Hawaii about 1931. It<br />
was attacked by three native predators, the coccinellid Scymnus mauritiusi,<br />
the drosophilid fly Gitonides perspicax, and the pineapple scale, Diaspis<br />
bromeliae (Jepson 1939b) but they produced little impact. The coccinellid<br />
Cryptolaemus montrouzieri was introduced from South Africa and liberated,<br />
but did not become established (Moutia and Mamet 1946). The mealybug is<br />
attended by Pheidole megacepahala, Solenopsis geminata and<br />
Technomyrmex detorquens (Jepson and Wieke 1939).<br />
PAPUA NEW GUINEA<br />
D. brevipes is recorded attacking taro where it is subject to predation by the<br />
coccinellids Cryptolaemus affinis, C. montrouzieri and C. wallacii (Shaw et<br />
al. 1979).<br />
PHILIPPINES<br />
Two strains (grey and pink) <strong>of</strong> the pineapple mealybug are present and it is<br />
suggested that they may have been introduced with planting material from<br />
Hawaii, in which case they would represent D. neobrevipes and D. brevipes<br />
respectively. The grey strain produces green spotting <strong>of</strong> the leaves, whereas<br />
the pink strain produces only chlorotic spots. Some pineapple cultivars can<br />
be seriously damaged by pineapple wilt which is caused by both strains. Two
154 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
species <strong>of</strong> ants are almost invariably in attendance on the mealybugs, namely<br />
Solenopsis geminata and Pheidole megacephala (Serrano 1934). Three<br />
predators from Hawaii were established in 1931 (Cleodiplosis koebelei:<br />
Cecidomyiidae, Scymnus margipallens and Hyperaspis silvestri: both<br />
Coccinellidae), but there is no information on their effectiveness (Clausen<br />
1978).<br />
PUERTO RICO<br />
D. brevipes is the most serious pest <strong>of</strong> pineapples, attacking the roots, leaves<br />
and fruits <strong>of</strong> all varieties grown. Anagyrus ananatis was imported in 1936<br />
from Brazil and both A. ananatis and Hambletonia pseudococcina in 1937<br />
from Hawaii where they had been introduced from Brazil and Venezuela<br />
respectively. Both were released in 1937 and 1938. Only half grown or older<br />
hosts were attacked by H. pseudococcina, the life cycle from oviposition to<br />
adult emergence taking 24 to 30 days. In one instance, 3 parasitoids emerged<br />
from the same host. Frequent recoveries were made from the field.<br />
Development from egg to adult took 19 to 21 days for A. ananatis and the sex<br />
ratio was 1:1 (Bartlett 1939). Recoveries <strong>of</strong> this species were reported later<br />
(Bartlett 1943). The larvae <strong>of</strong> a tineid moth, a pyralid moth and <strong>of</strong> a<br />
cecidomyiid fly were found living in the waxy secretions around large<br />
groups <strong>of</strong> mealybugs and were thought to be predators. Three species <strong>of</strong> ants,<br />
including Solenopsis geminata were frequently observed carrying young<br />
mealybugs around. There do not appear to be any recent reports <strong>of</strong> the<br />
effectiveness <strong>of</strong> the introduced parasitoids (Plank and Smith 1940).<br />
SOUTH AFRICA<br />
Larvae <strong>of</strong> the moth Drosica abjectella were observed preying on D. brevipes<br />
on pineapple in the Transvaal. The number <strong>of</strong> moth larvae and pupae per<br />
pineapple plant varied from 1 to 15 and these occurred in the leaf axils. Large<br />
nymphs and fully-fed mealybug females were preferred and moth larvae<br />
each consumed an average <strong>of</strong> 6.5 hosts in the laboratory. In winter,<br />
development <strong>of</strong> fourth instar D. abjectella took 15 to 26 days, the prepupal<br />
period 1 to 3 days and the pupal stage 35 to 45 days. The adults lived for 3 to<br />
6 days (BŸttiker 1957).<br />
SRI LANKA<br />
Both a bisexual and a parthenogenetic strain <strong>of</strong> D. brevipes are present. The<br />
former, which causes green spotting <strong>of</strong> pineapple leaves, occurs in the west<br />
and the latter in the Bibile area (Carter 1956).<br />
TAIWAN<br />
D. brevipes is widely distributed on pineapple up to about 750 m and also<br />
occurs on banana (Chiu and Cheng 1957). It appears that D. neobrevipes is<br />
also present. Natural enemies include the cecidomyiid predator<br />
Schizobremia formosana and also coccinellids, but these are less effective
TRINIDAD<br />
USA<br />
4.9 Dysmicoccus brevipes 155<br />
(Takahashi 1939). Cryptolaemus, imported for control <strong>of</strong> D. brevipes and<br />
other mealybugs, was only partially effective. A Scymnus sp., said to be<br />
effective against D. brevipes in Saipan, was introduced, but no further<br />
information is available (Sakimura 1935). The most abundant attendant ants<br />
were Pheidologeton diversus, Anoplolepis longipes and Camponotus<br />
friedae (Lee 1974).<br />
The encyrtid Pseudaphycus dysmicocci was reared as a solitary parasitoid <strong>of</strong><br />
second instar female nymphs <strong>of</strong> D. brevipes on pineapple (Bennett 1955).<br />
D. brevipes was a common pest in southern Florida and Hambletonia<br />
pseudococcina was introduced from Puerto Rico and liberated in 1954.<br />
Although it became established, information on its abundance and<br />
effectiveness is not available. It was postulated that the widespread use <strong>of</strong><br />
organic pesticides had probably reduced the parasitoid to very low levels<br />
(Clausen 1956).<br />
Major natural enemies<br />
Anagyrus ananatis Hym.: Encyrtidae<br />
This moderately polyphagous wasp is widespread in Brazil, where it is<br />
known as a parasitoid <strong>of</strong> D. brevipes, but also parasitises, inter alia, rhodes<br />
grass scale (Antonina graminis) and citrus mealybug (Planococcus citri)<br />
(Gabriel et al. 1982). It was established in Hawaii (Carter 1937), where it<br />
completes a generation in about 20 days.<br />
Hambletonia pseudococcina Hym.: Encyrtidae<br />
This parasitoid occurs as a bisexual form on D. brevipes in Brazil and as a<br />
parthenogenetic one in Colombia and Venezuela. The bisexual form failed<br />
to reproduce on D. brevipes in Hawaii, but the parthenogenetic form did so<br />
successfully (Carter 1937). About 24 to 30 days is required for the life cycle<br />
under tropical outdoor conditions and up to 4 individuals may emerge from a<br />
single host (Compere 1936). In laboratory trials, H. pseudococcina showed<br />
a high degree <strong>of</strong> specificity for D. brevipes and did not oviposit in 8 closely<br />
related mealybug species. Of 3 additional mealybugs tested, it attempted<br />
oviposition only in an unidentified species from a grass (Clancy and Pollard<br />
1947).<br />
Pseudiastata nebulosa Dipt.: Drosophilidae<br />
This predator is native to Guatemala. It was introduced to Hawaii in 1924 but<br />
did not become established. Except in the highlands <strong>of</strong> Guatemala, it is<br />
frequently found in large numbers on a single plant, both above and below<br />
the soil line. It was again introduced in 1932, but the colony died out. This
156 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
was possibly due to its requirement for large numbers <strong>of</strong> hosts, since 7 larvae<br />
consumed over 100 medium to large sized hosts during the last half <strong>of</strong> their<br />
larval lives. In Guatemala it is highly parasitised by two species <strong>of</strong> chalcidid<br />
wasps. Carter (1935) regarded it as a promising species for biological<br />
control.<br />
Vincentodiplosis pseudococci Dipt.: Cecidomyiidae<br />
This midge is native to Mexico and was established in Hawaii in 1950. In<br />
neglected, weedy pineapple plantations in Hawaii, where the ant Pheidole<br />
sp. was less reliant on mealybug honeydew, the midge was sufficiently<br />
effective in controlling D. brevipes as to almost completely eliminate the<br />
mealybug from the fruit. Many fruit were covered with the old webs<br />
produced by midge larvae, but there were no live mealybugs. The midge is<br />
rarely found on leaves, but its larvae are commonly found attacking large<br />
mealybugs at the base <strong>of</strong> the fruit (Carter 1935, 1944).<br />
Comments<br />
It is very likely that D. brevipes evolved in South and/or Central America<br />
and there is, therefore, a prima facie case to consider it as a candidate for<br />
classical biological control in <strong>Southeast</strong> Asia and the Pacific. Indeed, there<br />
are 2 parasitoid species (Anagyrus ananatis and Hambletonia<br />
pseudococcina: both Eulophidae) and 2 predator species (Vicentodiplosis<br />
pseudococci: Cecidomyiidae and Pseudiasta nebulosa: Diastadidae) that are<br />
capable <strong>of</strong> reducing the mealybug to subeconomic levels. However, when<br />
any one or more <strong>of</strong> a number <strong>of</strong> ant species attends the mealybug it is largely<br />
protected from natural enemies and is able to build up to damaging numbers.<br />
In the absence <strong>of</strong> both ants and natural enemies the unharvested honeydew it<br />
produces leads to heavy growth <strong>of</strong> sooty moulds and there is transmission <strong>of</strong><br />
pineapple mealybug wilt. Since apparently suitable natural enemies are<br />
available for introduction, the key to D. brevipes control is to deal with the<br />
attendant ants. There are several ant baits that have been used successfully<br />
for this purpose, but these are no longer registered for use in USA and,<br />
hence, cannot be recommended. No doubt suitable replacements will soon<br />
emerge. When extensive plantings <strong>of</strong> pineapples are made on areas where<br />
the soil has been worked to kill weeds, very few ant colonies survive.<br />
Recolonisation <strong>of</strong> the planted area occurs as colonies move along the rows<br />
towards the centre <strong>of</strong> the crop. One cultural method recommended to delay<br />
this spread is to plant several peripheral rows parallel to each boundary. Ants<br />
will then move along these, rather than into the crop and control measures<br />
can be concentrated on these rows (Rohrback et al. 1988). Promising results<br />
obtained with the integrated management <strong>of</strong> D. brevipes in Hawaii suggest<br />
that it would be well worth exploring similar methods elsewhere.
4.10 Hypothenemus hampei<br />
India<br />
20°<br />
0°<br />
20°<br />
Myanmar<br />
Laos<br />
+<br />
China<br />
Thailand<br />
+<br />
Cambodia<br />
P<br />
Vietnam<br />
++<br />
+<br />
++ Brunei<br />
Malaysia<br />
Singapore<br />
++<br />
Indonesia<br />
Taiwan<br />
+++<br />
Philippines<br />
Australia<br />
P<br />
Papua<br />
New Guinea<br />
157<br />
Hypothenemus hampei is native to Central Africa but has spread to most c<strong>of</strong>fee<br />
producing countries in Central and South America, to <strong>Southeast</strong> Asia and to several<br />
Pacific countries. Significant c<strong>of</strong>fee-growing areas not yet infested are Hawaii, Papua New<br />
Guinea, Vanuatu and Solomon Islands.<br />
It is a pest exclusively <strong>of</strong> c<strong>of</strong>fee berries and does not damage the vegetative parts. It is<br />
difficult to control with chemicals and, although plantation management methods can<br />
reduce damage, the c<strong>of</strong>fee berry borer remains an important pest.<br />
The most important natural enemies appear to be 3 parasitic wasps native to Africa,<br />
Cephalonomia stephanoderis,<br />
Phymastichus c<strong>of</strong>fea and Prorops nasuta.<br />
The last <strong>of</strong> these<br />
has been established in Brazil and Colombia without its own natural enemies, but has not<br />
so far produced spectacular results. C. stephanoderis has been established recently in<br />
Colombia, Ecuador, Mexico and New Caledonia, but it is too early to evaluate its impact.<br />
Phymastichus c<strong>of</strong>fea has not yet been established anywhere, but this is foreshadowed in<br />
Colombia. The fungus Beauveria bassiana shows early promise. A thorough study is in<br />
progress <strong>of</strong> the interactions <strong>of</strong> the parasites and other natural enemies <strong>of</strong> H. hampei and<br />
the influence on them <strong>of</strong> various components <strong>of</strong> the environment. Optimism has been<br />
expressed about the outcome <strong>of</strong> this program.<br />
20°<br />
0°<br />
20°
158 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Hypothenemus hampei (Ferrari)<br />
Rating<br />
Origin<br />
Distribution<br />
Coleoptera: Scolytidae<br />
c<strong>of</strong>fee berry borer<br />
<strong>Southeast</strong> Asia Southern and Western Pacific<br />
+++ Phil +++ N Cal<br />
12 ++ Viet, Msia, Indo 7 ++ Fiji, Fr P<br />
+ Thai, Laos, Brun<br />
P Camb P Pohnpei, Saipan<br />
This account updates the chapter on H. hampei in Waterhouse and Norris<br />
(1989) and the valuable review <strong>of</strong> Murphy and Moore (1990) in relation to<br />
prospects for biological control.<br />
FerrariÕs specimens, described in 1867 under the generic name<br />
Stephanoderes,<br />
were obtained from trade c<strong>of</strong>fee beans in France. There<br />
appears to be no record <strong>of</strong> the country <strong>of</strong> origin <strong>of</strong> the material, but in 1867<br />
infested beans could only have come from Africa or Saudi Arabia, because<br />
Hypothenemus hampei did not obtain a footing on other continents until<br />
later. The seed used to establish C<strong>of</strong>fea arabica in Saudi Arabia was<br />
probably obtained from the Ethiopian highlands centuries ago. There the<br />
c<strong>of</strong>fee berry borer is native, though scarce (Davidson 1967), but if it did not<br />
accompany the original seed it could easily have reached Saudi Arabia<br />
through Arabian-African commerce over the centuries.<br />
The wider range <strong>of</strong> parasitoids (3) in West Africa than in East Africa (2,<br />
with one shared with the West) suggests that H. hampei has been in the West<br />
for a very long time and may indeed have evolved there (L.O. Brun pers.<br />
comm.).<br />
This was given by CIE (1981) as: Africa (Angola, Benin, Burundi,<br />
Cameroon, Canary Is, Central African Republic, Chad, Congo, Ethiopia,<br />
Fernando Poo, Gabon, Ghana, Guinea, Ivory Coast, Kenya, Liberia,<br />
Malawi, Mozambique, Nigeria, Principe, Rio Muni, Rwanda, S‹o TomŽ,<br />
Senegal, Sierra Leone, Sudan, Tanzania, Togo, Uganda and Zimbabwe);<br />
Middle East (Saudi Arabia), Asia (Indonesia, Cambodia, Laos, Malaysia,<br />
Philippines, Sri Lanka, Thailand, Vietnam); Central America (Guatemala,
Biology<br />
4.10<br />
Hypothenemus hampei<br />
159<br />
Honduras, Greater West Indies); South America (Brazil, Peru, Surinam);<br />
Pacific (Caroline Is, Irian Jaya, Marianas Is, New Caledonia, Society Is).<br />
To these must be added: South America (Colombia in 1988 (D. Moore<br />
pers. comm. 1989), Ecuador (CIBC 1988a, b)); Central America (El<br />
Salvador, Mexico (Baker 1984)); Asia (India (Kumar et al. 1990)) and the<br />
Pacific (Fiji (Anon. 1979a), Tahiti (Johnston 1963)). In the West Indies,<br />
Reid (1983) reported the beetle from Jamaica and Puerto Rico, but it has not<br />
been reported from the lesser West Indies (Guadeloupe).<br />
Significant c<strong>of</strong>fee-growing or potential c<strong>of</strong>fee-growing areas not yet<br />
infested are Solomon Is, Vanuatu, Hawaii and Papua New Guinea, although<br />
the latter is at serious risk because it shares a common land frontier with Irian<br />
Jaya (Indonesia), where H. hampei has been present for many years<br />
(Thomas 1961). H. hampei is not present in Australia.<br />
The following description <strong>of</strong> the life cycle refers exclusively to the<br />
relationship <strong>of</strong> the beetle with C<strong>of</strong>fea spp., and principally with Arabian<br />
c<strong>of</strong>fee C. arabica and robusta c<strong>of</strong>fee C. canephora,<br />
the most important<br />
cultivated species. Infestations <strong>of</strong> H. hampei occur in c<strong>of</strong>fee seeds while<br />
they are enclosed in berries on the trees and in berries that fall to the ground.<br />
They will also continue vigorously in processed beans in storage, but not if<br />
the moisture content has been reduced below 12.5% (robusta beans) or<br />
13.5% (arabica beans) (Hargreaves 1935). Apart from dispersive flight by<br />
adult females and the walking by males from one berry to another on the<br />
same branch (P. Cochereau pers. comm. 1995), no part <strong>of</strong> the life cycle <strong>of</strong><br />
the c<strong>of</strong>fee berry borer is passed through outside <strong>of</strong> the c<strong>of</strong>fee bean.<br />
The length <strong>of</strong> adult females <strong>of</strong> H. hampei <strong>of</strong> American origin is given as<br />
1.4 to 1.7 mm (Wood 1982) and <strong>of</strong> Ugandan females and males as about<br />
1.9 mm and 1.3 mm respectively (Hargreaves 1926). Malaysian females<br />
averaged 1.58 mm and males 0.99 mm (Corbett 1933). Females outnumber<br />
males by at least 10 to 1 and the ratio is frequently much higher. The beetle is<br />
brown when it first emerges from the pupa but in the course <strong>of</strong> 4 or 5 days it<br />
becomes generally black, although the prothorax has a slightly reddish tinge.<br />
The prothorax is markedly humped, so that the down-turned head is not<br />
visible from above. The tibiae have strong spines which doubtless assist in<br />
such activities as tunneling through the pulp <strong>of</strong> c<strong>of</strong>fee berries, ejecting the<br />
resulting frass, and forcing a way to the soil surface should the berry become<br />
interred.<br />
Beetle attack tends to be aggregated on some trees or on particular<br />
branches within trees, rather than evenly distributed (Baker 1984). The
160 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
fertilised female flies to c<strong>of</strong>fee berries that have begun to ripen and bores an<br />
entrance hole at the apex, either in the terminal pore or in the calyx ridge or<br />
annulus <strong>of</strong> differentiated tissue that surrounds the pore. Sometimes this<br />
annulus is perforated by several holes, but boring into the fruit elsewhere is<br />
unusual. The colour <strong>of</strong> berries appears not to influence choice by females<br />
seeking oviposition sites (Morallo-Rejesus and Baldos 1980). Young<br />
berries, containing seeds with a watery endosperm, usually do not come<br />
under attack if more advanced berries are plentiful. If they do they are soon<br />
abandoned, after the female has fed on some <strong>of</strong> the pulp, and they then tend<br />
to fall prematurely, being particularly vulnerable to infection by disease<br />
organisms. The falling <strong>of</strong> such immature berries after being attacked <strong>of</strong>ten<br />
contributes significantly to the amount <strong>of</strong> crop lost. After the endosperm has<br />
passed from the watery to the milky stage in the course <strong>of</strong> maturation, beetles<br />
invading the berry will wait in the pulp until the seed tissue is firm enough to<br />
excavate (Penatos and Ochoa 1979). Rhodes and Mansingh (1981) cite<br />
opinions to the effect that females that become static in this fashion for<br />
several weeks (May to mid-July in the Jamaican lowlands) are in a state <strong>of</strong><br />
reproductive diapause. When available, berries are selected that are already<br />
suitable for colonisation. The green berry is favoured for feeding and the ripe<br />
(i.e. red) berry for breeding purposes, but the ripe berries are also very<br />
suitable for feeding (Corbett 1933). In a ripe berry the female bores in one<br />
operation through skin, pulp and the endocarp and pellicle surrounding one<br />
<strong>of</strong> the two seeds (beans) present in each berry. Ejected frass may surround<br />
the entrance hole during boring (Hutson 1936). Several days may be<br />
occupied in this boring process, and the female then tunnels into the<br />
endosperm, the substance <strong>of</strong> the seed, which is the basis <strong>of</strong> the worldÕs US$8<br />
billion annual c<strong>of</strong>fee crop (Bardner 1978). Berries that fall to the ground<br />
may generate considerable numbers <strong>of</strong> beetles, but these are from their ontree<br />
infestation, since the female berry borers do not appear to visit fallen<br />
fruit (Baker 1984).<br />
The eggs are laid at the rate <strong>of</strong> two or three a day in batches <strong>of</strong> 8 to 12 in<br />
chambers chewed out <strong>of</strong> the maturing bean tissue. Oviposition extends over<br />
a period <strong>of</strong> three to seven weeks, each female producing from about 30 to<br />
over 70 eggs. According to some authors, laying is not necessarily confined<br />
to one bean because the female that has initiated an infestation may fly to<br />
other berries during the oviposition period. According to others (e.g.<br />
Bergamin 1943) the female that has initiated an infestation only quits the<br />
bean when the first <strong>of</strong> her progeny emerge as adults. Others again (e.g.<br />
Hargreaves 1935) state that she remains until all the bean tissue is consumed<br />
or has deteriorated in some way. Most likely the pattern is quite flexible.<br />
Eggs hatch in three to nine days and young larvae bore into intact bean
4.10<br />
Hypothenemus hampei<br />
161<br />
tissue, making pockets opening <strong>of</strong>f the main tunnel made by the parent<br />
female. Male larvae pass through their two instars in the course <strong>of</strong> about 15<br />
days, and the females pass through three instars in about 19 days (Bergamin<br />
1943). Morallo-Rejesus and Baldos (1980) state that the female, like the<br />
male, passes through only two instars, indicating the need for further<br />
biological study. The long period over which oviposition is spread results in<br />
larvae in all stages <strong>of</strong> development being present in one bean. At the end <strong>of</strong><br />
the larval stage there is a non-feeding or prepupal stage lasting about two<br />
days. The insect then pupates, without any cocoon formation, in the galleries<br />
excavated by the larvae. The pupal stage is passed through in four to nine<br />
days. The period from egg-laying to the emergence <strong>of</strong> the adult is 25 to 35<br />
days. The temperatures at which the preceding records were made are<br />
generally not specified, but chiefly they relate to warm lowland c<strong>of</strong>fee<br />
plantations. Bergamin (1943) recorded that at 24.5¡C in Brazil the period<br />
from egg-laying to emergence <strong>of</strong> adult averaged 27.5 days. De Oliveira<br />
Filho (1927) found that in Brazil shade temperatures <strong>of</strong> 20 to 30¡C suited the<br />
females best. Below 15¡C they became inactive, endeavouring to hide,<br />
preferably in c<strong>of</strong>fee berries, but sometimes by boring into beans, maize,<br />
peanuts or cotton seed <strong>of</strong> suitably low moisture content. They can survive<br />
temperatures just below 0¡C, which however are rarely experienced in<br />
Brazilian c<strong>of</strong>fee growing areas. At higher elevations development is<br />
somewhat prolonged (Le Pelley 1968) and H. hampei has a low pest status in<br />
highland c<strong>of</strong>fee growing areas in East Africa and Java (Haarer 1962). Baker<br />
et al. (1989) conclude that the optimum mean annual temperature for the<br />
beetle is 23¡ to 25¡C and that parasitoids for biological control should be<br />
sought from a similar climate.<br />
The adult males emerge from the pupa earlier than the females. Their<br />
hindwings are short and they do not fly, but remain in the bean, fertilising<br />
their female siblings as they emerge. Each male can fertilise two females a<br />
day and up to 30 in his lifetime which may extend to 103 days, although<br />
averaging less. Corbett (1933) states that the males seldom leave the berries,<br />
and then only when they are near death. The vast majority <strong>of</strong> observers<br />
confirm that males never leave the berries. Quite likely they may move from<br />
bean to bean within a fruit, thereby gaining access to females other than their<br />
sisters. Parthenogenesis does not occur and, although unfertilised females<br />
may produce some eggs, these do not hatch. One insemination is sufficient to<br />
allow a female to lay fertile eggs throughout her reproductive period.<br />
Corbett (1933) stated that, if there are no males in the seed when the females<br />
emerge from the pupal skin after their hardening period <strong>of</strong> a few days, they<br />
leave via the entrance hole and seek males in other infested berries. Morallo-<br />
Rejesus and Baldos (1980) suggest that sex pheromones secreted by the<br />
males guide such females to appropriate berries.
162 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Females that have been fertilised remain in the ÔparentalÕ bean for three<br />
or four days, by which time they have become sexually mature. They then<br />
leave the berries via the entrance holes and enter others and, after a<br />
preoviposition period <strong>of</strong> 4 to 20 days, commence egg laying. Females have<br />
been known to live up to 282 days, and longevity was stated by Bergamin<br />
(1943) to average 156 days. According to Corbett (1933), in Malaysia<br />
females survived 81 days without food. There is time for a succession <strong>of</strong><br />
seven or eight generations a year in lowland c<strong>of</strong>fee growing areas but, on<br />
account <strong>of</strong> the long reproductive period, there are few clearcut population<br />
peaks to indicate generations.<br />
Life history studies have been carried out with artificial infestations <strong>of</strong><br />
c<strong>of</strong>fee trees in southern Mexico (Baker et al. 1992). Morallo-Rejesus and<br />
Baldos (1980) observed in the Philippines that beetles are to be observed in<br />
flight from 3.00 pm, considerable numbers being visible in the air between<br />
4.00 and 5.00 pm. Corbett (1933) observed in Malaysia that females fly at<br />
any time during the day, but in greatest numbers between 2.00 and 5.00 pm,<br />
reaching a peak between 3.30 and 4.30 pm. De Oliveira Filho (1927) states<br />
that, in Brazil, females Ôare activeÕ on warm nights, but it is unclear whether<br />
this implies flight activity. Kalshoven (1981) states that, in Java, females<br />
start flying during the midday period, and that they assemble under leaves<br />
and in other places where they dance up and down like gnats. Such activity<br />
can have no sexual significance, seeing that the males do not leave the seeds,<br />
and its function is obscure. In Java flights up to 345 m have been measured<br />
(Leefmans 1920). In Mexico, Baker (1984) carried out experimental studies<br />
on flight. Females flew freely in the laboratory for up to 22 minutes, tending<br />
to hover or move forward only slowly. In tethered flight, and thus relieved <strong>of</strong><br />
supporting their own weight, they could fly non-stop for 100 minutes, with a<br />
combined aggregate <strong>of</strong> three hours. Such enduring activity, combined with<br />
its afternoon peak <strong>of</strong> activity, suggests that, in their habits, the beetles<br />
resemble aphids and thrips in being adapted to exploiting periods <strong>of</strong><br />
maximum convection in the atmosphere, so achieving long-distance travel<br />
with their own contribution serving chiefly to keep them al<strong>of</strong>t. De Oliveira<br />
Filho (1927) states that local flight occurs when the fertilised female is<br />
seeking a place to lay, when (oviposition having commenced) she emerges<br />
to seek moister berries after having been driven out by the heat <strong>of</strong> the sun. It<br />
also occurs when unfertilised females seek males (as they do if there are<br />
none in the berry when they emerge), when seeds are waterlogged, are<br />
overcrowded with adults and larvae, or when the beetles are disturbed.<br />
Rhodes and Mansingh (1981) state that, in the Jamaican lowlands,<br />
beetles in dry berries remain in diapause for five months, from mid-<br />
December to mid-May. Baker (1984) found that in mid-spring in Mexico
Host plants<br />
4.10<br />
Hypothenemus hampei<br />
163<br />
females tended to remain in fallen c<strong>of</strong>fee berries at a time when temperatures<br />
in berries in the trees ranged up to an inimical 37¡C. Soaking the fallen<br />
berries in water induced many to emerge, but they did so in a specific<br />
pattern, some seven to eight hours after dawn. Possibly the soaking<br />
simulated rain that would have made the environment generally more<br />
favourable. Baker reminds us that c<strong>of</strong>fee is naturally an understorey plant in<br />
tropical forest and, by sheltering in fallen berries, beetles may avoid the<br />
harmful effects <strong>of</strong> strong, direct sunlight. Infestations are carried over<br />
between peaks <strong>of</strong> fruiting by the breeding that occurs in late-maturing<br />
berries, or else in those that have fallen to the ground. Females can survive<br />
for up to two months in buried beans (Clausen 1978).<br />
It is probable that intercontinental travel is brought about by the agency<br />
<strong>of</strong> man, rather than by travel in moving air masses. Infested beans are an<br />
obvious vehicle for dispersal, but there are other avenues to which<br />
quarantine measures should be applied. In Jamaica, Reid (1983) observed<br />
females among banana trash used in packing boxes on their way to the<br />
boxing plant. Commonly, beetles disperse in sacks, empty or otherwise, and<br />
on the clothing and equipment <strong>of</strong> plantation workers. Under some conditions<br />
beetles bore for protection into wood or other materials to the extent that<br />
Baker (1984) suggested that authorities in beetle-free areas should think<br />
very carefully before allowing entry <strong>of</strong> untreated plant material from an<br />
infested area.<br />
An important aspect <strong>of</strong> the biology <strong>of</strong> any insect pest is its host range. In<br />
Africa, in addition to its regular hosts in the genus C<strong>of</strong>fea,<br />
Hypothenemus<br />
hampei has been reported from fruit, pods or seeds <strong>of</strong> species <strong>of</strong> Centrosema,<br />
Crotalaria,<br />
Phaseolus and Tephrosia (Fabaceae), Leucaena (Mimosaceae),<br />
Caesalpinia (Caesalpiniaceae), Hibiscus (Malvaceae), Rubus and<br />
Oxyanthus (Rubiaceae), Vitis (Vitaceae) and Ligustrum (Oleaceae), but<br />
these associations are all considered to reflect only casual feeding by adults.<br />
In Africa, the only species outside <strong>of</strong> the genus C<strong>of</strong>fea in which immature<br />
stages have been found is Dialium lacourtianum (Caesalpiniaceae) (Le<br />
Pelley 1968).<br />
A review <strong>of</strong> hosts <strong>of</strong> the genus Hypothenemus was made by Johanneson<br />
and Mansingh (1984) who concluded that H. hampei was monophagous<br />
according to their criteria, as it attacked only six species <strong>of</strong> the genus C<strong>of</strong>fea.<br />
However, they listed 23 other species <strong>of</strong> plants in 11 families from which<br />
H. hampei has been recorded, but only as adult females. In contrast, in the<br />
Philippines, Morallo-Rejesus and Baldos (1980), whose paper was
164 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Damage<br />
overlooked by Johanneson and Mansingh, reported finding eggs, larvae and<br />
pupae <strong>of</strong> H. hampei in Leucaena leucocephala (Mimosaceae), Gliricidia<br />
sepium (Fabaceae), two species <strong>of</strong> Psychotria (Rubiaceae) and one <strong>of</strong><br />
Dioscorea (Dioscoreaceae). In laboratory tests they found that adults <strong>of</strong><br />
H. hampei fed on pods <strong>of</strong> four <strong>of</strong> those species and also on the pods <strong>of</strong> 19<br />
other species in 9 orders.<br />
Such feeding tests may be <strong>of</strong> little significance, however, since the<br />
survival times recorded are greatly exceeded by the periods for which the<br />
beetles are capable <strong>of</strong> withstanding starvation (Corbett 1933). If the insects<br />
were correctly identified, the host plants recorded in the Philippines may<br />
help to support a population <strong>of</strong> H. hampei when no c<strong>of</strong>fee berries are<br />
available. Reexamination <strong>of</strong> the host range is necessary. For example Cohic<br />
(1958) found H. hampei attacking loquat in New Caledonia, and this<br />
relationship, though abortive in the end, has not been reported anywhere else<br />
in the world. In connection with host records, Johanneson and Mansingh<br />
(1984) drew attention to the problem <strong>of</strong> misidentification <strong>of</strong> species <strong>of</strong><br />
Hypothenemus,<br />
a notoriously difficult genus, and also to misinterpretation<br />
<strong>of</strong> the relative roles <strong>of</strong> various host plants. Hargreaves (1935) found adults <strong>of</strong><br />
four species <strong>of</strong> Hypothenemus other than H. hampei in seed <strong>of</strong> Phaseolus<br />
lunatus (Fabaceae) in Uganda, and Gonzalez (1978) alludes to species <strong>of</strong><br />
Hypothenemus,<br />
known as false c<strong>of</strong>fee borers, which occur from Mexico to<br />
northern Argentina and greatly complicate quarantine procedures. Such<br />
insects would, <strong>of</strong> course, also raise difficulties in host plant studies. A<br />
thorough review <strong>of</strong> true hosts <strong>of</strong> H. hampei would be relevant to a number <strong>of</strong><br />
aspects <strong>of</strong> the control <strong>of</strong> this pest.<br />
Hypothenemus hampei is a pest exclusively <strong>of</strong> the immature and mature<br />
c<strong>of</strong>fee berries and does no damage whatsoever to the vegetative parts <strong>of</strong> the<br />
plant. Prates (1969) showed that adults <strong>of</strong> H. hampei were strongly attracted<br />
to extracts <strong>of</strong> green or ripe c<strong>of</strong>fee berries, but not to extracts <strong>of</strong> c<strong>of</strong>fee leaves<br />
or flowers. Significant losses are caused by the female beetles feeding on<br />
young berries which are too immature to colonise but which, after the beetle<br />
has gone, are invaded by decay organisms, and so fall prematurely. In Java<br />
Leefmans (1920) found that 80% <strong>of</strong> green berries that had fallen through<br />
being bored by the beetle contained decayed beans as against 46.5% in<br />
unbored beans that had fallen through other causes. In the Congo, Schmitz<br />
and Crisinel (1957) found that 64 to 82% <strong>of</strong> shed berries had fallen on<br />
account <strong>of</strong> H. hampei attack. Such losses caused by attack on immature<br />
fruits are serious enough, but the bulk <strong>of</strong> the damage done by this beetle is to
4.10<br />
Hypothenemus hampei<br />
165<br />
the endosperm <strong>of</strong> the mature beans, which may be extensively damaged or<br />
even completely destroyed. Even lightly bored beans acquire a distinctive<br />
blue-green staining which significantly reduces their market value (McNutt<br />
1975), but the further tunnelling by the beetles and their larvae brings about<br />
progressive degradation, so that the c<strong>of</strong>fee bean is reduced to a mass <strong>of</strong> frass.<br />
Market requirements demand the removal <strong>of</strong> damaged berries from the<br />
harvested crop, which is done by various mechanical processes (fortunately<br />
bored beans float), supplemented even by handpicking. The beans removed<br />
by such processing are not necessarily a total loss, but can go into only low<br />
grade fractions at a much reduced market rate.<br />
In New Caledonia, where no control measures had been implemented,<br />
H. hampei was found to have attacked 80% <strong>of</strong> berries (Cohic 1958). Other<br />
examples <strong>of</strong> losses due to Hypothenemus hampei are given by Le Pelley<br />
(1968). Severe infestations in Uganda may result in 80% <strong>of</strong> berries being<br />
attacked. In the Ivory Coast, damage <strong>of</strong> 5% to 20% <strong>of</strong> berries is common,<br />
rising to 50% to 80% in some cases. In the Congo, boring <strong>of</strong> up to 84% <strong>of</strong><br />
green berries and up to 96% <strong>of</strong> hard berries has been recorded and, in<br />
Tanzania, records indicate up to 96% boring <strong>of</strong> hard berries. In Malaysia<br />
there have been records <strong>of</strong> up to 90% <strong>of</strong> beans damaged. In Java crop loss <strong>of</strong><br />
40% was recorded in 1929, and in Brazil 60% to 80% losses have been<br />
experienced. The above figures apply for the most part to poorly managed<br />
situations, and crop losses can be reduced by appropriate management, but<br />
the beetle is a constant latent threat if vigilance is relaxed. In Jamaica, Reid<br />
(1983) estimated that 27% <strong>of</strong> the berries harvested were damaged. The<br />
studies <strong>of</strong> Reid and Mansingh (1985) showed that H. hampei was<br />
responsible for 20.9% reduction <strong>of</strong> exportable beans in the Jamaican crop <strong>of</strong><br />
1980Ð81. Baker (1984) reported that, in southern Mexico, the attack <strong>of</strong><br />
H. hampei on c<strong>of</strong>fee plantations was so severe that, in spite <strong>of</strong> application <strong>of</strong><br />
insecticides in some places in 1982, no berries were harvested because it<br />
would not have been economical to do so.<br />
Proper processing results in beans <strong>of</strong> moisture content too low to permit<br />
the borer to multiply. This is below 13.5% for arabica c<strong>of</strong>fee and below<br />
12.5% for robusta c<strong>of</strong>fee. If c<strong>of</strong>fee beans are stored with significantly higher<br />
moisture content, beetle reproduction continues. Thus Morallo-Rejesus and<br />
Baldos (1980) found that, in the Philippines, infestation in c<strong>of</strong>fee beans<br />
stored before drying rose from 20% to 100% in six weeks.
166 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Natural enemies<br />
The cryptic nature <strong>of</strong> the immature stages and the male <strong>of</strong> H. hampei makes<br />
them relatively inaccessible victims for predators, and the only one recorded<br />
is the non-specific Javanese bug Dindymus rubiginosus. This bug draws the<br />
borers from the berries with its beak and sucks them dry. Le Pelley (1968)<br />
states that it is <strong>of</strong> little importance.<br />
The most important parasitic wasps, Cephalonomia stephanoderis,<br />
Prorops nasuta, Phymastichus c<strong>of</strong>fea and Heterospilus c<strong>of</strong>feicola are, <strong>of</strong><br />
course, African in origin and are dealt with in some detail by Klein Koch et<br />
al. (1988) and Feldhege (1992). C. stephanoderis which is restricted to West<br />
Africa is the most important species in Ivory Coast, parasitising up to 50% <strong>of</strong><br />
H. hampei in black berries (Ticheler 1961). The potential <strong>of</strong> H. c<strong>of</strong>feicola in<br />
biological control requires further study because its larvae are not very<br />
specific, but the other three species appear to have a narrow enough host<br />
range to make them acceptable from this point <strong>of</strong> view. A fifth parasite,<br />
Goniozus sp. is recorded, but without further data, from Ivory Coast<br />
(Cochereau and Potiaroa 1994).<br />
In addition to the identified arthropod natural enemies (Table 4.10.1),<br />
Leefmans (1924a) recorded a non-specific parasite that attacks beetles in<br />
newly infested berries and Hargreaves (1926) found an unidentified<br />
hymenopterous parasitoid in Uganda, now known also from Togo as<br />
Aphanogmus dictynna and considered to be a hyperparasitoid <strong>of</strong><br />
C. stephanoderis or P. nasuta (Feldhege 1992). Morallo-Rejesus and<br />
Baldos (1980) reported the presence in the Philippines <strong>of</strong> a braconid and an<br />
encyrtid parasitoid <strong>of</strong> H. hampei, both unidentified, and presumably nonspecific<br />
members <strong>of</strong> the local fauna.<br />
Some ants attack the borer. Swallows and other small birds that feed on<br />
the wing consume flying adults <strong>of</strong> H. hampei.<br />
The parasitic fungus Beauveria bassiana has been observed attacking<br />
H. hampei in Brazil (Averna-Sacc‡ 1930; Villacorta 1984), Jamaica<br />
(Rhodes and Mansingh 1981), Cameroon (Pascalet 1939), Congo (Sladden<br />
1934; Steyaert 1935), Ivory Coast (Ticheler 1961), India (Balakrishnan et al.<br />
1994), Java (Friederichs and Bally 1922) and in New Caledonia (Cochereau<br />
and Potiaroa 1994). Steyaert (1935) and Averna-Sacc‡ (1930) studied the<br />
seasonal cycle and the former also made studies <strong>of</strong> the infectivity and<br />
epidemiology <strong>of</strong> the fungus <strong>of</strong> which there are many strains. In an analysis <strong>of</strong><br />
16 isolates from H. hampei adults from 10 countries in Latin America,<br />
Africa, Asia and the Pacific, 13 formed a homogenous group with very<br />
similar electrophoretic and physiological characteristics, suggesting a<br />
distinct strain associated widely with the c<strong>of</strong>fee berry borer. Of the
4.10 Hypothenemus hampei 167<br />
remaining 3 strains, one from Sri Lanka is suspected as having degenerated<br />
during some 63 years in storage, but the others (from New Caledonia and<br />
Kenya) are probably distinct entities (Bridge et al. 1990). The New<br />
Caledonian strain presumably attacked some other host until H. hampei<br />
arrived there in 1948. It is a particularly virulent strain and can cause death <strong>of</strong><br />
H. hampei in 5 days (Cochereau et al. 1994). Moist, warm conditions favour<br />
the incidence <strong>of</strong> this pathogen, and heavy rain is thought to enhance the rate<br />
<strong>of</strong> infection. If spraying with fungal preparations is avoided on the day <strong>of</strong><br />
release <strong>of</strong> parasitoids, adverse effects on the latter are not observed (Reyes et<br />
al. 1995). Friederichs (1922) recommended the encouragement <strong>of</strong> heavy<br />
shade to increase the incidence <strong>of</strong> fungal pathogens, but this runs counter to<br />
the fact that intensity <strong>of</strong> shade must <strong>of</strong>ten be reduced to encourage<br />
hymenopterous parasitoids which, however, may still prove to be <strong>of</strong> minor<br />
significance in population regulation. Certainly, Klein Koch (1989a)<br />
considered Beauveria to be the most important natural enemy <strong>of</strong> H. hampei<br />
in Ecuador. In Colombia, preparations <strong>of</strong> selected strains <strong>of</strong> Beauveria in oil<br />
have produced 20 to 95% adult mortality, slightly higher than the 20 to 90%<br />
produced by selected strains <strong>of</strong> Metarhizium (P. Cochereau, pers. comm.<br />
1995). Varela and Morales (1996) have characterised a number <strong>of</strong> Beauveria<br />
isolates and their virulence against H. hampei.<br />
Pascalet (1939) advocated the spraying <strong>of</strong> suspensions <strong>of</strong> spores, before<br />
sunrise, but no results are available. As with so many parasitic fungi, its<br />
application would be limited by intolerance <strong>of</strong> dry conditions. Another<br />
fungus that attacks H. hampei, Paecilomyces javanicus, is Afro-<strong>Asian</strong> in<br />
distribution and wide spectrum in its host range (Samson 1974), attacking<br />
also Lepidoptera. Its use against H. hampei appears not to have been<br />
attempted.<br />
There appears to be only one record <strong>of</strong> nematodes attacking H. hampei in<br />
the field (Varaprasad et al. 1994), but in addition, Allard and Moore (1989)<br />
showed that a Heterorhabditis sp. could cause high mortality <strong>of</strong> both adult<br />
and larval H. hampei under laboratory conditions and that infective<br />
juveniles were produced from adults and larger larvae. Spraying <strong>of</strong><br />
nematodes on fallen berries might remove the need to collect them (which<br />
involves much labor), leaving them to provide mulch. Dispersal <strong>of</strong> infected<br />
adults may also spread the nematodes into the pest population. Further work<br />
with nematodes is clearly desirable.
Table 4.10.1 Natural enemies <strong>of</strong> Hypothenemus hampei<br />
Species and type<br />
HEMIPTERA<br />
Country Reference Comment<br />
PYRRHOCORIDAE (Predator)<br />
Dindymus rubiginosus<br />
HYMENOPTERA<br />
Java Wurth 1922 Not specific<br />
BETHYLIDAE (ectoparasites <strong>of</strong> immature stages)<br />
Cephalonomia stephanoderis Ivory Coast Betrem 1961; Ticheler 1961;<br />
Cochereau & Potiaroa 1994;<br />
A promising parasite<br />
Togo<br />
Klein Koch et al. 1988<br />
Goniozus sp. Ivory Coast Cochereau & Potiaroa 1994<br />
Prorops nasuta Cameroon Klein Koch et al. 1988<br />
Congo<br />
Klein Koch et al. 1988<br />
Ivory Coast Klein Koch et al. 1988<br />
Kenya<br />
Klein Koch et al. 1988<br />
Tanzania Rangi et al. 1988<br />
Togo<br />
Klein Koch et al. 1988<br />
Uganda<br />
Klein Koch et al. 1988;<br />
Klein Koch et al. 1988;<br />
Waterston 1923<br />
Scleroderma cadaverica<br />
CERAPHRONIDAE<br />
Uganda Benoit 1957 Causes severe dermatitis in man<br />
Aphanogmus (= Calliceras) dictynna<br />
EULOPHIDAE<br />
Uganda Waterston 1923 Possibly hyperparasitic<br />
Phymastichus c<strong>of</strong>fea (attacks<br />
Ivory Coast Cochereau & Potiaroa 1994<br />
adult beetles)<br />
Kenya<br />
La Salle 1990<br />
Togo<br />
Klein Koch et al. 1988<br />
168 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong>
Table 4.10.1 (contÕd) Natural enemies <strong>of</strong> Hypothenemus hampei<br />
Species and type<br />
HYMENOPTERA<br />
Country Reference Comment<br />
BRACONIDAE<br />
(ectoparasitoid and predator)<br />
Heterospilus c<strong>of</strong>feicola<br />
Uganda<br />
Tanzania<br />
Cameroon<br />
Congo<br />
Schmiedeknecht 1924<br />
CIBC 1988b<br />
Klein Koch et al. 1988<br />
Klein Koch et al. 1988<br />
Kills larvae with sting<br />
Attacks larvae <strong>of</strong> other parasites <strong>of</strong><br />
H. hampei Ñ also may be<br />
cannibalistic<br />
FORMICIDAE<br />
(predator)<br />
Crematogaster curvispinosa<br />
ACARI<br />
Brazil Pinto da Fonseca & Araujo 1939 Can cause high mortality <strong>of</strong> immature<br />
stages in c<strong>of</strong>fee berries<br />
Pyemotid mite<br />
NEMATODA<br />
New Caledonia P. Cochereau pers. comm.<br />
Heterorhabditis sp. Moore & Prior 1988<br />
Panagrolaimus sp.<br />
FUNGI<br />
India Varaprasad et al. 1994<br />
HYPHOMYCETES<br />
Beauveria bassiana<br />
(= Botrytis stephanoderis)<br />
Java<br />
Cameroon<br />
Friederichs & Bally 1922<br />
Pascalet 1939<br />
Cosmopolitan, in a variety <strong>of</strong> strains<br />
Metarhizium anisopliae Moore & Prior 1988<br />
Nomuraea rileyi Moore & Prior 1988 Usually recorded from Lepidoptera<br />
Paecilomyces (= Spicaria) javanicus Java Friederichs and Bally 1922;<br />
Samson 1974<br />
Indonesia, Asia, Africa<br />
P. tenuipes Moore & Prior 1988<br />
4.10 Hypothenemus hampei 169
170 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Attempts at biological control<br />
Africa<br />
Published information is summarised in Table 4.10.2, but there were<br />
probably a number <strong>of</strong> transfers <strong>of</strong> parasites within Africa and perhaps South<br />
America that have gone unrecorded. In the past decade the International<br />
Institute for <strong>Biological</strong> <strong>Control</strong> had adopted the policy <strong>of</strong> breeding African<br />
parasitoids in England on H. hampei in c<strong>of</strong>fee beans from the country <strong>of</strong><br />
destination. This is because <strong>of</strong> the possibility that the wasps might carry<br />
spores <strong>of</strong> fungal diseases <strong>of</strong> c<strong>of</strong>fee, especially new strains <strong>of</strong> c<strong>of</strong>fee leaf rust<br />
(Hemileia vastatrix) and c<strong>of</strong>fee berry disease (Colletotrichium c<strong>of</strong>feanum)<br />
(Moore and Prior 1988; Rangi et al. 1988; Nemeye et al. 1990; Murphy and<br />
Rangi 1991). The danger <strong>of</strong> fungal transmission could also be reduced by<br />
breeding H. hampei on an artificial diet (Brun et al. 1993; Perez et al. 1995;<br />
Villacorta 1985).<br />
CAMEROON<br />
Pascalet (1939) recommended the introduction <strong>of</strong> Heterospilus c<strong>of</strong>feicola,<br />
Prorops nasuta and Beauveria bassiana to any plantations lacking them.<br />
There is no record that this was implemented anywhere, nor whether any or<br />
all <strong>of</strong> the organisms were not already generally present.<br />
CONGO<br />
Sladden (1934) and Leroy (1936) suggested that, by breeding and liberating<br />
them, it would be possible to increase the efficiency <strong>of</strong> P. nasuta and<br />
H. c<strong>of</strong>feicola, which he knew to be already present in the Congo and he made<br />
a similar suggestion for fungus diseases. However, there is no indication <strong>of</strong><br />
the extent to which this was done.<br />
KENYA<br />
Prorops nasuta was sent from Uganda to Kenya in 1930 (Greathead 1971),<br />
but according to Evans (1965) that wasp and H. c<strong>of</strong>feicola were probably<br />
native there. Abasa (1975) considered that parasites were <strong>of</strong> doubtful value<br />
in controlling H. hampei in Kenya.<br />
UGANDA<br />
Prorops nasuta and Heterospilus c<strong>of</strong>feicola are both native to Uganda.<br />
Hargreaves (1935) considered that some areas lacked these parasites, and so<br />
he introduced cultures from Kampala County, north <strong>of</strong> Lake Victoria, to<br />
Bwamba County on the western border. He stated that this introduction<br />
resulted in a great reduction in the previously intense infestation <strong>of</strong> c<strong>of</strong>fee<br />
berry borer but, in view <strong>of</strong> the natural occurrence <strong>of</strong> P. nasuta over a wide<br />
area to the west <strong>of</strong> the Ugandan border (Le Pelley 1968), it seems unlikely<br />
that the distribution was discontinuous and that it was lacking in Bwamba
Asia<br />
4.10 Hypothenemus hampei 171<br />
County. HargreavesÕ claims that the introduction brought about a great<br />
reduction in the impact <strong>of</strong> the c<strong>of</strong>fee berry borer in Bwamba County must be<br />
treated with reserve, the more so since De Toledo Piza and Pinto da Fonseca<br />
(1935) state that neither P. nasuta nor H. c<strong>of</strong>feicola appeared to control the<br />
borer in nearby Kampala. More recently, P. nasuta was reported to achieve<br />
20% parasitisation in western Kenya in the dry season (Barrera et al. 1990b).<br />
SRI LANKA<br />
Stock <strong>of</strong> P. nasuta and H. c<strong>of</strong>feicola from Uganda were liberated in Sri<br />
Lanka in 1938, but neither species became established (Hutson 1939).<br />
INDONESIA<br />
The introduction <strong>of</strong> Prorops nasuta to Java from Uganda in 1923 was the<br />
earliest attempt to bring about the biological control <strong>of</strong> H. hampei which had<br />
first been reported in Java in 1909 (Kalshoven 1981). P. nasuta was found to<br />
be easily propagated (Leefmans 1924a), was distributed widely in<br />
considerable numbers (Begemann 1926) and became established (Le Pelley<br />
1968). However, it apparently could not maintain itself and was still being<br />
bred for distribution in 1928 (UltŽe 1928) and in 1932 (Betrem 1932;<br />
Schweizer 1932; UltŽe 1932).<br />
Leefmans (1924a) drew attention to the fact that the P. nasuta did not<br />
thrive in shade, and that it flourished best in black berries which tend to be<br />
most abundant after harvest, when the parasite is least needed. The former<br />
problem was solved by appropriate pruning but, despite improvements in<br />
management to favour it and the long period spent in breeding and<br />
disseminating it, the parasite seems not to have become established in Java<br />
(Clausen 1978; Kalshoven 1981).<br />
Cultures <strong>of</strong> Heterospilus c<strong>of</strong>feicola were taken to Java from Uganda<br />
along with those <strong>of</strong> P. nasuta in 1923, but the wasp appears not to have been<br />
released. Leefmans (1924a) seems to have concluded that it was likely to be<br />
incompatible with P. nasuta.
Table 4.10.2 Introductions for the biological control <strong>of</strong> Hypothenemus hampei<br />
Country and species liberated<br />
BRAZIL<br />
Year From Result Reference<br />
Cephalonomia stephanoderis ? ? + Benassi & Berti-Filho 1989<br />
Prorops nasuta<br />
COLOMBIA<br />
1929 Uganda + Hempel 1933; Yokoyama et al. 1978<br />
Cephalonomia stephanoderis 1988 Kenya via U.K. + C. Klein Koch pers. comm. 1995;<br />
Sponagel 1993<br />
Prorops nasuta<br />
ECUADOR<br />
1995 + Bustillo et al. 1995;<br />
Portilla & Bustillo 1995<br />
Cephalonomia stephanoderis 1988 Togo via U.K. + CIBC 1988b; Klein Koch 1989a,b,c;<br />
Klein Koch et al. 1988; Delgado et al.<br />
1990; Sponagel 1993<br />
Prorops nasuta 1987Ð1990<br />
1988<br />
Kenya via U.K.<br />
West Africa<br />
Ð<br />
Ð<br />
CIBC 1988a; Klein Koch et al. 1988;<br />
Rangi et al. 1988; Murphy & Rangi<br />
1991; Sponagel 1993<br />
EL SALVADOR<br />
Cephalonomia stephanoderis<br />
GUATEMALA<br />
1988 Kenya via U.K. + Sponagel 1993; C. Klein Koch pers.<br />
comm. 1995<br />
Cephalonomia stephanoderis 1988<br />
1993-1995<br />
HONDURAS<br />
Kenya via U.K. ? Sponagel 1993<br />
Garcia & Barrios 1996<br />
Cephalonomia stephanoderis 1988 Kenya via U.K. ? Sponagel 1993<br />
172 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong>
Table 4.10.2 (contÕd) Introductions for the biological control <strong>of</strong> Hypothenemus hampei<br />
Country and species liberated<br />
INDONESIA<br />
Year From Result Reference<br />
Cephalonomia stephanoderis 1988 ? Sponagel 1993<br />
Heterospilus c<strong>of</strong>feicola 1923<br />
1931<br />
Prorops nasuta 1923Ð<br />
1925<br />
KENYA<br />
Uganda<br />
Uganda<br />
not liberated<br />
?<br />
Kalshoven 1981<br />
Le Pelley 1968<br />
Uganda Ð Begemann 1926<br />
Kalshoven 1981<br />
Prorops nasuta 1930 Uganda already present Evans 1965<br />
MEXICO<br />
Cephalonomia stephanoderis 1988Ð1989 Togo via U.K. + Barrera et al. 1990 a, b;<br />
CIBC 1988b<br />
Prorops nasuta<br />
NEW CALEDONIA<br />
1988Ð1989 Kenya and Togo via U.K. ? Barrera et al. 1990a, b;<br />
Murphy & Rangi 1991<br />
Cephalonomia stephanoderis<br />
UGANDA (BWAMBA COUNTY)<br />
1993 Ivory Coast + Cochereau & Potiaroa 1994<br />
Prorops nasuta<br />
PERU<br />
1932 Uganda (Kampala county) +<br />
?already present<br />
Prorops nasuta 1962<br />
1964? Brazil Ð<br />
SRI LANKA<br />
Hargreaves 1935<br />
Clausen 1978<br />
De Ingunza 1964<br />
Heterospilus c<strong>of</strong>feicola 1938 Uganda Ð Hutson 1939<br />
Prorops nasuta 1938 Uganda Ð Hutson 1939<br />
4.10 Hypothenemus hampei 173
174 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Pacific<br />
NEW CALEDONIA<br />
Infestation <strong>of</strong> c<strong>of</strong>fee berries with H. hampei ranges from 0% to 100%, with<br />
an overall average <strong>of</strong> 33%. C. stephanoderis from West Africa was released<br />
in 1993 and recovered almost a year later. However, wherever the<br />
aggressive, little red fire ant (Wasmannia auropunctata, introduced around<br />
1970) is present this parasitoid is unable to survive. When the fire ant is<br />
eliminated from a plantation by banding the trees with insecticide the wasp is<br />
established. Ant control is, thus, a prerequisite for biological control<br />
(Cochereau and Potiara 1994; P. Cochereau pers. comm. 1995).<br />
Phymastichus c<strong>of</strong>fea is currently under consideration for liberation<br />
(P. Cochereau pers. comm.). Cochereau et al. (1994) have examined in some<br />
detail the effectiveness against H. hampei <strong>of</strong> a virulent New Caledonian<br />
strain <strong>of</strong> Beauveria bassiana, which shows considerable promise in the field.<br />
Central America<br />
GUATEMALA<br />
Cephalonomia stephanoderis was mass produced and released during 1993<br />
to 1995. Infestation by H. hampei was reduced 75%, to 2.7 to 5.3% in 1993<br />
and to 0.4 to 0.9% in 1994, but an increase was observed in 1995 (1.6 to<br />
2.4%, compared with the control <strong>of</strong> 3.8% infestation), resulting in 48%<br />
control (Garcia and Barrios 1996). The cost <strong>of</strong> mass liberations <strong>of</strong><br />
C. stephanoderis was comparable with that <strong>of</strong> chemical control (Decazy et<br />
al. 1995).<br />
MEXICO<br />
Cephalonomia stephanoderis from Togo and Prorops nasuta from both<br />
Togo and Kenya were raised in U.K. in c<strong>of</strong>fee beans from Mexico before<br />
being sent there for mass production and liberation during 1988 and 1989.<br />
C. stephanoderis has become established, but the situation with P. nasuta is<br />
unclear (Barrera et al. 1990a, b). The impact <strong>of</strong> these parasitoids remains to<br />
be reported.<br />
South America<br />
BRAZIL<br />
Prorops nasuta was imported into Brazil from Uganda in 1929, and by 1933<br />
it was established in several c<strong>of</strong>fee plantations (Hempel 1933, 1934). As in<br />
Java, breeding and distribution continued and in 1937 (Anon. 1937) it was<br />
stated to be <strong>of</strong> considerable value in controlling the c<strong>of</strong>fee berry borer in S‹o<br />
Paulo, but only if its numbers were boosted by rearing between c<strong>of</strong>fee<br />
production seasons. Puzzi (1939) studied the reproduction <strong>of</strong> the parasite in
4.10 Hypothenemus hampei 175<br />
relation to that <strong>of</strong> its host in Brazil and concluded that, in theory, it was more<br />
prolific, but that the efficiency <strong>of</strong> the parasite was limited by the tendency <strong>of</strong><br />
the female to remain in one berry. De Toledo (1942) examined rates <strong>of</strong><br />
parasitisation, but his figures do not suggest that the wasp could have been<br />
having any significant impact. De Toledo et al. (1947) were only mildly<br />
enthusiastic about the value <strong>of</strong> the wasp, mentioning a continuing need for<br />
repeated liberations and the requirement for boosting the effect <strong>of</strong> the<br />
parasite by cultural practices. Le Pelley (1968) stated that, at that time,<br />
Brazilian entomologists appeared satisfied that P. nasuta was <strong>of</strong> value in<br />
their country, but he could find no conclusive evidence that the amount <strong>of</strong><br />
routine work required for the control <strong>of</strong> H. hampei had decreased.<br />
Yokayama et al. (1978), considered that the climate <strong>of</strong> the S‹o Paulo district<br />
in Brazil was unfavourable for this wasp, in which the growers lost interest<br />
when BHC was found to give satisfactory control. Nevertheless, they<br />
reported that it had recently been recovered in c<strong>of</strong>fee plantations in S‹o<br />
Paulo, having survived pesticide usage, severe droughts and winter frosts.<br />
The fact that P. nasuta has been transferred more recently to areas in Brazil<br />
where it is not established indicates that the wasp is considered to be <strong>of</strong> some<br />
value (Ferreira and Batistela Sobrinho 1987).<br />
Although there is no record <strong>of</strong> its release, a species <strong>of</strong> Cephalonomia,<br />
presumably C. stephanoderis was recovered in the field in Brazil between<br />
1986 and 1988 (Benassi and Berti-Filho 1989).<br />
A survey <strong>of</strong> natural enemies <strong>of</strong> H. hampei in northern Espirito Santo<br />
from 1986 to 1994 revealed 3 parasitoids (Prorops nasuta, Cephalonomia<br />
sp., and a species <strong>of</strong> Proctotrupoidea), an ant predator (Crematogaster<br />
curvispinosa) and a fungus (Beauveria bassiana) (Benassi 1995).<br />
COLOMBIA<br />
The rate <strong>of</strong> population increase <strong>of</strong> H. hampei in the field has been studied by<br />
Gaviria et al. (1995). A major program <strong>of</strong> integrated management<br />
commenced in 1992, involving local strains <strong>of</strong> the fungi Beauveria bassiana<br />
and Metarhizium anisopliae and parasitoids, in particular Cephalonomia<br />
stephanoderis and Prorops nasuta (CABI: IIBC 1993; C. Klein Koch pers.<br />
comm. 1994; Bustillo et al. 1995). Methods for mass production <strong>of</strong><br />
C. stephanoderis and Prorops nasuta are provided by Portilla and Bustillo<br />
(1995). Sixty million parasitoids were released and 100 tons <strong>of</strong> B. bassiana<br />
and M. anisopliae were applied (Bustillo et al. 1995).<br />
ECUADOR<br />
Klein Koch (1986) proposed the introduction <strong>of</strong> 3 parasitoids from Africa,<br />
Prorops nasuta, Heterospilus c<strong>of</strong>feicola and Cephalonomia stephanoderis.<br />
C. stephanoderis was first introduced from Togo in 1988 and P. nasuta from<br />
Tanzania and Kenya in 1987Ð88 and Togo in 1988 (Klein Koch 1989c).
176 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
PERU<br />
Liberations continued in subsequent years with some 920000 <strong>of</strong> the former<br />
species and 30,000 <strong>of</strong> the latter being released in 1992. Both species are now<br />
well established and having a significant effect. In field experiments with<br />
caged c<strong>of</strong>fee trees 86% <strong>of</strong> berries on the bush and 87% on the ground<br />
contained C. stephanoderis and up to 52% and 31% parasitisation<br />
respectively was recorded from tree and ground berries in the open (Klein<br />
Koch 1989b,c, 1990). However the results are poor in the Amazon region <strong>of</strong><br />
the country where the rainfall is very high. Elsewhere, as part <strong>of</strong> an<br />
integrated approach, infestation levels on C<strong>of</strong>fea arabica are as low as 0.4 to<br />
1.4%. Recently introduced catimor varieties are resistant to c<strong>of</strong>fee rust, so<br />
sprays are no longer required (Klein Koch pers. comm. 1994). In addition,<br />
the fungus Beauveria bassiana and the ant Azteca sp. cause considerable<br />
mortality <strong>of</strong> H. hampei when humid weather conditions prevail (Klein Koch<br />
et al. 1987).<br />
According to De Ingunza (1964) Prorops nasuta was introduced from Brazil<br />
to Peru in 1962, but failed to become established.<br />
Major parasite species<br />
Cephalonomia stephanoderis Hym.: Bethylidae<br />
Cephalonomia stephanoderis is a small black bethylid wasp which is native<br />
to West Africa (Ivory Coast, Togo). The females, which are 1.6 to 2.0 mm in<br />
length, enter bored c<strong>of</strong>fee berries and deposit eggs on the ventral surface <strong>of</strong><br />
final stage larvae and prepupae <strong>of</strong> H. hampei. Its larvae feed as ectoparasites,<br />
exhausting the tissues <strong>of</strong> the host in 4 to 6 days, then spinning a silken<br />
cocoon in which to pupate. The pupal stage lasts about 15 days. Fertilisation<br />
takes place in the berry where the wasps emerge, and seemingly, the males,<br />
although fully winged, remain there after the females have left. Females<br />
must feed for two days at 27¡C or 6 to 11 days at 24¡C before they can<br />
mature eggs. Adult females feed by preference on H. hampei eggs and<br />
young larvae but also on prepupae and they chew holes in the intersegmental<br />
membrane <strong>of</strong> adult beetles, between the prothoracic and mesothoracic<br />
tergites, and feed on the haemolymph. Females cannot produce eggs on a<br />
diet <strong>of</strong> borer eggs or adults alone, but need to feed first on the larvae and/or<br />
prepupae <strong>of</strong> the borer. They can lay up to 70 eggs (Barrera et al. 1989, 1993;<br />
Abraham et al. 1990; Wegbe 1990; Infante et al. 1994a ) and can distinguish<br />
between parasitised and unparasitised hosts (Barrera et al. 1994). In the<br />
Ivory Coast, Koch (1973) found that adult C. stephanoderis each required<br />
two eggs, two larvae or two adults per day for survival. At the end <strong>of</strong> the<br />
c<strong>of</strong>fee season H. hampei populations were reduced by parasitisation by 20%
4.10 Hypothenemus hampei 177<br />
to 30%, but by not more than 5% between seasons. Ticheler (1961) recorded<br />
up to 50% parasitisation by this, the most important parasitoid in the Ivory<br />
Coast. In West Africa C. stephanoderis is commoner than P. nasuta<br />
(Abraham et al. 1990). A major mass production and release program<br />
commenced in Colombia in 1993, with a production <strong>of</strong> 10 million wasps per<br />
month. When 400 000 wasps were released in a 2 million ha area <strong>of</strong> c<strong>of</strong>fee<br />
trees 85% parasitisation was attained when there was 80% infestation <strong>of</strong><br />
c<strong>of</strong>fee berries and 20% parasitisation when there was 5% infestation. The<br />
target for the releases was 12.5 wasps per berry and 300 berries containing<br />
wasps for each 15 trees, giving 20 000 to 30 000 wasps per ha (P. Cochereau<br />
pers. comm. 1995). Life tables were developed for C. stephanoderis in<br />
Mexico (Infante and Luis 1993; Infante et al. 1994b).<br />
Studies <strong>of</strong> mass releases <strong>of</strong> C. stephanoderis suggest that they can<br />
control low density populations <strong>of</strong> H. hampei in commercial c<strong>of</strong>fee<br />
plantations, adult predation by the wasp probably being the most important<br />
mortality factor. However, mass production costs are too high for releases to<br />
be economically viable and cheap artificial diets for mass rearing are being<br />
investigated, with successful rearing already achieved for four generations<br />
(CABI:IIBC 1996).<br />
Heterospilus c<strong>of</strong>feicola Hym.: Braconidae<br />
Heterospilus c<strong>of</strong>feicola is a braconid wasp about 2.5 mm long. It does not<br />
enter the borehole <strong>of</strong> the beetle, but travels from berry to berry inserting its<br />
ovipositor into the boreholes in the course <strong>of</strong> seeking Hypothenemus larvae.<br />
Only one small egg is laid in each berry, and the larva that emerges after<br />
about six days feeds on beetle eggs and larvae over a period <strong>of</strong> 18 to 20 days,<br />
consuming 10 to 15 eggs and larvae per day. In this regard it is more <strong>of</strong> a<br />
predator than a parasite. According to De Toledo Piza and Pinto da Fonseca<br />
(1935) the larva kills the adult H. hampei before pupating inside a white<br />
silken cocoon. The wasp emerges after a comparatively brief pupal period<br />
(Hargreaves 1926; De Toledo Piza and Pinto da Fonseca 1935, Le Pelley<br />
1968). In Uganda it is attacked by a chalcidid <strong>of</strong> the genus Closterocerus<br />
(Schmiedeknecht 1924).<br />
Hargreaves (1926, 1935) stated that Heterospilus c<strong>of</strong>feicola contributed<br />
substantially to the control <strong>of</strong> H. hampei in Uganda. The Brazilian<br />
entomologists De Toledo Piza and Pinto da Fonseca (1935) studied the wasp<br />
in Uganda with a view to assessing its potential value as a biological control<br />
agent in Brazil. They concluded that H. c<strong>of</strong>feicola can thrive only in areas<br />
with a continuous production <strong>of</strong> c<strong>of</strong>fee berries throughout the year, and as<br />
such conditions prevail nowhere in Brazil they recommended against its<br />
importation. One possible disadvantage <strong>of</strong> this wasp is that its larvae feed on
178 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
the larvae <strong>of</strong> other wasps as well as those <strong>of</strong> H. hampei, and it may even be<br />
cannibalistic (Hargreaves 1924). If these statements are verified then there<br />
may be reservations about the employment <strong>of</strong> H. c<strong>of</strong>feicola in biological<br />
control. A further difficulty associated with this species as a biological<br />
control agent is that a number <strong>of</strong> workers have been unable to breed it in the<br />
laboratory, a problem also experienced by CIBC (1987) during its current<br />
program, although Rangi et al. (1988) have reported limited success. The<br />
free-living existence <strong>of</strong> the adults may involve special nutritional or mating<br />
requirements that have not yet been met experimentally.<br />
Phymastichus c<strong>of</strong>fea Hym.: Eulophidae<br />
This parasitoid was first recorded in Togo as recently as 1989 (Borbon-<br />
Martinez 1989), causing up to 30% parasitisation, but is now known also<br />
from Ivory Coast and Kenya. It is an endoparasite <strong>of</strong> the adult female<br />
H. hampei, which is usually attacked as she is commencing to tunnel in to a<br />
c<strong>of</strong>fee berry. P. c<strong>of</strong>fea also enters the berry to parasitise male H. hampei.<br />
Oviposition occurs in both the thorax and the abdomen <strong>of</strong> the host, the<br />
former producing a male and the latter a female. Although several eggs may<br />
be laid in each host, only two wasps are produced. Males range in length<br />
from 0.45 to 0.55 mm and females from 0.8 to 1.0 mm. Females do not<br />
require to be fertilised before they commence oviposition shortly after<br />
emergence. At 27¡C, larval development takes about 21 days, the pupal<br />
stage about 8 days and adult longevity appears to be a few days only. In the<br />
field 20 females were produced for each male and it was estimated that<br />
between 4 and 7 hosts could be parasitised in a 4-hour period.<br />
P. c<strong>of</strong>fea was the most important parasitoid <strong>of</strong> H. hampei in the majority<br />
<strong>of</strong> c<strong>of</strong>fee holdings on the Togolese Plateau at about 800m above sea level<br />
and is fairly common around Man (West Ivory Coast) near the Liberian<br />
border (La Salle 1990; Feldhege 1992; P. Cochereau pers. comm. 1994;<br />
Infante et al. 1994a, ).<br />
Mass production methods have been developed in Colombia for<br />
P. c<strong>of</strong>fea and a decision to release is expected shortly (CABI:IIBC 1996).<br />
Prorops nasuta Hym.: Bethylidae<br />
Prorops nasuta is native to Uganda, Kenya, Tanzania and Cameroon, Ivory<br />
Coast and Togo. It is a dark brown bethylid wasp about 2.3mm in length, the<br />
name nasuta referring to the characteristic bilobed ÔnoseÕ protruding<br />
forwards above the antennal bases. This wasp parasitises and preys upon<br />
several species <strong>of</strong> Hypothenemus (Clausen 1978). Males that emerge first<br />
from pupae always stay on the remaining cocoons within the c<strong>of</strong>fee berry<br />
and mate with the females as they emerge. Because P. nasuta populations<br />
are, thus, highly inbred it is probable that there are different strains <strong>of</strong>
4.10 Hypothenemus hampei 179<br />
P. nasuta in West Africa and East Africa, (Abraham et al. 1990; Griffiths<br />
and Godfray 1988; Murphy and Rangi 1991) and testing these may be highly<br />
relevant to biological control. On c<strong>of</strong>fee the fertilised female enters an<br />
infested berry via the borehole <strong>of</strong> the adult H. hampei, choosing berries on<br />
the trees rather than those on the ground. If the parent borer beetle is still<br />
present she may kill it and use the cadaver to plug the entrance hole, over<br />
which she stands guard. According to CIBC (1987) the female wasp does not<br />
feed upon borer beetles she may kill, but other authors state that she will do<br />
so if no other life history stages are available, but that she cannot mature eggs<br />
on a diet <strong>of</strong> adults alone. Several larvae and pupae may be injured with the<br />
ovipositor before any oviposition occurs, and these victims succumb in a<br />
few days. P. nasuta feeds by preference on the eggs and young larvae <strong>of</strong><br />
H. hampei and oviposit on the late third stage larvae and pupae. The hosts<br />
chosen for oviposition are stung, sometimes several times, and thus<br />
paralysed before one, or sometimes two, eggs are laid upon them. Eggs are<br />
placed ventrally on larvae and on the abdominal dorsum <strong>of</strong> pupae. The eggs<br />
(0.55 ´ 0.18mm) are large for a wasp <strong>of</strong> this size. They hatch in an average <strong>of</strong><br />
about three days and the larval stages last three to eight days. The<br />
ectoparasitic larva may consume more than one host. There is a prepupal<br />
(non-feeding) period <strong>of</strong> about three days, passed inside a silken cocoon spun<br />
by the fully fed larva. It is common to find 20 cocoons in a c<strong>of</strong>fee bean, and<br />
up to 62 have been recorded. The pupal stage lasts on an average about 21<br />
days, varying from 9 to 27 days according to temperature.<br />
The life cycle from egg to adult lasts 17 to 33 days (average 29) at 25¡C<br />
and may be as long as 66 days at 18¡C. There are considerable discrepancies<br />
between figures given by various authors for the duration <strong>of</strong> the life history<br />
stages, but there is general agreement that the female is quite long-lived Ñ<br />
up to 135 days being cited in Brazil, given an abundant supply <strong>of</strong> larvae and<br />
prepupae as food. By contrast it appears that the males do not feed and they<br />
do not survive longer than 13 days. Females outnumber the males, a figure <strong>of</strong><br />
three to one being recorded. Statements as to duration <strong>of</strong> the preoviposition<br />
period give rather diverse figures. Usually a few days are indicated but one<br />
record is <strong>of</strong> 17 days. Parthenogenesis may occur, when only male progeny<br />
are produced. Females may lay up to 66 eggs at a rate <strong>of</strong> one or two a day,<br />
utilising several berries in the process.<br />
In feeding, females consume several eggs and unparasitised larvae per<br />
day and they will also eat pupae. Normally all stages <strong>of</strong> the beetle in a berry<br />
are killed either by parasitisation, predation or merely by stabbing before the<br />
female leaves (Leefmans 1924a; Begemann 1926; Hempel 1933; De Toledo<br />
Piza and da Fonseca 1935; Hargreaves 1935; Hutson 1936; Puzzi 1939; De<br />
Toledo 1942; Le Pelley 1968; Abraham et al. 1990; ). The low abundance <strong>of</strong>
180 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
P. nasuta in Western Kenya suggests that it may not be suited to high<br />
altitudes (Murphy and Rangi 1991). In one locality in S‹o Paulo (Brazil) the<br />
percentage <strong>of</strong> infested berries that contained parasitoids rose to a maximum<br />
<strong>of</strong> 2.4 in autumn (De Toledo 1942).<br />
Scleroderma cadaverica Hym.: Bethylidae<br />
Scleroderma cadaverica is listed by Herting and Simmonds (1973) as a<br />
natural enemy <strong>of</strong> H. hampei, but only some <strong>of</strong> the specimens before Benoit<br />
(1957) when he prepared the taxonomic description had been reared from<br />
that species, others being stated to come from small beetles boring in cane<br />
furniture. The North American species <strong>of</strong> Scleroderma are stated by<br />
Krombein et al. (1979) to be parasitic on the larvae <strong>of</strong> small wood-boring<br />
beetles, the female wasps frequently stinging people inhabiting infested<br />
houses. The African specimens <strong>of</strong> S. cadaverica were submitted to European<br />
specialists for identification and description because stinging by females<br />
(which may be either winged or apterous) had caused severe dermatitis to<br />
African and European people. No responsible person would consider using<br />
this insect in biological control projects.<br />
Comments<br />
Hypothenemus hampei has established itself in most <strong>of</strong> the c<strong>of</strong>fee growing<br />
areas <strong>of</strong> the world, but there are still uninfested countries, such as Australia,<br />
Hawaii, Papua New Guinea, Vanuatu and the Solomon Islands. Quarantine<br />
is <strong>of</strong> critical importance to these countries and it is important to ensure that<br />
c<strong>of</strong>fee imported into clean areas has been completely disinfested. Thorough<br />
drying <strong>of</strong> the seed c<strong>of</strong>fee is an indispensable supplement to disinfestation<br />
techniques.<br />
Although great advances have been made in recent years in the chemical<br />
control <strong>of</strong> H. hampei, it would be a great advantage to have the support <strong>of</strong><br />
additional measures. Rhodes and Mansingh (1981) found chemical control<br />
inadequate on its own in Jamaica and advocated its integration with cultural<br />
practices and Bardner (1978) emphasised the need to harmonise cultural,<br />
biological and chemical control <strong>of</strong> c<strong>of</strong>fee pests. Hernandez Paz and Penagos<br />
Dardon (1974) found, in Guatemala, that low-volume sprays <strong>of</strong> endosulfan<br />
could completely destroy H. hampei in berries on the bushes and, according<br />
to Mansingh and Rhodes (1983), this chemical is in extensive use in Central<br />
and South America. However, a very high level <strong>of</strong> resistance to endosulfan<br />
in H. hampei is reported in New Caledonia (Brun et al. 1989, 1990), raising<br />
concern that this valuable insecticide may not remain an effective material<br />
for long.
4.10 Hypothenemus hampei 181<br />
Plantation sanitation is an old-established tradition in pest control, and<br />
the c<strong>of</strong>fee berry borer has long been attacked from this angle. The life cycle<br />
<strong>of</strong> the borer, indeed, lends itself to this approach, as it is narrowly specific to<br />
the c<strong>of</strong>fee berries. In Java, Roepke (1912) and Leefmans (1924b)<br />
recommended the total destruction <strong>of</strong> infested or susceptible berries over a<br />
period long enough to break the life cycle <strong>of</strong> the c<strong>of</strong>fee berry borer. A period<br />
<strong>of</strong> three months was aimed at, although some records <strong>of</strong> the longevity <strong>of</strong><br />
beetles exceed this. Measures taken involve the collection <strong>of</strong> all fallen<br />
berries and the picking <strong>of</strong> any that may have escaped the harvest, plus the<br />
continuous removal <strong>of</strong> all young berries on which adult female beetles might<br />
feed. Friederichs (1922) and Rutgers (1922) reported successful application<br />
<strong>of</strong> the method in Java. The latter reported that, on estates which had applied<br />
the measures, the percentage <strong>of</strong> infested berries fell from 40% to 90% to<br />
between 0.5% and 3.0%. In New Caledonia application <strong>of</strong> the method<br />
reduced infestation from 80% to 10% (Cohic 1958). In Malaysia, Corbett<br />
(1933) recommended picking at weekly intervals (or the shortest period<br />
practicable, according to size <strong>of</strong> holding) <strong>of</strong> all bored green, ripe and<br />
blackened berries on the bush and from the ground. A host-free period <strong>of</strong> six<br />
months was recommended for the eradication <strong>of</strong> the beetle from isolated<br />
plantations. In Mexico, Baker (1984) concluded that berries are not infested<br />
while lying on the ground surface. Nevertheless any infested berries allowed<br />
to lie where they fell ultimately generated large numbers <strong>of</strong> beetles.<br />
Turning to biological control, it is generally possible to gain some idea <strong>of</strong><br />
the likely effectiveness <strong>of</strong> natural enemies in a new country from their<br />
impact in their country <strong>of</strong> origin. In a new country the enemies may be more<br />
effective if freed from hyperparasites before transfer or less effective if they<br />
are poorly adapted to their new environment. Based on their evaluation <strong>of</strong><br />
the population dynamics <strong>of</strong> H. hampei and its parasitoids, Moore and Prior<br />
(1988) and Murphy and Moore (1990) were optimistic about the value <strong>of</strong><br />
biological control as a key component <strong>of</strong> successful integrated management<br />
<strong>of</strong> H. hampei. Although this indeed seems probable, the reports from Africa<br />
are far from uniform and the c<strong>of</strong>fee berry borer is a problem in some areas.<br />
This may be due to whether arabica or robusta c<strong>of</strong>fee is involved and to the<br />
widely different conditions under which c<strong>of</strong>fee is grown and harvested<br />
since, for example, parasitoids may be less effective when c<strong>of</strong>fee is shaded<br />
(Hargreaves 1935).<br />
Hargreaves (1926, 1935) considered that both Prorops nasuta and<br />
Heterospilus c<strong>of</strong>feicola were important in regulating populations <strong>of</strong><br />
H. hampei in commercial c<strong>of</strong>fee at about 1200m in Uganda and Pascalet<br />
(1939) arrived at a similar view concerning these parasitoids in Cameroon.<br />
On the other hand, Abasa (1975) in Kenya, De Toledo Piza and Fonseca
182 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
(1935) and Ingram (quoted by Le Pelley 1968, p 125) in Uganda and Sladden<br />
(1934) and Schmitz and Crisinel (1957) in Zaire concluded that parasitoids<br />
had little influence on the number <strong>of</strong> bored berries. Murphy and Moore<br />
(1990) considered that P. nasuta did not have a major impact in Western<br />
Kenya where H. c<strong>of</strong>feicola was not encountered. There, c<strong>of</strong>fee berries<br />
infested with H. hampei, ranged in being attacked by P. nasuta from 0% in<br />
the wet season to 25% in the dry season and P. nasuta populations did not<br />
build up until after the annual c<strong>of</strong>fee harvest when populations <strong>of</strong> H. hampei<br />
had already crashed from their annual peak.<br />
Although P. nasuta is present in Ivory Coast (Diro, Man) it is usually<br />
rare and H. c<strong>of</strong>feicola apparently absent. However, Cephalonomia<br />
stephanoderis was common and up to 50% <strong>of</strong> colonies <strong>of</strong> H. hampei in black<br />
berries were parasitised, resulting in important population reduction<br />
(Ticheler 1961), a conclusion contested by Koch (1973) on unsubstantiated<br />
grounds. It is relevant that, in the presence <strong>of</strong> C. stephanoderis, but without<br />
any chemical treatment, Cochereau and Potiaroa (1994) reported that only<br />
2% <strong>of</strong> the c<strong>of</strong>fee beans (4% <strong>of</strong> the berries) were attacked in Ivory Coast by<br />
H. hampei. They contrast this with the situation in New Caledonia where it<br />
was common to find 50% <strong>of</strong> the berries bored, even in plantations treated<br />
with endosulfan.<br />
Experience in biological control suggests that the negative assessments<br />
<strong>of</strong> the value <strong>of</strong> parasites in Africa may reflect the restraining influence <strong>of</strong><br />
hyperparasites (such as Aphanogmus (= Ceraphron) dictynna), predators,<br />
competitors and diseases, but it seems that no such impediments were taken<br />
to Brazil with the original stocks <strong>of</strong> Prorops nasuta. Nevertheless, Le Pelley<br />
(1968) stated that there was no evidence that, 35 years after the introduction,<br />
Brazilian c<strong>of</strong>fee growers had to invest less effort in other control measures<br />
and Yokayama et al. (1978), 45 years after its introduction, gave a<br />
depressing picture <strong>of</strong> its impact. Greater success may attend the more recent<br />
employment <strong>of</strong> Cephalonomia stephanoderis.<br />
In spite <strong>of</strong> the conflicting reports, there are, on balance, good reasons for<br />
maintaining optimism that natural enemies can play an important role in<br />
reducing the losses caused by H. hampei. There are the still to be evaluated<br />
prospects that C. stephanoderis will prove useful and there are various<br />
strains (and intraspecific crosses) <strong>of</strong> P. nasuta, in addition to Phymasticus<br />
c<strong>of</strong>fea, Heterospilus c<strong>of</strong>feicola and Goniozus to consider. It is quite possible<br />
also that further work, especially in countries such as Ethiopia, which have<br />
not been studied, will reveal additional parasitoid species. It is possible that<br />
nematodes may prove effective in controlling infestations in fallen berries<br />
and likely that applications <strong>of</strong> a virulent strain <strong>of</strong> Beauveria bassiana may<br />
prove to be a valuable alternative to pesticides. It is fortunate for <strong>Southeast</strong>
4.10 Hypothenemus hampei 183<br />
<strong>Asian</strong> countries that active work on both parasitoids and B. bassiana is in<br />
progress in a number <strong>of</strong> South American countries and in New Caledonia,<br />
from which valuable new information will emerge. However, this is not a<br />
justification to delay action if H. hampei is high enough on the priority list,<br />
since it is likely that parasitoid cultures and relevant expertise will be more<br />
readily and economically available now than they will be some years hence.
4.11 Leucinodes orbonalis<br />
India<br />
Myanmar<br />
++<br />
20°<br />
Laos<br />
+<br />
0°<br />
20°<br />
China<br />
++<br />
Thailand<br />
+<br />
Cambodia<br />
++<br />
Vietnam<br />
+++<br />
+++<br />
++ Brunei<br />
Malaysia<br />
+<br />
Singapore<br />
P<br />
Indonesia<br />
Taiwan<br />
++<br />
P<br />
Philippines<br />
Australia<br />
Papua<br />
New Guinea<br />
185<br />
It appears that Leucinodes orbonalis has been introduced from India to <strong>Southeast</strong> Asia.<br />
In India and most other countries, egg plant (brinjal) is protected against insect pests with<br />
heavy applications <strong>of</strong> broad spectrum pesticides, which must also suppress parasitoids<br />
and predators. Several parasitoids, and especially the ichneumonid Trathala flavoorbitalis,<br />
are capable <strong>of</strong> producing in excess <strong>of</strong> 50% combined mortality. If levels <strong>of</strong> this magnitude<br />
are combined with the widespread adoption <strong>of</strong> partially resistant egg plant cultivars, there<br />
are good reasons for believing that substantial pest control would be achieved.<br />
20°<br />
0°<br />
20°
186 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Leucinodes orbonalis GuenŽe<br />
Rating<br />
Origin<br />
Distribution<br />
Lepidoptera: Pyralidae<br />
brinjal fruit borer, eggplant fruit and shoot borer<br />
<strong>Southeast</strong> Asia China Southern and Western Pacific<br />
+++ Viet, Brun, Phil<br />
18 ++ Myan, Camb, Msia ++ absent<br />
+ Thai, Laos, Sing<br />
P Indo<br />
According to Purseglove (1968), eggplant ( Solanum melongena)<br />
is native to<br />
India and there are certainly many varieties in cultivation there. It is<br />
probably safe to conclude that L. orbonalis is also native to India, although it<br />
is not confined to eggplant and also occurs widely in Africa (see below).<br />
Leucinodes orbonalis is closely related to Central American Neoleucinodes<br />
species such as N. elegantalis,<br />
which is a widespread pest there <strong>of</strong> eggplant,<br />
and other Solanaceae. This suggests that parasitoids <strong>of</strong> Neoleucinodes may<br />
be <strong>of</strong> interest for Leucinodes and vice versa.<br />
L. orbonalis occurs in the Indian subcontinent (Andaman Is, India,<br />
Pakistan, Nepal, Bangladesh, Sri Lanka), Southern Asia all 10 <strong>Southeast</strong><br />
<strong>Asian</strong> nations, also Hong Kong, China, Taiwan, Japan, Africa (Burundi,<br />
Cameroon, Congo, Ethiopia, Ghana, Kenya, Lesotho, Malawi,<br />
Mozambique, Nigeria, Rwanda, Sierra Leone, Somalia, South Africa,<br />
Tanzania, Uganda, Zimbabwe) (CIE 1976; Tamaki and Miyara 1982;<br />
Whittle and Ferguson 1987; Veenakumari et al. 1995) and is also reported<br />
from Congo (Dhankar 1988) and Saudi Arabia (FAO 1982). It is not<br />
recorded from Australasia, Oceania, the Americas or Europe and was<br />
apparently absent until recently from the Philippines (CIE 1976), although<br />
Navasero (1983) has now reported it there. Between 1977 and 1987 there<br />
were 1291 interceptions <strong>of</strong> L. orbonalis at U.S. ports <strong>of</strong> entry, most on eggplant<br />
fruit in passenger baggage (Whittle and Ferguson 1987), and there<br />
must be significant risks also <strong>of</strong> its entry to Australia and the Pacific.
Biology<br />
Host plants<br />
4.11<br />
Leucinodes orbonalis<br />
187<br />
The flat, oval eggs are mostly laid at night, either singly or in groups <strong>of</strong> 2 to 4<br />
(and up to 200 per female), on the lower surface <strong>of</strong> young shoots, flower<br />
buds and calyces <strong>of</strong> developing fruits. They hatch in 4 days at an optimum<br />
temperature <strong>of</strong> 30¡C and relative humidity <strong>of</strong> 70% to 90%. Larval<br />
development occupies 14 days and pupal development 19 days. With a<br />
preoviposition period <strong>of</strong> 2 days, this results in a generation time <strong>of</strong> about one<br />
month. Details from three authors are shown in Table 4.11.1. Up to 9 larvae<br />
have been found in a single fruit and, when mature, pupate within a tough<br />
silken cocoon on the fruit, stem or among ground litter (Tamaki and Miyara<br />
1982; Khoo et al. 1991; Yin 1993). In the absence <strong>of</strong> fruit, larvae feed on the<br />
growing points <strong>of</strong> the plant. In the plains <strong>of</strong> India it occurs throughout the<br />
year but, at higher altitudes, cold weather interrupts its development and it<br />
overwinters as a larva in a silken cocoon, usually 1 to 3 cm below the soil<br />
surface. It is capable <strong>of</strong> surviving temperatures as low as Ð6.5¡C (Lal 1975).<br />
It thrives best under warm, moist monsoonal conditions. L. orbonalis can be<br />
reared in the laboratory on dried eggplant fruit or on a semi-synthetic diet<br />
(Islam et al. 1978, Patil 1990). Virgin females produce a pheromone that<br />
attracts males (Gunawardena 1992; Yasuda and Kawasaki 1994).<br />
In addition to eggplant, which is its main host, L. orbonalis is reported to<br />
feed on several other Solanum species, e.g. S. tuberosum (potato: shoots<br />
only) (Fletcher 1916; Mehto et al. 1980; Isahaque and Chaudhuri 1983),<br />
S. nigrum (black berry nightshade) (Nair 1967; Das and Patnaik 1970;<br />
Isahaque and Chaudhuri 1983 ), S. indicum,<br />
S. myriacanthum (shoots only)<br />
(Menon 1962; Isahaque and Chaudhuri 1983) and S. xanthocarpum (Menon<br />
1962). It has also been reported from tomato ( Lycopersicon esculentum)<br />
(Hargreaves 1937; Das and Patnaik 1970), potato ( Solanum tuberosum)<br />
and<br />
several unexpected plants, including cape gooseberry (Pillai 1922), green<br />
pea pods (Hussain 1925), mango shoots (Hutson 1930), cucumber, sweet<br />
potato and capsicum (Whittle and Ferguson 1987). Screening egg plant<br />
varieties for resistance to L. orbonalis has revealed several that are relatively<br />
resistant. Thick-skinned varieties appear to be more resistant (Patil and Ajri<br />
1993; Patel et al. 1995).
Table 4.11.1<br />
Development times, and other life history data (rounded) <strong>of</strong> Leucinodes orbonalis<br />
Reference Atwal and Verma 1972 Baang and Corey 1991 Mehto et al. 1983<br />
Temperature 20¡C 25¡C 30¡C 35¡C<br />
egg development(days) 9 6 4 3 5 6<br />
egg survival (%) 63 69 78 55<br />
larval development(days) 29 20 14 12 18 15<br />
larval survival (%) 53 72 69 48<br />
pupal development(days) 17 13 9 7 10 12<br />
pupal survival (%) 57 65 71 55<br />
longevity females(days) 11 7 6 3 3 8<br />
males(days) 9 7 4 2 2 4<br />
eggs/female 188 225 248 86 85Ð254 122<br />
188 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong>
Damage<br />
4.11<br />
Leucinodes orbonalis<br />
189<br />
All stages <strong>of</strong> eggplant are attacked by L. orbonalis,<br />
which is regarded as one<br />
<strong>of</strong> its major insect pests. Larvae bore into the tender shoots <strong>of</strong> both seedlings<br />
and after transplantation older plants, causing wilting and death <strong>of</strong> the<br />
growing tips. Later, they bore into flower buds and fruits. The damaged buds<br />
are shed and the fruits carry circular holes, sometimes plugged with frass.<br />
Such fruits are unmarketable. The yield loss varies with location and season<br />
and is greatest when temperature and humidity is high. Losses range from 20<br />
to 60% (Krishnaiah 1980; Dhankar 1988, Roy and Pande 1994) or even<br />
higher (Akhtar and Khawaja 1973; Lal 1991). It is reported that Vitamin C in<br />
bored fruit can be reduced by 60% (Hami 1955).<br />
There is an extensive literature dealing with the screening, mainly in<br />
India, for resistance <strong>of</strong> eggplant cultivars to L. orbonalis.<br />
Some cultivars are<br />
far less damaged than others, although no information is available on the<br />
genes involved. The less susceptible cultivars generally have one or more<br />
morphological characteristics, including compact vascular bundles in a thick<br />
layer, lignified cells and less area <strong>of</strong> pith in the shoots, tougher fruit skin and<br />
a tight calyx to hinder larval entry. Biochemical factors involved include a<br />
low protein and sugar content in resistant genotypes and a higher silica and<br />
crude fibre content in the shoots, which adversely affects growth rate, pupal<br />
period, survival, sex ratio and fecundity. The wild relatives <strong>of</strong> Solanum<br />
melongena that are not attacked by L. orbonalis <strong>of</strong>ten have a high alkaloid<br />
content, which may be responsible, but this attribute would not be desirable<br />
in an edible product (Dhankar 1988).<br />
Natural enemies<br />
Table 4.11.2 lists the natural enemies <strong>of</strong> L. orbonalis.<br />
It is striking that most<br />
are from India and Sri Lanka, that the records from Malaysia and the<br />
Philippines are the only ones from <strong>Southeast</strong> Asia and that the species there<br />
have not been reported elsewhere. Although it is possible that some <strong>of</strong> the<br />
Indian and Sri Lankan species have a restricted distribution, the lack <strong>of</strong><br />
records from elsewhere probably means that the necessary investigations<br />
have not been carried out.
Table 4.11.2<br />
Natural enemies <strong>of</strong> Leucinodes orbonalis<br />
Species Stage attacked % parasitisation Country Reference<br />
NEUROPTERA<br />
CHRYSOPIDAE<br />
Chrysopa kulingensis<br />
DERMAPTERA<br />
egg<br />
larva<br />
12.5<br />
2-4<br />
China Yang 1982<br />
UNIDENTIFIED (15) Philippines Navasero 1983<br />
DIPTERA<br />
SARCOPHAGIDAE<br />
Amobia sp.<br />
TACHINIDAE<br />
Malaysia Thompson 1946<br />
Pachyophthalmus sp. Malaysia Corbett 1929; Yunus & Ho 1980<br />
Pseudoperichaeta sp. 5.7 India Patel et al. 1971<br />
Sturmia parachrysops<br />
India Thompson 1946<br />
UNIDENTIFIED<br />
HYMENOPTERA<br />
BRACONIDAE<br />
pupa 5Ð8 China Yang 1982<br />
Apanteles sp. 3.1-11.1 Philippines Navasero 1983<br />
Bracon greeni<br />
larva; ecto lab only India Venkatraman et al. 1948<br />
Bracon sp. larva; ecto 9.2Ð28.1 India Tewari & Sardana 1987a<br />
Campyloneura sp. larva 1Ð2 India Tewari & Moorthy 1984<br />
Chelonus sp.<br />
15.5<br />
Phanerotoma sp. 4.6<br />
7.4<br />
Phanerotoma sp. nr.<br />
hindecasisella<br />
Philippines<br />
Sri Lanka<br />
India<br />
Sri Lanka<br />
Navasero 1983<br />
Sandanayake & Edirisinghe 1992<br />
Patel et al. 1971<br />
Sandanayake & Edirisinghe 1992<br />
larva 1Ð2 India Patel et al. 1971;<br />
Tewari & Moorthy 1984<br />
190 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong>
Table 4.11.2 (contÕd) Natural enemies <strong>of</strong> Leucinodes orbonalis<br />
Species<br />
HYMENOPTERA<br />
Stage attacked % parasitisation Country Reference<br />
CHALCIDIDAE<br />
Brachymeria lasus<br />
Philippines Navasero 1983<br />
Brachymeria sp.<br />
EULOPHIDAE<br />
Philippines Navasero 1983<br />
Dermatopelte<br />
(= Dermatopolle)<br />
sp.<br />
ICHNEUMONIDAE<br />
pupa 16 China Yang 1982<br />
Cremastus hapaliae<br />
Malaysia Yunus & Ho 1980<br />
Eriborus argenteopilosus larva 0.5Ð2 India Tewari & Sardana 1987b<br />
Itamoplex sp. pupa 9Ð15 India Verma & Lal 1985<br />
Pristomerus testaceus<br />
larva; ecto India Ayyar 1927<br />
Trathala flavoorbitalis<br />
Trathala striata<br />
Xanthopimpla punctata<br />
BACTERIUM<br />
BACULOVIRUS<br />
larva 36.2<br />
3.6Ð9.1<br />
12.9Ð18.2<br />
Sri Lanka<br />
India<br />
India<br />
India<br />
Malaysia<br />
Sandanayake & Edirisinghe 1992<br />
Mallik et al. 1989<br />
Naresh et al. 1986a, b,<br />
Patel et al. 1967<br />
Yunus & Ho 1980<br />
Malaysia C.L. Tan pers. comm. 1994<br />
Philippines Navasero 1983<br />
larva 2Ð3 China Yang 1982<br />
larva 1.1Ð6.4 India Tewari & Singh 1987<br />
4.11<br />
Leucinodes orbonalis<br />
191
192 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
CHINA<br />
The life history <strong>of</strong> L. orbonalis was studied by Yin (1993), in Hunan where<br />
up to 6 generations were completed annually and overwintering occurred in<br />
the pupal stage. Yang (1982) recorded two pupal parasitoids, the<br />
polyembryonic wasp Dermatopelte sp., causing 16% parasitisation (and<br />
producing 12 to 21 individuals per host pupa) and an unidentified dipteran<br />
causing 5% to 8% mortality. A predator, Chrysopa kulingensis consumed<br />
12.5% <strong>of</strong> eggs and 2% to 4% <strong>of</strong> larvae. A disease, presumably <strong>of</strong> bacterial<br />
origin, killed 2% to 3% <strong>of</strong> larvae.<br />
INDIA, SRI LANKA<br />
The ichneumonid Trathala flavoorbitalis appears to be the most effective<br />
parasitoid so far recorded, with an average parasitisation rate in Sri Lanka <strong>of</strong><br />
36.2% (Sandanayake and Edirisinghe 1992). In Hanyana, India, Naresh et al.<br />
(1986a) recorded rates from 12.9% to 18.2% and in Bihar, Mallik et al.<br />
(1989) 3.6% to 9.1%. These were higher than the combined rates <strong>of</strong> 1% to<br />
2% by Phanerotoma sp. and Campyloneura sp.. T. flavoorbitalis is a very<br />
widespread species and attacks the larvae <strong>of</strong> many species <strong>of</strong> Lepidoptera.<br />
Bracon sp. from near Bangalore, India, with a parasitisation rate ranging up<br />
to 28.1% (Tewari and Sardana 1987a), Chelonus sp. ranging up to 5.5%<br />
(Sandanayake and Edirisinghe 1992) and Itamoplex sp. up to 15% (Verma<br />
and Lal 1985) are all capable <strong>of</strong> producing significant mortality.<br />
PHILIPPINES<br />
A dermapteran larval predator and 5 parasitoids (the braconids Apanteles sp.<br />
(on larvae) and Chelonus sp. (on pupae), the chalcidids Brachymeria lasus<br />
(= B. obscurata) and Brachymeria sp. (larvae and pupae), and the<br />
ichneumonid Xanthopimpla punctata) were found attacking L. orbonalis in<br />
the field. Apanteles sp. and the dermapteran were the most abundant<br />
(Navasero 1983). It appears that L. orbonalis has only been recognised in the<br />
Philippines since the early 1970s.<br />
Attempts at biological control<br />
There have been none.<br />
Major natural enemies<br />
Bracon sp. Hym.: Braconidae<br />
This larval ectoparasitoid was found near Bangalore, India attached to the<br />
thorax <strong>of</strong> the host larva. It pupated in a silk cocoon inside the tunnel made by<br />
its host and caused parasitisation ranging from 9.2% to 28.1%. It was<br />
regarded as a promising parasitoid (Tewari and Sardana 1987a).
4.11 Leucinodes orbonalis 193<br />
Itamoplex sp. Hym.: Ichneumonidae<br />
Adult Itamoplex sp., 8 to 10 mm in length, were reported from Kulu Valley,<br />
Himachal Pradesh, India where the winter temperature drops as low as Ð8¡C.<br />
L. orbonalis overwinters in the larval (?prepupal) stage in an earthen cocoon<br />
attached to the host plant, usually 1 to 3 cm below the soil surface. The<br />
parasitoid emerged from 9% to 15% <strong>of</strong> these cocoons. Itamoplex (= Cryptus)<br />
sp. is recorded attacking a range <strong>of</strong> host Lepidoptera in cocoons (Verma and<br />
Lal 1985).<br />
Trathala (= Cremastus) flavoorbitalis Hym.: Ichneumonidae<br />
This is a widespread, non-specific parasitoid <strong>of</strong> lepidopterous larvae. It<br />
occurs naturally in India, Japan, Myanmar, Sri Lanka, the Philippines and<br />
Singapore and has been established in Canada, Hawaii and continental USA<br />
for biological control <strong>of</strong> several important lepidopterous pests. It is recorded<br />
from at least 5 families <strong>of</strong> Lepidoptera, involving over 40 different hosts,<br />
most <strong>of</strong> them pest species (Bradley and Burgess 1934). It is not known<br />
whether there are strains that prefer to attack particular hosts.<br />
T. flavoorbitalis is recorded from L. orbonalis in India and also in Sri<br />
Lanka where L. orbonalis is its major host and where an average<br />
parasitisation level <strong>of</strong> 36.2% is reported (Sandanayake and Edirisinghe<br />
1992, 1993). In Hissar, India, Trathala was the only parasitoid <strong>of</strong><br />
L. orbonalis, with levels <strong>of</strong> attack on larvae ranging from 13.2% to 18.2% in<br />
winter to 12.9% in summer at a time when 95.2% <strong>of</strong> fruit was infested<br />
(Naresh et al. 1986a, b).<br />
T. flavoorbitalis females commence ovipositing 4 days after emergence,<br />
with a preference for 3rd, 4th and 5th instar host larvae. In the laboratory,<br />
only a fraction <strong>of</strong> 1st instar larvae are stung and all soon die from the<br />
encounter, a fate shared by about half the 2nd instar larvae that are stung. In<br />
later instars 68Ð91% were stung, but without early mortality. Not all <strong>of</strong> these<br />
received eggs, although some received up to 5, with only one parasitoid larva<br />
developing beyond the first instar. Parasitoid development time from egg to<br />
adult was about 23 days at 28¡C. Successful development occurred in 53%<br />
<strong>of</strong> the 3rd, 57% <strong>of</strong> the 4th and 41% <strong>of</strong> 5th instar host larvae, adult wasps<br />
emerging after pupation <strong>of</strong> the host. In the field, the parasitoid attacks the<br />
host larva by inserting its ovipositor into the hole bored into the fruit and<br />
through which the larva pushes out frass (Bradley and Burgess 1934; Naresh<br />
et al. 1986a; Sandanayake and Edirisinghe 1992, 1993).
194 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Comments<br />
It is not clear how many <strong>of</strong> the natural enemies are sufficiently host specific<br />
to be confidently transferred as biological control agents although, where<br />
alternative hosts are known, these are also pests. Thus, Eriborus<br />
argenteopilosus has a wide host range, including Condica (= Prospalta)<br />
capensis, which attacks safflower and sunflower, Helicoverpa armigera,<br />
and Spodoptera exigua (Tewari and Sardana 1987a,b). The ichneumonid<br />
Pristomerus testaceus has been bred from the brinjal stem-borer Euzophera<br />
ferticella (Ayyar 1927). It is not known whether the ichneumonid<br />
Phanerotoma sp. nr hindecasisella is a distinct species. True<br />
P. hindecasisella has been reported from several lepidopterous families in<br />
India or Sri Lanka: Gelechiidae (Dichomeris eridantis), Noctuidae (Earias<br />
insulana), Pyralidae (Eutectona (= Pyrausta) macheralis, Hendecasis<br />
duplifascialis, Maruca vitrata (= M. testulalis), Nephopterix rhodobasalis,<br />
Syllepte derogata) and Tortricidae (Leguminivora (= Cydia) ptychora)<br />
(Thompson 1953; Fellowes and Amarasena 1977; Kumar et al. 1980;<br />
Tewari and Moorthy 1984).<br />
Bracon greeni is best known as a primary ectoparasitoid <strong>of</strong> the<br />
lepidopterous lac predator Eublemma amabilis (Noctuidae) and has not been<br />
reported to parasitise any other host in nature. However, under laboratory<br />
conditions, it was successfully reared on L. orbonalis (Venkatraman et al.<br />
1948).<br />
Although entomopathogenic nematodes have not been recorded<br />
attacking L. orbonalis in the field, Steinernema (=Neoaplectana)<br />
carpocapsae (DD136 strain) produced 73.3% mortality <strong>of</strong> larvae in the<br />
laboratory in 72 hours (Singh and Bardhan 1974).<br />
The weight <strong>of</strong> evidence suggests that L. orbonalis originated in India and<br />
spread into <strong>Southeast</strong> Asia. It is most surprising that it has only<br />
comparatively recently become a pest Ñ and a serious one Ñ in the<br />
Philippines which, in 1990, had 16 000ha under egg plant and produced<br />
113 000 tonnes, second only to Indonesia in production in <strong>Southeast</strong> Asia<br />
(FAO 1991). In view <strong>of</strong> the steady spread around the world <strong>of</strong> so many other<br />
pests it is also surprising that Australia, the Pacific, the Americas and Europe<br />
are still free from L. orbonalis.<br />
It might well be assumed that not all <strong>of</strong> its natural enemies in India have<br />
accompanied it during its spread. However, reports <strong>of</strong> high damage levels to<br />
susceptible egg plant cultivars in India do not provide much confidence that<br />
the natural enemies there are particularly effective, unless their efficacy is,<br />
perhaps, reduced by insecticide applications or so-far-unreported<br />
hyperparasitoids.
4.11 Leucinodes orbonalis 195<br />
Nevertheless, parasitisation in Sri Lanka is quite impressive, with<br />
Trathala flavoorbitalis averaging 36.2%, Chelonus sp. 15.5% and<br />
Phanerotoma sp. 7.4%. The combined average rate <strong>of</strong> 59.1% would<br />
certainly be capable <strong>of</strong> causing an important lowering <strong>of</strong> moth populations,<br />
particularly if associated with the high levels <strong>of</strong> host plant resistance that are<br />
available. It is probable that the widespread high rates <strong>of</strong> application <strong>of</strong><br />
broad spectrum insecticides currently used are preventing natural enemies<br />
from exerting much effect. An investigation <strong>of</strong> what natural enemies <strong>of</strong><br />
L. orbonalis are already present in <strong>Southeast</strong> Asia is required to enable a<br />
decision on the attractiveness <strong>of</strong> this pest as a biological control target. It is<br />
probably safe to conclude that T. flavoorbitalis, the most effective parasitoid<br />
so far reported, is present in most, if not all, <strong>of</strong> <strong>Southeast</strong> Asia, but it would<br />
be relevant, in relation to possible host-preferring strains, to determine<br />
whether it attacks L. orbonalis throughout the region.<br />
It is interesting that the closely related South American Neoleucinodes<br />
elegantalis has a quite different suite <strong>of</strong> parasitoids. Of 2500 larvae collected<br />
in the field 1.6% were parasitised by the encyrtid Copidosoma sp. and 0.08%<br />
by the tachinid Lixophaga sp. Of 527 pupae, 0.38% were parasitised by the<br />
ichneumon Calliephialtes sp. The fungus Beauveria sp. caused 55%<br />
mortality. Trichogramma sp. parasitised 89% <strong>of</strong> eggs laid on egg plant and<br />
tomato (Plaza et al. 1992). Of these species, Beauveria and the<br />
Trichogramma sp. might be relevant to the <strong>Southeast</strong> <strong>Asian</strong> scene.
4.12 Nezara viridula<br />
India<br />
Myanmar<br />
++<br />
20°<br />
Laos<br />
+<br />
0°<br />
20°<br />
China<br />
++<br />
Thailand<br />
+<br />
Cambodia<br />
+<br />
Vietnam<br />
++<br />
P<br />
+ Brunei<br />
Malaysia<br />
+<br />
Singapore<br />
+<br />
Indonesia<br />
Taiwan<br />
++<br />
P<br />
Philippines<br />
Australia<br />
Papua<br />
New Guinea<br />
++<br />
197<br />
Nezara viridula is probably native to the Ethiopian region, but is now dispersed widely<br />
throughout the warmer regions <strong>of</strong> the world.<br />
There have been a number <strong>of</strong> major successes with biological control <strong>of</strong> the green<br />
vegetable bug, particularly with an egg parasitoid, Trissolcus basalis.<br />
<strong>Control</strong> has been<br />
supplemented in Hawaii by two parasitoids <strong>of</strong> adults, Trichopoda pilipes and T. pennipes,<br />
which, however, have failed to establish in most other places.<br />
The main areas where biological control using T. basalis has been unsuccessful are<br />
<strong>of</strong>ten associated with extensive plantings <strong>of</strong> soybean. It has been suggested that the<br />
surface hairyness <strong>of</strong> most soybean cultivars reduces the effectiveness <strong>of</strong> this parasitoid. If<br />
this proves to be correct the way is open for the selection <strong>of</strong> cultivars with this attribute<br />
modified.<br />
Over 80 parasitoids <strong>of</strong> N. viridula are known, amongst which there are several<br />
potentially valuable, untried species that are clearly worthy <strong>of</strong> investigation.<br />
N. viridula is an attractive target for biological control, especially where it is a problem in<br />
an area not closely associated with extensive soybean plantings.<br />
20°<br />
0°<br />
20°
198 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Nezara viridula (Linnaeus)<br />
Rating<br />
Origin<br />
Hemiptera, Pentatomidae<br />
green vegetable bug, southern green stink bug (USA)<br />
<strong>Southeast</strong> Asia China Southern and Western Pacific<br />
10 ++ Myan, Viet ++ 14 ++ Cook Is, Fr P, Niue,<br />
PNG, Sam<br />
+ Thai, Laos, Camb,<br />
Msia,<br />
Distribution<br />
+ Fiji, Kiri, N Cal,<br />
A Sam<br />
Sing, Indo<br />
P Brun, Phil P FSM, Guam, Sol Is,<br />
Tong, Van<br />
This account updates the chapter on Nezara viridula in Waterhouse and<br />
Norris (1987), and the valuable account <strong>of</strong> Jones (1988), in relation to<br />
prospects for biological control.<br />
The locality <strong>of</strong> the holotype Ôin IndiisÕ (Linnaeus 1758) has been interpreted<br />
as India or the East Indies. However it may well have been brought there by<br />
European travel since an analysis <strong>of</strong> genetic colour variants and the relative<br />
abundance <strong>of</strong> fairly specific insect parasitoids led Hokkanen (1986) and<br />
Jones (1988) to the conclusion that the original home <strong>of</strong> N. viridula was in<br />
the Ethiopian (Afrotropical) or Mediterranean region, rather than in<br />
<strong>Southeast</strong> Asia (Yukawa and Kiritani 1965). Furthermore, Africa is<br />
considered to be the centre <strong>of</strong> the genus Nezara (Freeman 1940).<br />
Azores, Canary Is, Bermuda, the Mediterranean littorale, most <strong>of</strong> Africa and<br />
the Middle East, Madagascar, Mauritius, Reunion, Rodriguez, Seychelles,<br />
Asia (exclusive <strong>of</strong> desert areas and those with very cold winters) Korea and<br />
southern Japan, <strong>Southeast</strong> Asia, Papua New Guinea (including Bismarck<br />
Archipelago), most <strong>of</strong> the oceanic Pacific nations (Butcher 1981), Australia,<br />
New Zealand, Hawaii, southern USA, Mexico and other Central American<br />
countries, West Indies generally, and much <strong>of</strong> South America (Anon. 1970).<br />
It is not known to occur in Tokelau, Tuvalu or the Marquesas (Waterhouse<br />
1985, 1997).
Biology<br />
Damage<br />
4.12<br />
Nezara viridula<br />
199<br />
The entirely green colour form smaragdula <strong>of</strong> N. viridula is the one that<br />
occurs widely in the Pacific area. In other areas <strong>of</strong> the world it is<br />
accompanied by several other colour forms (Yukawa and Kiritani 1965).<br />
The barrel-shaped eggs are usually laid at night in neat rafts, commonly <strong>of</strong><br />
80 to 120 or more eggs and they are cemented firmly to one another and to<br />
the sheltered surface <strong>of</strong> a leaf. The eggs hatch in 4 to 9 days, and the newly<br />
emerged nymphs remain together near the eggshells for a day or two, a<br />
degree <strong>of</strong> gregariousness persisting also in the next instar or two. There are<br />
five nymphal stages before the adult emerges, 24 to 60 days after hatching,<br />
depending on temperature. At 25Ð28¡C, 55Ð65% RH and 14 hours daylight<br />
the development periods (in days) were: egg, 4.8; 1st instar nymph, 3.8; 2nd<br />
instar, 5.2; 3rd instar, 4.5; 4th instar, 6.4; 5th instar, 11.9 (Harris and Todd<br />
1980). The nymphal stages are multicoloured, but the adults are a uniform<br />
green in the form smaragdula.<br />
The adults can fly strongly. There may be<br />
four generations in a year in coastal New South Wales, and perhaps more in<br />
areas with no perceptible winter.<br />
The adult bugs live up to 3 weeks in hot weather. In regions with a cold<br />
winter those <strong>of</strong> the autumn generation may live much longer, hibernating in<br />
debris, under bark, or in buildings, inactive and non-reproductive. Such<br />
hibernating bugs change colour from green to brown. In areas with a less<br />
severe winter the still-green adults may remain active, although nonreproductive.<br />
Waite (1980) showed that the adults and nymphs tend to bask<br />
exposed on the surface <strong>of</strong> the plant canopy in the early daylight<br />
hoursÑbehaviour that can be availed <strong>of</strong> in applying chemical control<br />
measures (Kamal 1937; Clausen 1978; Hely et al. 1982; Singh and Rawat<br />
1982; Todd 1989).<br />
When not controlled by chemicals or natural enemies, this bug can be a<br />
serious pest <strong>of</strong> a very wide range <strong>of</strong> crops and ornamental plants. In Australia<br />
there are recommendations for its chemical control on beans, cucurbits,<br />
peas, potatoes, tomatoes, passionfruit, groundnuts, sorghum, soybeans,<br />
sunflowers and tobacco (Anon. 1967, 1979b; Miller et al. 1977), but it also<br />
attacks maize, crucifers, spinach, lucerne and many other legumes, grapes,<br />
oranges and many other fruits and seeds, and macadamia (Ironside 1979; La<br />
Croix 1986) and pecan nuts (Seymour and Sands 1992). Undoubtedly there<br />
would be recommendations for a much wider range <strong>of</strong> cultivated plants were<br />
it not for the fact that biological control is effective now in many situations
200 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
(Anon. 1967; Hely et al. 1982). In other countries it can be a pest <strong>of</strong> rice,<br />
sesame and other grains, lablab, guavas, cowpeas, capsicums, cotton and<br />
many other plants. It also infests and breeds on many weeds, which afford it<br />
harborage.<br />
The attack <strong>of</strong> the bugs is concentrated chiefly on fruits and fruiting<br />
bodies, which, through the removal <strong>of</strong> sap and the injection <strong>of</strong> saliva, show<br />
discoloration, malformation, stunting and shrivelling. Heavily attacked<br />
tomatoes, for instance, are repulsive and inedible and even lightly attacked<br />
ones are unmarketable. Passlow and Waite (1971), Goodyer (1972) and<br />
Romano and Kerr (1977) attribute serious losses in Australian soybean crops<br />
to the attack <strong>of</strong> this pest. Miller et al. (1977) measured the yield reduction on<br />
soybeans and showed that an important effect <strong>of</strong> the bugs feeding was a<br />
reduction in the germinability <strong>of</strong> seed. On soybean in Brazil, Corso et al.<br />
(1975) showed that green vegetable bug attack affected pod development,<br />
increased pod fall, and reduced the number <strong>of</strong> seeds per pod. Link et al.<br />
(1973) demonstrated reductions in germination percentage and oil content<br />
and an increase in relative protein content in heavily damaged seeds. A list <strong>of</strong><br />
references dealing with N. viridula and its association with soybeans has<br />
been prepared by DeWitt and Godfrey (1972).<br />
Some cultivars <strong>of</strong> soybeans are less damaged by N. viridula than others<br />
and a strain <strong>of</strong> soybean that appears to have a high level <strong>of</strong> resistance has<br />
been selected (Gilman et al. 1982). Mild antibiosis and non-preference were<br />
factors contributing to resistance (Kester et al. 1984). The adverse effects <strong>of</strong><br />
the genotype (PI 171444) on the biology <strong>of</strong> a parasite Telenomus chloropus<br />
attacking the eggs <strong>of</strong> N. viridula feeding on it have been studied by Orr et al.<br />
(1985b).<br />
The green vegetable bug has been shown to carry spores <strong>of</strong> fungal<br />
diseases from plant to plant (Corso et al. 1975) and to transmit plant<br />
pathogens during feeding (Kaiser and Vakili 1978).<br />
Natural enemies<br />
Introduced parasitoids have brought about successful biological control <strong>of</strong><br />
this pest in a number <strong>of</strong> countries and there is an extensive literature on the<br />
subject. Nine species <strong>of</strong> parasite attacking Nezara viridula were listed by<br />
Thompson (1944), 27 by Hokkanen (1986) and 57 by Jones (1988). At least<br />
eighty are now included in Table 4.12.1 which is a modification and<br />
extension <strong>of</strong> that in Jones (1988).<br />
The species belong to two families <strong>of</strong> Diptera and six families <strong>of</strong><br />
Hymenoptera. It is notable from the entries that relatively little is known <strong>of</strong><br />
the parasitoids <strong>of</strong> N. viridula in the Ethiopian or Mediterranean regions,
4.12<br />
Nezara viridula<br />
201<br />
which constitute the presumed area <strong>of</strong> origin <strong>of</strong> the green vegetable bug. Egg<br />
parasitoids are the most numerous and all are Hymenoptera, whereas<br />
nymphal and adult parasitoids are, with one exception, all Diptera. Two<br />
hyperparasitoids <strong>of</strong> the major egg parasitoid Trissolcus basalis are known<br />
from Australia, (both are species <strong>of</strong> Acroclissoides:<br />
Clarke and Seymour<br />
1992) and one hyperparasitoid <strong>of</strong> the fly Trichopoda pennipes from Hawaii<br />
( Exoristobia philippinensis:<br />
Davis and Krauss 1965). Predators are not dealt<br />
with as all are known to be, or suspected as being, widely polyphagous and<br />
hence unlikely to be approved for introduction to new areas. Nevertheless,<br />
they play a significant role in maintaining N. viridula populations at low<br />
levels. For example, in one study in soybeans in Louisiana, Stam et al.<br />
(1987) found 18 insect and 6 spider species to be predators and that they<br />
were responsible for 33.6% <strong>of</strong> the total mortality <strong>of</strong> N. viridula that occurred<br />
from egg to adult.<br />
The Scelionidae is the most important <strong>of</strong> the six families <strong>of</strong><br />
hymenopterous egg parasitoids. In it Trissolcus basalis is not only the most<br />
important species, but also the most widespread. It attacks the eggs <strong>of</strong> a<br />
number <strong>of</strong> other pentatomids, but appears to have a preference for<br />
N. viridula:<br />
it is the dominant parasitoid <strong>of</strong> N. viridula eggs wherever it<br />
occurs. Many other species are listed, in particular in the genera Trissolcus,<br />
Telenomus, Ooencyrtus and Gryon,<br />
but a number appear to have no close<br />
relationship with N. viridula and are unlikely to be <strong>of</strong> value as potential<br />
biological control agents. Some drought-resistant species from the<br />
Mediterranean may prove useful. The better known <strong>of</strong> the more promising<br />
species are discussed later.<br />
The tachinid parasitoids <strong>of</strong> adult N. viridula also oviposit on the cuticle<br />
<strong>of</strong> 4th and 5th instar nymphs. Often these eggs are shed before hatching<br />
along with the cuticle at moulting but, if hatching and penetration <strong>of</strong> the bug<br />
occurs, the parasitoid larva matures in the adult. More <strong>of</strong> the species <strong>of</strong><br />
tachinids are native to South America than elsewhere. They were clearly<br />
dependent upon other pentatomids before the arrival <strong>of</strong> N. viridula,<br />
but<br />
several now appear to have a preference for it. The most promising <strong>of</strong> the<br />
three tachinid parasitoids occuring in the Ethiopian region is the widespread<br />
Bogosia antinorii,<br />
which is known only from N. viridula (van Emden 1945;<br />
Barraclough 1985).<br />
A picorna-like and a toti-like virus are known from N. viridula<br />
(Williamson and Wechmar 1992, 1995).
Table 4.12.1<br />
Parasitoids <strong>of</strong> the green vegetable bug, Nezara viridula (modified after Jones 1988)<br />
Parasitoid Geographic range Known host relations Selected references<br />
DIPTERA<br />
SARCOPHAGIDAE<br />
Sarcodexia innota<br />
Sarcodexia sternodontis<br />
TACHINIDAE<br />
Bogosia antinorii<br />
Cylindromyia rufifemur<br />
Ectophasia crassipennis<br />
Ectophasiopsis arcuata<br />
Euclytia flava<br />
Gymnosoma clavata<br />
Gymnosoma kuramanum<br />
Gymnosoma rotundata<br />
Trichopoda giacomellii<br />
(= Trichopoda nigrifrontalis,<br />
= T. gustavoi<br />
= Eutrichopodopsis nitens)<br />
Trichopoda lanipes<br />
Trichopoda pennipes<br />
Southern USA Two records ex N. viridula;<br />
wide host range as primary<br />
parasitoid and scavenger<br />
Widespread in Africa Recorded only ex N. viridula<br />
Australia One record ex N. viridula<br />
Italy Bred ex N. viridula<br />
Chile Well adapted to N. viridula<br />
Drake 1920; Temerak &<br />
Whitcomb 1984<br />
Hokkanen 1986<br />
Greathead 1966, 1971;<br />
Barraclough 1985<br />
Cantrell 1984<br />
Colazza & Bin 1995<br />
Jones 1988<br />
USA Generalist parasitoid Aldrich 1995<br />
Palaearctic, Israel One record ex N. viridula<br />
Herting 1960<br />
Japan Takano 1956<br />
Palaearctic Wide host range; attacks Nezara spp. in Japan Kiritani et al. 1963; Kiritani<br />
& Sasaba 1969<br />
Argentina, Brazil, Well adapted to N. viridula<br />
Blanchard 1966; Mallea et<br />
Colombia, Paraguay<br />
al. 1968; Gastal 1977a,b;<br />
Liljesthršm 1980, 1981;<br />
Ferreira 1984<br />
Southern USA One record ex N. viridula;<br />
attacks other species Drake 1920<br />
North America, Hawaii Well adapted to N. viridula<br />
Drake 1920; Todd & Lewis<br />
1976; Jones 1979;<br />
Buschman & Whitcomb<br />
1980; McPherson et al.<br />
1982<br />
202 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong>
Table 4.12.1 (contÕd) Parasitoids <strong>of</strong> the green vegetable bug, Nezara viridula (modified after Jones 1988)<br />
Parasitoid<br />
DIPTERA<br />
Geographic range Known host relations Selected references<br />
TACHINIDAE (contÕd)<br />
Trichopoda pilipes<br />
West Indies, Hawaii Well adapted to N. viridula<br />
Trichopoda sp. Uruguay One record ex N. viridula<br />
Myers 1931; Nishida 1966;<br />
Davis 1967<br />
Guido & Ruffinelli 1956<br />
HYMENOPTERA<br />
EULOPHIDAE<br />
Pleurotropitiella albipes<br />
EURYTOMIDAE<br />
Argentina Esquivel 1950<br />
Neorileya sp.<br />
EUPELMIDAE<br />
Brazil Recorded only ex N. viridula Ferreira 1981, 1984, 1986<br />
Anastatus bifasciatus Italy Bred ex N. viridula Colazza & Bin 1990<br />
Anastatus dasyni Malaysia Pentatomidae, Coreidae; described ex N. viridula van der Vecht 1933<br />
Anastatus japonicus East Asia Lepidoptera, Heteroptera; produces only males in<br />
N. viridula<br />
Hokyo et al. 1966b; Kiritani<br />
& Sasaba 1969<br />
Anastatus sp. Thailand Emerged ex imported eggs <strong>of</strong> N. viridula Jones 1988<br />
Anastatus sp. Southern USA Two records ex N. viridula Jones 1988<br />
Anastatus sp. Australia<br />
(Queensland)<br />
One record ex N. viridula Seymour & Sands 1993<br />
Unidentified sp.<br />
ENCYRTIDAE<br />
Hawaii ex N. viridula on macadamia Jones 1992<br />
Hexacladia hilaris USA One record ex N. viridula Buschman & Whitcomb<br />
1980<br />
Ooencyrtus californicus California ex N. viridula and other pentatomids H<strong>of</strong>fmann et al. 1991<br />
4.12<br />
Nezara viridula<br />
203
Table 4.12.1 (contÕd) Parasitoids <strong>of</strong> the green vegetable bug, Nezara viridula (modified after Jones 1988)<br />
Parasitoid<br />
HYMENOPTERA<br />
Geographic range Known host relations Selected references<br />
ENCYRTIDAE (contÕd)<br />
Ooencyrtus fecundus Morocco VoegelŽ 1961<br />
Ooencyrtus johnsoni California ex N. viridula and other pentatomids H<strong>of</strong>fmann et al. 1991<br />
Ooencyrtus malayensis Malaysia, Philippines Pentatomidae, Coreidae, Lepidoptera van der Vecht 1933; Jones<br />
et al. 1983<br />
Ooencyrtus nezarae East Asia Coreidae, Pentatomidae, Plataspidae; not uncommon<br />
on N. viridula in Japan<br />
Hokyo & Kiritani 1966<br />
Ooencyrtus pityocampae Italy Breeds in eggs <strong>of</strong> N. viridula and other pentatomids in<br />
the laboratory<br />
Tiberi et al. 1991<br />
Ooencyrtus submetallicus West Indies, Central<br />
and South America<br />
Pentatomidae, Coreidae Gahan 1927; Lee 1979; de<br />
Santis 1985; Ferreira 1986<br />
Ooencyrtus trinidadensis West Indies, Argentina Pentatomidae, Coreidae Davis & Krauss 1963;<br />
Davis 1967; de Santis 1985<br />
Ooencyrtus sp. Brazil One record ex N. viridula Ferreira 1986<br />
Ooencyrtus sp. Thailand Emerged ex imported eggs <strong>of</strong> N. viridula Jones 1988<br />
Ooencyrtus sp. Philippines Possibly is O. malayensis Davis 1967; Corpuz 1969<br />
Ooencyrtus sp. France One record; recovered ex other pentatomids Jones 1988<br />
Ooencyrtus sp. Italy Bred ex N. viridula Colazza & Bin 1995<br />
Ooencyrtus sp. (spp.?) Southern USA Taxonomy and host range not known Drake 1920; Buschman &<br />
Whitcomb 1980;<br />
Jones 1988<br />
Xenoencyrtus hemipterus<br />
(= X. niger)<br />
Australia Seymour & Sands 1993<br />
Xenoencyrtus rubricatus Australia Described ex N. viridula Riek 1962<br />
Xenoencyrtus sp. Australia Bred ex N. viridula Forrester 1979<br />
204 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong>
Table 4.12.1 (contÕd) Parasitoids <strong>of</strong> the green vegetable bug, Nezara viridula (modified after Jones 1988)<br />
Parasitoid<br />
HYMENOPTERA<br />
Geographic range Known host relations Selected references<br />
PTEROMALIDAE<br />
Pteromalus sp. Egypt One record ex N. viridula Adair 1918<br />
3 spp. Brazil Ferreira & Moscardi 1995<br />
SCELIONIDAE<br />
Gryon fulviventris Africa, Asia Pentatomoidea; attacks N. viridula only in Thailand Anderson 1919; Dry 1921,<br />
Jones 1988<br />
Gryon japonicum Japan, Brazil native <strong>of</strong> Japan Kishino & Teixeira 1994<br />
Gryon obesum Southern USA, Brazil Records ex N. viridula; attacks other pentatomids Buschman & Whitcomb<br />
1980; Masner 1983;<br />
H<strong>of</strong>fmann et al. 1991;<br />
Correa & Moscardi 1995<br />
Gryon sp. Australia Minor attack on N. viridula Titmarsh 1979<br />
Gryon sp. Laos One record ex N. viridula; may be G. fulviventris Grist & Lever 1969; Dean<br />
1978a,b<br />
Gryon sp. India One record ex N. viridula, may be G. fulviventris Yadava et al. 1982<br />
Psix lacunatus Asia, Australia Pentatomidae, Scutelleridae; ex N. viridula in Pakistan Johnson & Masner 1985<br />
Psix striaticeps Africa, India<br />
Togo<br />
Pentatomidae; recorded once ex ÔNezaraÕ<br />
Common on N. viridula and 2 other pentatomids<br />
Telenomus chloropus Palearctic Pentatomidae; major parasitoid <strong>of</strong> Nezara spp. in<br />
E. Asia; females only in Japan<br />
Fouts 1934; Johnson &<br />
Masner 1985;<br />
Poutouli 1995<br />
Kiritani & Hokyo 1962;<br />
Hokyo & Kiritani 1963;<br />
Johnson 1984a<br />
Telenomus comperei Philippines Cadapan & Alba 1987<br />
Telenomus cristatus Southern USA, West<br />
Indies<br />
Known only ex N. viridula and Acrosternum hilare Johnson 1984a; Orr et al.<br />
1986<br />
4.12 Nezara viridula 205
Table 4.12.1 (contÕd) Parasitoids <strong>of</strong> the green vegetable bug, Nezara viridula (modified after Jones 1988)<br />
Parasitoid<br />
HYMENOPTERA<br />
Geographic range Known host relations Selected references<br />
SCELIONIDAE (contÕd)<br />
Telenomus cyrus Java, Philippines,<br />
Taiwan<br />
Descr. ex N. viridula; host relations unknown Nixon 1936; Taiwan<br />
Agricultural Research<br />
Institute 1984; Jones 1988<br />
Telenomus gifuensis East Asia Pentatomidae, Coreidae; not well adapted to N. viridula Hidaka 1958; Hokyo &<br />
Kiritani 1963<br />
Telenomus mormideae South America Attacks N. viridula and other pentatomids Ferreira 1986; Liljestršm &<br />
Bernstein 1990<br />
Telenomus pacificus Philippines Cadapan & Alba 1987<br />
Telenomus podisi North and South<br />
America<br />
Pentatomidae; not well adapted to N. viridula Buschman & Whitcomb<br />
1980; Correa & Moscardi<br />
1995; Orr et al. 1985a,<br />
1986<br />
Telenomus seychellensis East Africa Attacks other spp.; may be common on N. viridula Nixon 1935;<br />
Croix & Thindwa 1986<br />
Telenomus sp. Argentina<br />
Vietnam<br />
Minor attack on N. viridula eggs Liljestršm & Bernstein 1990<br />
van Lam 1996<br />
Trissolcus aloysiisabaudiae East Africa Reportedly common on N. viridula in cotton Fouts 1930; Chiaromonte<br />
1931; Paoli 1933<br />
Trissolcus basalis N. & S. America, S.<br />
Europe, Africa, Hawaii,<br />
Australia, New<br />
Zealand, Fiji<br />
Most important parasitoid <strong>of</strong> N. viridula outside central<br />
Africa and eastern Asia<br />
Miller 1928; Kamal 1937;<br />
Lever 1941; Buschman &<br />
Whitcomb 1980; Ferreira<br />
1980; Orr et al. 1986;<br />
Colazza & Bin 1995<br />
Trissolcus brochymenae N. and S. America Recorded ex N. viridula; attacks other pentatomids Johnson 1984b<br />
206 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong>
Table 4.12.1 (contÕd) Parasitoids <strong>of</strong> the green vegetable bug, Nezara viridula (modified after Jones 1988)<br />
Parasitoid<br />
HYMENOPTERA<br />
Geographic range Known host relations Selected references<br />
SCELIONIDAE (contÕd)<br />
Trissolcus crypticus Pakistan ex Plautia crossota and Acrosternum gramineum; bred Clarke 1993a<br />
readily in N. viridula in lab. in Australia and Hawaii<br />
Trissolcus euschisti California ex N. viridula and other pentatomids H<strong>of</strong>fmann et al. 1991<br />
Trissolcus hullensis North America,<br />
Venezuela<br />
Recorded ex N. viridula; attacks other pentatomids Johnson 1985<br />
Trissolcus lepelleyi Central Africa Descr. ex N. viridula, an apparently common host Nixon 1936; Le Pelley 1979<br />
Trissolcus lodosi Turkey Descr. ex N. viridula; nothing else known Szab— 1981<br />
Trissolcus maro Southern Africa N. viridula is only known host Nixon 1935; Croix &<br />
Thindwa 1986<br />
Trissolcus mitsukurii Japan Important parasitoid <strong>of</strong> N. viridula in Japan Kiritani & Hokyo 1962;<br />
Hokyo & Kiritani 1963<br />
Trissolcus oenone Australia Johnson 1991<br />
Trissolcus ogyges Australia one recent record Seymour & Sands 1993<br />
Trissolcus rudus Vietnam van Lam 1996<br />
Trissolcus scuticarinatus South America One record ex N. viridula; attacks other pentatomids Ferreira 1986<br />
Trissolcus sipius East Africa Descr. ex N. viridula but not reported since Nixon 1936<br />
Trissolcus solocis Florida, Mexico Recorded ex N. viridula; attacks other pentatomids Buschman & Whitcomb<br />
1980; Johnson 1985<br />
Trissolcus thyantae Eastern N. America Recorded ex N. viridula; attacks other pentatomids Johnson 1985<br />
Trissolcus urichi Brazil Ferreira & Moscardi 1995<br />
Trissolcus utahensis California ex N. viridula and other pentatomids H<strong>of</strong>fmann et al. 1991<br />
Trissolcus sp. Taiwan<br />
India<br />
Recorded ex N. viridula; host relations unknown<br />
Recorded ex N. viridula<br />
Taiwan Agricultural<br />
Research Institute 1984;<br />
Nath & Dutta 1994<br />
4.12 Nezara viridula 207
208 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
The role <strong>of</strong> pheromones and other chemical<br />
secretions<br />
Sexually mature male N. viridula release a pheromone that is a powerful<br />
attractant for mature females and also attracts males and older nymphs, but<br />
to a lesser extent. It is produced from the ventral abdominal epidermis<br />
(Lucchi 1994). The principal ingredient <strong>of</strong> the mixture <strong>of</strong> compounds<br />
(obtained from extracts <strong>of</strong> male cuticle), isolated from bugs collected in<br />
southern France, is the sesquiterpene (z)-a-bisabolene trans epoxide,<br />
whereas the accompanying cis isomer is not attractive (BrŽzot et al. 1993).<br />
The pheromone blend differs between some Brazilian (which do not produce<br />
the cis compound) (Baker et al. 1987) and North American, Hawaiian and<br />
Japanese bug populations which do, but in differing ratios. Other related<br />
pentatomids (e.g. the North American Acrosternum hilare) emit mixtures<br />
containing distinctive ratios (but containing more cis isomers) <strong>of</strong> the same<br />
sesquiterpenes as in N. viridula (Aldrich et al. 1989). Males <strong>of</strong> the native<br />
Japanese Nezara antennata and Acrosternum aseadum produce speciesspecific<br />
pheromone blends based on the same compounds as Nezara<br />
viridula. However, whereas the trans/cis 1,2 epoxide ratio <strong>of</strong> N. antennata is<br />
within the range for most USA. N. viridula populations (3 to 4.4:1), the<br />
blend from Japanese N. viridula males is 0.82:1 to 1:1. The ratio for Italy<br />
was 2.16:1, for Brazil 2.28 to 4.67:1, and for Australia 3.90:1. The ratios for<br />
Acrosternum hilare, A. marginatum and A. pennsylvanicum are 6:100, 7:100<br />
and 94:100 respectively (Aldrich et al. 1989, 1993). However, the situation<br />
is not clearcut. Brezot et al. (1994) studied the proportions <strong>of</strong> cis and trans<br />
bisabolene epoxides in individuals <strong>of</strong> a southern France (SF) and a French<br />
West Indies (FWI) strain <strong>of</strong> N. viridula. The trans isomer composed 42 to<br />
82% <strong>of</strong> bisabolene epoxides in SF males and 74 to 94% <strong>of</strong> FWI males.<br />
Means differed significantly in spite <strong>of</strong> this inter-individual variation. Ryan<br />
et al. (1995) also found variability in the ratio <strong>of</strong> isomers within a single<br />
N. viridula population in Australia.<br />
It is interesting that the pheromone mixture from mature males in<br />
southeastern United States is also highly attractive to the tachinid parasitoid<br />
Trichopoda pennipes (Aldrich et al. 1987), which lays far more eggs on male<br />
than on female N. viridula. Trichopoda spp. and a group <strong>of</strong> related tachinids<br />
are native to the Americas, where they attack a small group <strong>of</strong> native<br />
pentatomid bugs. It is postulated that the chemical similarity <strong>of</strong> the<br />
Acrosternum and Nezara pheromones facilitated the adoption by the<br />
tachinids <strong>of</strong> N. viridula when it reached the Americas (see below).<br />
Furthermore, that the immigrant populations <strong>of</strong> N. viridula released both the<br />
trans and cis isomers and that parasitisation by the tachinids preferentially
4.12 Nezara viridula 209<br />
attracted to the cis isomer provided the major selection pressure leading to<br />
the present N. viridula populations having predominantly the trans isomer in<br />
their pheromone mix. Perhaps in parallel, in the 200 years or so <strong>of</strong> interaction<br />
between N. viridula and T. pennipes in tropical America, N. viridula has<br />
evolved a shorter pre-oviposition period and a longer developmental period<br />
than an Italian population (Hokkanen and Pimentel 1984; Aldrich et al.<br />
1989). At all events, at least two distinct pheromone strains <strong>of</strong> N. viridula<br />
can now be distinguished, based on the presence or absence in the volatile<br />
secretions <strong>of</strong> the cuticle <strong>of</strong> cis-(z)-a-bisobolene epoxide. Mature N. viridula<br />
males <strong>of</strong> one population from Brazil produce the trans, but not the cis isomer<br />
(Baker et al. 1987), whereas males from 2 other populations do in ratios <strong>of</strong><br />
2.28:1 and 4.67:1 respectively (Aldrich et al. 1993), similar to males from<br />
southern USA which produce the trans and cis isomers in a 3:1 ratio (Aldrich<br />
et al. 1987) and those from Southern France in a 2:1 ratio (Baker et al. 1987).<br />
A quite different, but somewhat analogous situation, occurs with the<br />
hymenopteran Trissolcus basalis. A short chain unsaturated aldehyde, (E)-<br />
2-decenal present in the defensive scent produced in the adult N. viridula<br />
metathoracic gland (Gilby and Waterhouse 1965) is attractive to female<br />
T. basalis. A different compound, secreted on the eggs by the ovipositing<br />
female N. viridula, attracts female T. basalis to the egg raft (Mattiacci et al.<br />
1991, 1993). This compound is produced in the bug ovary and serves as an<br />
adhesive for attaching the eggs to the oviposition substrate. The adhesive<br />
and the material(s) responsible for the kairomone activity were partly<br />
soluble in water and completely in acetone and elicited recognition<br />
behaviour from T. basalis females when applied to glass beads. The<br />
adhesive appears to be a mucopolysaccharide-protein complex (Bin et al.<br />
1993).<br />
Attempts at biological control<br />
Early attempts at biological control were made chiefly in Australia and the<br />
Pacific, countries where the introduction <strong>of</strong> the bug was comparatively<br />
recent, but many other countries have been involved in more recent times.<br />
AUSTRALIA<br />
N. viridula was first reported in Australia in 1916 and soon became a<br />
widespread and serious pest. In 1933 the scelionid wasp Trissolcus basalis<br />
was introduced from Egypt into Western Australia (Table 4.12.2) where it<br />
readily became established and produced a great reduction in the pest status<br />
<strong>of</strong> the green vegetable bug. From Western Australia the parasitoid was<br />
distributed widely throughout south and southeastern Australia, making a<br />
considerable impact on the bug, except in cultivated areas in inland eastern<br />
Australia, where the cold winters affected its abundance (Wilson 1960).
210 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Additional strains were later liberated <strong>of</strong> what, at the time, was believed to be<br />
T. basalis. These originated from the West Indies (1953), Italy (1956) and<br />
Pakistan (1961). However, doubt has been cast on the specific identity <strong>of</strong> the<br />
Italian material, although T. basalis is known to occur there. The Pakistan<br />
material, originally identified as T. basalis (but probably containing two<br />
species) has recently been shown (Clarke 1993a) to have consisted mainly <strong>of</strong><br />
a new species Trissolcus crypticus. This bred readily in N. viridula eggs in<br />
the laboratory and, later, was widely distributed in Australia and Hawaii.<br />
However, T. crypticus has not been reported since in field collections.<br />
For many years up to the mid 1960s N. viridula was a very common and<br />
serious pest in the Canberra district, damaging tomatoes and beans in<br />
particular. This situation changed dramatically following the liberation <strong>of</strong><br />
the material from Pakistan (presumably both T. basalis and T. crypticus)<br />
and, since then, N. viridula has become a very uncommon insect, appearing<br />
only in extremely limited numbers late in the season and only every few<br />
years. A great improvement also resulted about the same time in other<br />
subcoastal eastern cultivated areas, credited by Ratcliffe (1965) also to the<br />
liberation <strong>of</strong> material from Pakistan.<br />
However, since T. crypticus apparently did not become established its<br />
role, if any, in the changed situation is unclear. It is not known whether any<br />
cross mating with T. basalis might have occurred, which might have<br />
resulted in greater adaptability <strong>of</strong> T. basalis to the Canberra environment.<br />
The continuing very low abundance <strong>of</strong> N. viridula is possibly attributable to<br />
a heavy attack on its eggs by a resident population <strong>of</strong> T. basalis which is<br />
maintained on native pentatomid hosts, <strong>of</strong> which there are several (see later).
Table 4.12.2 Introductions for the biological control <strong>of</strong> Nezara viridula<br />
Country and species<br />
ANTIGUA<br />
Liberated From Result Reference<br />
Anastatus sp. 1961Ð62 Pakistan Ð Cock 1985<br />
Trichopoda pilipes 1949<br />
1955<br />
Florida<br />
Montserrat<br />
Ð<br />
Ð<br />
Cock 1985<br />
Cock 1985<br />
Xenoencyrtus hemipterus (= X. niger) 1963 Australia Ð Cock 1985<br />
ARGENTINA<br />
Trissolcus basalis 1981 Australia, Hawaii + Crouzel & Saini 1983; Porta & Crouzel 1984<br />
AUSTRALIA<br />
Bogosia antinorii 1958 Kenya _ Greathead 1971<br />
Telenomus chloropus<br />
(= T. nakagawai)<br />
1962<br />
1980<br />
1981<br />
Japan<br />
Japan<br />
Japan<br />
Ð<br />
Ð<br />
?<br />
Callan 1963<br />
Field 1984<br />
Field 1984; J. Turner pers. comm. 1984<br />
Trissolcus basalis 1933 Egypt + Kamal 1937; Wilson 1960<br />
1953 West Indies + Wilson 1960<br />
1956 Italy + Wilson 1960<br />
1961 Pakistan + Ratcliffe 1965<br />
1979Ð82 USA + Field 1984<br />
1979Ð82 Brazil + J. Turner pers. comm. 1984<br />
1979Ð82 South Africa + J. Turner pers. comm. 1984<br />
Trissolcus crypticus 1961 Pakistan Ð Clarke 1993a<br />
Trissolcus mitsukurii 1962 Japan + Callan 1963<br />
Trichopoda giacomelli Argentina ? Liljesthršm 1994<br />
4.12 Nezara viridula 211
Table 4.12.2 (contÕd) Introductions for the biological control <strong>of</strong> Nezara viridula<br />
Country and species Liberated From Result Reference<br />
Trichopoda pennipes 1941Ð43<br />
1949Ð50<br />
1952Ð53<br />
1980<br />
1980<br />
?<br />
Trichopoda pilipes 1952Ð54<br />
1980<br />
1980<br />
Ooencyrtus submetallicus 1953Ð57<br />
1962<br />
BRAZIL<br />
Florida<br />
Florida<br />
Florida<br />
Florida<br />
Florida<br />
Italy<br />
West Indies<br />
West Indies<br />
Hawaii<br />
Trinidad<br />
Trinidad<br />
Ð<br />
Ð<br />
Ð<br />
Ð<br />
Ð<br />
+<br />
Ð<br />
Ð<br />
Ð<br />
Ð<br />
Ð<br />
Wilson 1960<br />
Wilson 1960<br />
Wilson 1960<br />
Michael 1981<br />
Michael 1981<br />
Giangiuliani et al. 1994; Colazza & Bin 1995<br />
Wilson 1960<br />
Michael 1981<br />
Michael 1981<br />
Wilson 1960<br />
CSIRO files<br />
Gryon japonicum Japan + Kishino & Teixeira 1994<br />
Gryon obesum USA + Correa & Moscardi 1995<br />
Ooencyrtus nezarae Japan Ð Kobayashi & Cosenza 1987<br />
Telenomus chloropus Japan Ð Kobayashi & Cosenza 1987<br />
Telenomus gifuensis Japan Ð Kobayashi & Cosenza 1987<br />
Trissolcus mitsukurii Japan + Kobayashi & Cosenza 1987;<br />
Kishino & Teixeira 1994,<br />
Trissolcus sp. Japan Ð Kobayashi & Cosenza 1987<br />
CALIFORNIA<br />
Trissolcus basalis 1987<br />
1987<br />
1987<br />
France<br />
Italy<br />
Spain<br />
+<br />
+<br />
+<br />
H<strong>of</strong>fman et al. 1991<br />
H<strong>of</strong>fman et al. 1991<br />
H<strong>of</strong>fman et al. 1991<br />
212 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong>
Table 4.12.2 (contÕd) Introductions for the biological control <strong>of</strong> Nezara viridula<br />
Country and species<br />
CHILE (EASTER ISLAND)<br />
Liberated From Result Reference<br />
Ectophasiopsis arcuata 1982, 1985/6 Chile + Ripa & Rojas 1989; Ripa et al. 1995<br />
Trissolcus basalis 1982 Chile Ð Ripa & Rojas 1989; Ripa et al. 1995<br />
COOK IS<br />
Trissolcus basalis<br />
FIJI<br />
1950 New Zealand ? Cumber 1953, Walker and Deitz 1979;<br />
A. Walker pers. comm. 1984<br />
Trissolcus basalis 1941 Australia + Lever 1941, 1943<br />
Trichopoda pennipes<br />
HAWAII<br />
1949 Florida ? OÕConnor 1950<br />
Trissolcus basalis 1962 Australia + Davis 1964, 1967<br />
Xenoencyrtus hemipterus<br />
(= X. niger)<br />
1962 Australia Ð Davis 1964, 1967<br />
Telenomus chloropus 1967 Japan Ð Davis & Chong 1968<br />
Telenomus sp. 1962 Australia Ð Davis 1964, 1967<br />
Trichopoda pilipes 1962 West Indies + Davis 1964, 1967<br />
Trichopoda pennipes 1962 Florida + Davis 1964, 1967<br />
Ooencyrtus submetallicus 1962 West Indies Ð Davis 1964, 1967<br />
Ooencyrtus trinidadensis 1962 West Indies Ð Davis 1964, 1967<br />
Trissolcus mitsukurii 1966 Japan Ð Davis & Krauss 1967<br />
4.12 Nezara viridula 213
Table 4.12.2 (contÕd) Introductions for the biological control <strong>of</strong> Nezara viridula<br />
Country and species<br />
ITALY<br />
Liberated From Result Reference<br />
Trichopoda pennipes 1984 or<br />
earlier<br />
1989<br />
Trissolcus basalis 1989 ? + Colazza & Bin 1995<br />
KIRIBATI<br />
?<br />
?<br />
+<br />
+<br />
G.K. Waite pers. comm.<br />
Gianguiliani & Farinelli 1995; Colazza et al. 1996a<br />
Trissolcus basalis<br />
MONTSERRAT<br />
1979 Fiji + Anon. 1979b; Williams 1979;<br />
Dhamaraju pers. comm. 1985<br />
Anastatus sp. 1961Ð62 Pakistan Ð Cock 1985<br />
Trissolcus mitsukurii<br />
NEW ZEALAND<br />
1966 Japan Ð Cock 1985<br />
Ooencyrtus submetallicus West Indies Ð Jones 1988<br />
Trichopoda pennipes 1965Ð67 Florida Ð Cumber 1967; Clausen 1978<br />
Trissolcus basalis 1949 Australia + Cumber 1951<br />
Xenoencyrtus hemipterus 1962 Australia Ð Jones 1988<br />
NEW CALEDONIA<br />
Trissolcus basalis 1942Ð43 Fiji + Lever 1943<br />
PAPUA NEW GUINEA<br />
Trissolcus basalis 1978 Australia +<br />
+<br />
Anon. 1983<br />
Young 1982<br />
Trichopoda pennipes 1977 Hawaii Ð J.W. Ismay pers. comm. 1985<br />
Trichopoda pilipes 1980Ð81 Hawaii Ð Young 1982<br />
214 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong>
Table 4.12.2 (contÕd) Introductions for the biological control <strong>of</strong> Nezara viridula<br />
Country and species<br />
PITCAIRN IS<br />
Liberated From Result Reference<br />
Trissolcus basalis 1952 Fiji ? Dumbleton 1957<br />
POHNPEI<br />
Trissolcus basalis 1989 Hawaii + Esguerra et al. 1993<br />
AMERICAN SAMOA<br />
Trissolcus basalis 1953 Fiji ? Dumbleton 1957<br />
SAMOA<br />
Trissolcus basalis 1953 ? + Clausen 1978<br />
SOLOMON IS<br />
Trissolcus basalis 1940 Australia + CSIRO files<br />
Trichopoda pennipes 1940, 1949,<br />
1950<br />
SOUTH AFRICA<br />
Florida _ Dumbleton 1957<br />
OÕConnor 1950<br />
Trissolcus basalis 1955 Australia + Bedford 1964; Greathead 1971; Annecke & Moran<br />
1982; Bennett 1990<br />
Trichopoda pennipes 1986<br />
1994<br />
ST KITTS AND NEVIS<br />
Florida<br />
USA, Italy<br />
?<br />
?<br />
Bennett 1990<br />
Farinelli et al. 1994<br />
Anastatus sp. 1961Ð62 Pakistan Ð Cock 1985<br />
Trissolcus mitsukurii 1966 Japan Ð Cock 1985<br />
ST VINCENT<br />
Anastatus sp. 1961Ð62 Pakistan Ð Cock 1985<br />
TAIWAN<br />
Trissolcus basalis 1983 ? + Su & Tseng 1984<br />
4.12 Nezara viridula 215
Table 4.12.2 (contÕd) Introductions for the biological control <strong>of</strong> Nezara viridula<br />
Country and species<br />
TONGA<br />
Liberated From Result Reference<br />
Trissolcus basalis<br />
USA (CALIFORNIA)<br />
1941 Australia + Dumbleton 1957<br />
Clausen 1978<br />
Trissolcus basalis<br />
ZIMBABWE<br />
1992 eastern USA + Pickett et al. 1996<br />
Trissolcus basalis 1955 Australia ? Annecke & Moran 1982<br />
216 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong>
4.12 Nezara viridula 217<br />
Doubt has been cast (Clarke 1990) both on the policy and effectiveness<br />
<strong>of</strong> introducing strains <strong>of</strong> T. basalis from different regions having differing<br />
environmental conditions. This practice has either led in Australia to<br />
effective biological control <strong>of</strong> N. viridula extending into additional<br />
environments or, alternatively, the initial genetic make-up <strong>of</strong> T. basalis has<br />
steadily undergone changes to allow progressive adaptation to new<br />
environments. It is possible that T. basalis consists <strong>of</strong> a complex <strong>of</strong> sibling<br />
species but, if not, it would be surprising if T. basalis has remained a<br />
homogenous species worldwide. Johnson (1985) found that American<br />
specimens <strong>of</strong> T. basalis showed much less morphological variation than<br />
those <strong>of</strong> Africa suggesting an African origin for the species. Furthermore,<br />
Handley (1975) reported that Australian T. basalis females would not mate<br />
with American males, although Powell and Shepherd (1982) found that<br />
reproductive isolation did not occur within any <strong>of</strong> 3 Australian strains<br />
examined; or between them and a strain from Florida. Nevertheless, the<br />
latter strain proved least fecund. Ferreira and Zamataro (1989) found no<br />
differences in reproductive capacity or longevity between an Australian and<br />
a Brazilian strain <strong>of</strong> T. basalis and Awan et al. (1989) concluded that Italian,<br />
French and Spanish populations consist <strong>of</strong> a single biotype, although several<br />
significant differences were observed in their biology. On the other hand,<br />
differences in courtship behaviour have been observed between different<br />
populations <strong>of</strong> T. basalis (Bin et al. 1988; Clarke and Walter 1992).<br />
However, it is not known whether any <strong>of</strong> these differences has any<br />
significance for biological control.<br />
Three other wasps have been introduced, Trissolcus mitsukurii (Japan<br />
1962), Telenomus chloropus (Japan 1962, Japan via USA 1980) and<br />
Ooencyrtus submetallicus (West Indies 1952Ð53). Only the former is<br />
believed to have become established but its impact has not been reported<br />
(Field 1984; J. Turner pers. comm. 1985).<br />
The only parasitoid reported from adult or nymphal N. viridula is the<br />
native tachinid Cylindromyia rufifemur (Cantrell 1984; Coombs and Khan<br />
1997). Three exotic species <strong>of</strong> parasitic tachinid fly have also been<br />
employed in attempts at biological control <strong>of</strong> Nezara viridula, Trichopoda<br />
pennipes from Florida, T. pilipes from the West Indies and Bogosia antinorii<br />
from Kenya. The Trichopoda species were introduced into Australia in the<br />
1940s and 1950s, but failed to become established (Wilson 1960). More<br />
recently the Trichopoda species were introduced into Western Australia<br />
from their native countries and also from Hawaii where they have been<br />
successfully established (Michael 1981). They have not become established<br />
in Australia. It is tempting to postulate that the pheromone blend secreted by<br />
Australian N. viridula does not attract Trichopoda pennipes and T. pilipes
218 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
females, whereas that <strong>of</strong> the Hawaiian population does. Whether or not this<br />
is true, it is clear that the responses by tachinid females to host pheromones<br />
introduces a considerable degree <strong>of</strong> host specificity to some particular<br />
populations <strong>of</strong> a host.<br />
The Argentinian Trichopoda giacomellii has been introduced to<br />
Australia for examination in quarantine for host specificity. Tests indicate<br />
that it has a limited host range involving only Nezara and its very close<br />
relations and it has now been approved for release in Australia (D.P.A.<br />
Sands pers. comm. 1997).<br />
At least three native wasps parasitise Nezara eggs, Telenomus sp.,<br />
Xenoencyrtus hemipterus and ÔCoruna sp.Õ (certainly a misidentification <strong>of</strong><br />
genus, Z. Boucek pers. comm. 1986), but these are <strong>of</strong> minor importance.<br />
The successful biological control <strong>of</strong> N. viridula in southern Australia has<br />
been repeated more recently in northwestern Australia where the green<br />
vegetable bug was first recorded in 1974. By 1976 populations were<br />
immense, for example over 33 nymphs and adults being recorded per square<br />
metre on a tomato crop and over 20 per head on badly affected sorghum.<br />
Since T. basalis was not present, it was introduced from southwestern<br />
Australia. Although more than 44 000 were released, initial establishment<br />
was poor. However, after 4 months, the situation improved dramatically,<br />
parasitisation was close to 100% and damage was reduced to sub-economic<br />
levels (Strickland 1979), although there are still periods <strong>of</strong> crop growth<br />
when the pest may be a problem (Michael 1981).<br />
T. basalis attacks, sometimes heavily, the egg masses <strong>of</strong> a range <strong>of</strong><br />
pentatomid bugs. In southern Australia common pentatomid hosts are the<br />
horehound bug Agonoscelis rutila (Noble 1937; Clarke and Walter 1994)<br />
Cermatulus nasalis and Oechalia schellembergii (Awan 1989). In<br />
northwestern Australia alternative pentatomid hosts include Piezodorus<br />
hybneri and Oechalia schellenbergii, the egg masses <strong>of</strong> which suffer<br />
respectively 68% and 51% parasitisation. The coreid bug Riptortus serripes<br />
is also attacked, 38% <strong>of</strong> its egg masses being parasitised (Strickland 1979).<br />
Nezara viridula is under excellent biological control and is generally a<br />
very uncommon insect throughout southern Australia. However regular or<br />
occasional damage occurs in a subcoastal zone extending from south east<br />
Queensland (Titmarsh 1979) to north central New South Wales (e.g.<br />
Forrester 1979) and into northern Victoria (Clarke 1992a) and especially on<br />
soybeans.<br />
In an attempt to control these damaging populations, nine strains <strong>of</strong><br />
T. basalis were introduced between 1979 and 1981, mass reared and<br />
released in southeastern Queensland. One strain came from each <strong>of</strong> South<br />
Carolina, Florida and Mississippi (USA), two from Brazil, two from South
4.12 Nezara viridula 219<br />
Africa and two from northern Australia (Darwin and Kununurra). Although<br />
a slightly higher level <strong>of</strong> parasitisation has resulted, the problem has not been<br />
resolved (J. Turner, pers. comm. 1984; Clarke 1992a). It is notable that<br />
N. viridula is seldom a pest except in regions where soybean is a major crop.<br />
Turner (1983) showed that the rate <strong>of</strong> movement <strong>of</strong> T. basalis on soybean<br />
Glycine max was a third <strong>of</strong> that on cowpea Vigna unguiculata, mungbean<br />
V. radiata radiata, bean Phaseolus vulgaris, or sunflower Helianthus<br />
annuus. The proportion <strong>of</strong> N. viridula eggs parasitised on soybean was down<br />
to 25% <strong>of</strong> that on cowpea, mungbean and sunflower. Observations, since<br />
contested by Kelly (1987), suggested that the arrangement and height <strong>of</strong> the<br />
soybean leaf hairs, which are neither evenly spaced nor patterned, were<br />
responsible for interfering with the waspsÕ searching activities and this<br />
suggestion needs further investigation. Many soybean varieties have beeen<br />
selected for cicadellid resistance, which is directly correlated with the<br />
density, length and orientation <strong>of</strong> the leaf hairs (Broersma et al. 1972).<br />
Sesame Sesamum indicum leaves were repellent to the wasps, and those that<br />
did alight left immediately and engaged in vigorous grooming elsewhere<br />
(Turner 1983). Thus the nature <strong>of</strong> crops in an area can materially affect the<br />
success <strong>of</strong> biological control <strong>of</strong> the green vegetable bug.<br />
In spite <strong>of</strong> the foregoing, a claim has been made (Clarke 1990, 1992a,b,<br />
1993a,b, Clarke and Walter 1992), that, in Australia, Ôthere is little evidence<br />
to support claims <strong>of</strong> successful biological control <strong>of</strong> N. viridulaÕ (Clarke<br />
1993b). It is a mystery how such a view can be maintained in the light <strong>of</strong> the<br />
abundant evidence, available to its authors, from Western Australia, South<br />
Australia and coastal and southern New South Wales (Wilson 1960; Callan<br />
1963; Ratcliffe 1965; Strickland 1979; Michael 1981; Field 1984;<br />
Waterhouse and Norris 1987). Furthermore, until the 1970s, when<br />
increasing plantings <strong>of</strong> soybean, in particular, have provided highly suitable<br />
conditions for N. viridula populations to increase greatly in southern<br />
Queensland and northern New South Wales, there were even publications by<br />
entomologists in the Queensland Department <strong>of</strong> Primary Industries that,<br />
with the exception <strong>of</strong> the Darling Downs, Ôcontrol <strong>of</strong> the pest has become<br />
virtually unnecessary resulting from the introduction and establishment <strong>of</strong> a<br />
tiny parasite É Õ (Passlow and Waite 1971) and Trissolcus basalis Ôhas<br />
reduced the importance <strong>of</strong> Nezara viridula (L.) in coastal QueenslandÕ<br />
(Smith 1977). The highly effective control progressively achieved over a<br />
vast area <strong>of</strong> southern and Western Australia is in no way diminished in<br />
validity by the fact that N. viridula is, indeed, an important pest in a much<br />
smaller area extending from southeast Queensland, through central NSW to<br />
northern Victoria (Clarke 1992a). Although it is most commonly associated<br />
there with soybean (see later under Italy) it is also found on a range <strong>of</strong> other<br />
crops, including grain legumes, tomatoes and beans.
220 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Clarke and Walter (1992) postulate that, because N. viridula oviposits<br />
only rarely during summer in southeast Queensland, T. basalis is largely<br />
without its preferred host for 60Ð90 days during which daily temperatures<br />
average more than 25¡C. This exceeds the average survival time <strong>of</strong> adult<br />
females at this temperature. Adult survival <strong>of</strong> T. basalis in summer is thus<br />
held to be the most likely factor limiting its populations. This postulate<br />
assumes that the egg masses <strong>of</strong> native pentatomid hosts <strong>of</strong> T. basalis are also<br />
in short supply over summer. It also appears not to apply to the far hotter but<br />
moister climate <strong>of</strong> the Ord Irrigation Area in far northern Western Australia,<br />
where N. viridula continues to be under generally excellent control.<br />
AFRICA<br />
N. viridula does not appear to be regarded as an important pest in northern<br />
Africa where its eggs are attacked by a number <strong>of</strong> Scelionidae, including<br />
Trissolcus aloysiisabaudiae, T. basalis, T. lepelleyi, T. maro, T. sipius and<br />
Telenomus seychellensis. Trissolcus basalis occurs mainly in coastal areas<br />
and the others are reported primarily from the eastern and central half <strong>of</strong> the<br />
continent. T. basalis was introduced to South Africa from Australia, New<br />
Zealand and USA, although there is some evidence that it may have already<br />
occurred there prior to these introductions (Giliomee 1958; Greathead<br />
1971). In Malawi eggs laid on macadamia were reported to experience an<br />
average <strong>of</strong> 74% parasitisation by Trissolcus maro and Telenomus<br />
seychellensis (Croix and Thindwa 1986). In Somalia Trissolcus<br />
alloysiisabaudiae is very abundant and may cause 100% parasitisation <strong>of</strong><br />
N. viridula eggs on cotton (Paoli 1933). T. lepelleyi and T. sipius attack<br />
N. viridula eggs in East Africa, the latter being known only from Kenya.<br />
Psix striaticeps, which occurs in tropical Africa and India, has been bred<br />
from N. viridula eggs (Jones 1988).<br />
ARGENTINA<br />
N. viridula was first recorded in 1919. Since the native tachinid parasitoid<br />
Trichopoda giacomellii was unable to maintain populations at sufficiently<br />
low levels, three parasitoid wasps were introduced, <strong>of</strong> which the most<br />
effective is Trissolcus basalis (Crouzel and Saini 1983; Porta and Crouzel<br />
1984).<br />
In Buenos Aires Province, mortality <strong>of</strong> N. viridula eggs was found to be<br />
due mainly to parasitisation by T. basalis, that <strong>of</strong> 1st to 3rd instar nymphs to<br />
predation and that <strong>of</strong> adults (together with reduction in egg production) to<br />
parasitisation by Trichopoda giacomellii. Adult mortality and reduction in<br />
egg production was found to be density dependent. Three egg parasitoids<br />
were present, Trissolcus basalis (95% <strong>of</strong> total parasitisation), Telenomus<br />
mormideae and Telenomus sp.. Nymphal mortality was principally due to<br />
spiders and predatory bugs (Podisus sp.), although there was also some
BRAZIL<br />
4.12 Nezara viridula 221<br />
parasitisation by Trichopoda giacomellii. Adverse climatic conditions<br />
(heavy rain) played a minor role in nymphal mortality (Liljesthršm and<br />
Bernstein 1990). Parasitisation <strong>of</strong> N. viridula eggs by T. basalis rose to a<br />
maximum <strong>of</strong> 90% in autumn although 33% <strong>of</strong> the parasitoids died, the<br />
majority (60%) in the pupal stage (Liljestršm and Camean 1992).<br />
In Rio Grande do Sul the main causes <strong>of</strong> mortality <strong>of</strong> N. viridula eggs laid<br />
throughout the season on soybean were infertility (2.7%: relatively<br />
constant), failure to hatch (14.1%: fluctuating), parasitisation (24%:<br />
relatively constant) and predation (17.3%: relatively constant) (Moreira and<br />
Becker 1986a). Three scelionid parasitoids were present, T. basalis,<br />
Trissolcus sp. and Telenomus mormideae. T. basalis killed a greater number<br />
<strong>of</strong> eggs and attacked a larger number <strong>of</strong> egg rafts than the other species<br />
(Moreira and Becker 1986b). A complex <strong>of</strong> polyphagous predators did not<br />
discriminate between parasitised and unparasitised eggs and were<br />
responsible for 25.5% mortality. The predators were responsible for 17.3%<br />
mortality <strong>of</strong> N. viridula and 34% <strong>of</strong> T. basalis (Moreira and Becker 1986c).<br />
Predation on host eggs was the main cause <strong>of</strong> mortality <strong>of</strong> T. basalis in the<br />
pre-emergence period (Moreira and Becker 1987).<br />
The tachinid fly Trichopoda giacomellii (= Eutrichopodopsis nitens) is<br />
the most important parasitoid <strong>of</strong> N. viridula in northern Paran‡ State. The<br />
level <strong>of</strong> parasitoid attack varies according to the plant on which the host is<br />
feeding and is highest when soybean is not available (Panizzi 1989).<br />
Although T. basalis (introduced) and Telenomus mormideae (native) were<br />
already present on the Cerrados area, Kobayashi and Cosenza (1987)<br />
introduced from Japan 5 additional species. In order <strong>of</strong> decreasing efficacy<br />
in parasitisation and adult emergence these were Trissolcus mitsukurii,<br />
Ooencyrtus nezarae, Telenomus chloropus, Telenomus gifuensis and<br />
Trissolcus sp.. Of the introduced species, T. mitsukurii parasitised eggs <strong>of</strong> all<br />
major pentatomid species throughout the year and also survived the dry<br />
winter season. In addition, it was the dominant competitor on egg masses.<br />
However, Bennett (1990) reports, more recently that it is not definite that<br />
permanent establishment has been achieved. When compared with<br />
T. basalis, the latter parasitised about 90% <strong>of</strong> exposed eggs with 60% adult<br />
emergence, whereas T. mitsukurii achieved about 70% parasitisation and<br />
40% emergence (Kobayashi and Cosenza (1987). In northern Brazil,<br />
Ferreira (1986) reported 40% parasitisation <strong>of</strong> N. viridula eggs by T. basalis<br />
and that Telenomus mormideae was also abundant.<br />
In the Federal District <strong>of</strong> Brazil the egg parasitoids Trissolcus mitsukurii<br />
and Gryon japonicum, introduced from Japan, gave good levels <strong>of</strong><br />
parasitisation (Kishino and Teixeira 1994). In Parana State T. basalis,
222 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Telenomus podisi and Gryon obesum parasitised up to 60% <strong>of</strong> Nezara eggs<br />
on soybean (Correa and Moscardi 1995).<br />
In southern Brazil, T. basalis is the main parasitoid, attacking between<br />
97.5% and 100% <strong>of</strong> N. viridula eggs laid on soybean. In 1988, 36.6% <strong>of</strong> the<br />
egg masses were attacked and 10.3% in 1989, with 21.8% and 6.3% <strong>of</strong> the<br />
individual eggs being parasitised respectively (Foerster and QueirÏz 1990).<br />
COOK IS<br />
T. basalis was introduced to Mangaia in 1950, but is not known to have<br />
become established (Cumber 1953; Walker and Deitz 1979) and this must be<br />
assumed not to have occurred.<br />
EASTER IS<br />
<strong>Control</strong> <strong>of</strong> N. viridula was achieved by the establishment <strong>of</strong> the tachinid<br />
Ectophasiopsis arcuata from mainland Chile, so that it is now difficult to<br />
find a bug. T. basalis was also introduced but was not recovered (Ripa and<br />
Rojas 1989; Ripa et al. 1992).<br />
FIJI<br />
N. viridula was first recorded in 1939 and Trissolcus basalis was introduced<br />
from Australia in 1941 (Lever 1941). Success was immediate and good<br />
control resulted (OÕConnor 1950). Large populations <strong>of</strong> N. viridula are<br />
reported to develop sometimes on cowpeas, but the insect is not troublesome<br />
on other legumes (Swaine 1971).<br />
Trichopoda pennipes was introduced from Florida in 1949, but its<br />
establishment is not recorded (OÕConnor 1950).<br />
HAWAII<br />
N. viridula was first recorded in 1961 and the wasps Trissolcus basalis,<br />
Xenoencyrtus hemipterus (= X. niger) and Telenomus sp. (all egg<br />
parasitoids) from Australia were released in 1962. Other importations in<br />
1962 were the tachinid fly, Trichopoda pilipes, which parasities last instar<br />
nymphs and adults, and two egg parasitic wasps Ooencyrtus submetallicus<br />
and O. trinidadensis from the West Indies. In 1963 Trichopoda pennipes<br />
was imported from Florida. Of these parasites, Trissolcus basalis,<br />
Trichopoda pennipes and T. pilipes became established (Davis and Krauss<br />
1964; Davis 1964, 1967; Croix and Thindwa 1967; Clausen 1978). Nezara<br />
populations declined steadily to sub-economic levels, with only sporadic<br />
outbreaks, and the species is generally under effective biological control<br />
(C.J. Davis, pers. comm. 1985). Average parasitisation by Trissolcus basalis<br />
ranged up to about 95% and by Trichopoda pilipes up to 86%. Trichopoda<br />
pupae are occasionally parasitised by the encyrtid Exoristobia philippensis<br />
(Davis 1964).<br />
More recently (1990Ð91) egg rafts <strong>of</strong> N. viridula placed in weeds at the<br />
border <strong>of</strong> macadamia nut plantations had significantly higher rates <strong>of</strong>
4.12 Nezara viridula 223<br />
parasitisation (49.9%) than rafts placed in the canopy <strong>of</strong> macadamia trees<br />
(14.7%). Predators were more effective at locating rafts placed in trees than<br />
in weeds and were always more efficient than T. basalis, regardless <strong>of</strong> their<br />
location. The egg parasitoid, Anastatus sp. was equally inefficient in both<br />
habitats. In 1990 only 1.2% <strong>of</strong> the eggs in the trees were parasitised and 8.6%<br />
in the weeds. During the same period, predators destroyed 26.0% and 14.5%<br />
in trees and weeds respectively.<br />
In 1991 parasitisation <strong>of</strong> eggs dropped to 0.2% and 1.7% in trees and<br />
weeds, whereas predation increased to 47.7% and 36.9% respectively.<br />
Doubt was, therefore, cast upon T. basalis having a prominent role in<br />
biological control <strong>of</strong> N. viridula in Hawaii (Jones 1995). Although this<br />
conclusion appears to follow in the macadamia agroecosystem studied, it<br />
would be <strong>of</strong> interest to know whether it applies also to other susceptible<br />
crops. Predation was attributed mainly to ants, including Pheidole<br />
megacephala.<br />
INDIA<br />
Singh (1973) reported no parasitoids in life-table studies <strong>of</strong> N. viridula on<br />
soybeans.<br />
INDONESIA<br />
Partial life tables showed on soybeans in Northern Sumatra that mortality <strong>of</strong><br />
N. viridula until the late 1st instar was 50 to 87%, <strong>of</strong> which 18 to 85%<br />
occurred during the egg stage and was caused mainly by predators. Only 2 to<br />
26% <strong>of</strong> the eggs were parasitised. The main predators were two species <strong>of</strong><br />
ants (Solenopsis geminata and Dolichoderus sp. a staphylinid beetle<br />
(Paederus sp.) and several crickets, although other egg predators belonging<br />
to the families Tettigoniidae, Lygaeidae and Anthocoridae were also<br />
observed feeding on the eggs. Trissolcus basalis parasitised the eggs but no<br />
evidence was obtained <strong>of</strong> the presence <strong>of</strong> tachinid parasitoids that attack late<br />
nymphs and adults (van den Berg et al. 1995).<br />
ITALY<br />
Before production <strong>of</strong> soybeans began in Italy in 1981 N. viridula was only<br />
important occasionally. Crops attacked included tomatoes and legumes. The<br />
increasing production (over a decade more than a thousand fold increase in<br />
area planted to soybeans) filled a temporal and food gap for N. viridula and<br />
other pentatomids (Colazza and Bin 1990). The second generation <strong>of</strong><br />
N. viridula now migrates each summer into soybeans at the beginning <strong>of</strong><br />
development <strong>of</strong> pods, which then provide the main food for reproduction<br />
and larval development. Abundance <strong>of</strong> N. viridula increased steadily in the<br />
eighties throughout northern and central Italy to a level at which it became a<br />
key pest. Three parasitoids were recorded from egg rafts, Anastatus<br />
bifasciatus, Ooencyrtus sp. and Trissolcus basalis. However the first 2
224 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
JAPAN<br />
species were never bred from egg rafts collected from soybeans.<br />
Approximately 20% <strong>of</strong> egg rafts were parasitised in 1986 and 1987,<br />
increasing to 50% in 1988 to 1992. Efficiency <strong>of</strong> parasitisation (% eggs<br />
parasitised divided by the number <strong>of</strong> egg masses discovered) was 65% in<br />
1986, but rose to 92% in 1988 and 1990.<br />
The early larval instars were generally free from parasitoid attack<br />
although two tachinids were occasionally recorded, the native Ectophasia<br />
crassipennis (5 to 10% parasitisation) and the accidentally introduced<br />
Trichopoda pennipes (2 to 15% parasitisation).<br />
T. basalis was also recovered from the eggs <strong>of</strong> 2 other pentatomid bugs<br />
present at low levels in soybean fields, Carpocoris mediterraneus and<br />
Piezodorus lituratus (Colazza and Bin 1995).<br />
The widespread Telenomus chloropus (= T. nakagawi) and also Trissolcus<br />
(= Asolcus) mitsukurii which is known only from Japan are the two most<br />
important parasitoids <strong>of</strong> N. viridula and have been studied intensively in<br />
Fukuoka. The host-finding ability <strong>of</strong> T. chloropus is superior to that <strong>of</strong><br />
T. mitsukurii and it parasitises egg masses more rapidly and more<br />
efficiently. Females lay about 100 eggs (which is 1.6 times that <strong>of</strong><br />
T. mitsukurii and live 11 days, or 3 days longer than T. mitsukurii (Nakasuji<br />
et al. 1966), although the latter may have at least 11 generations a year,<br />
whereas the former has some 9 generations (Hokyo et al. 1966b). Hokyo et<br />
al. (1966a) have shown experimentally that the two species do not<br />
discriminate between each otherÕs parasitised and unparasitised eggs, with<br />
T. mitsukurii larvae usually being successful in competition with<br />
T.chloropus larvae (Hokyo et al. 1966a). Furthermore, female T. mitsukurii<br />
<strong>of</strong>ten bite and kill female T. chloropus when they meet on the egg mass<br />
(Hokyo and Kiritani 1966). It follows that the effectiveness <strong>of</strong> T. chloropus<br />
is reduced by the presence <strong>of</strong> T. mitsukurii (Nakasuji et al. 1966), so it would<br />
be undesirable, in a biological control program, to introduce the latter along<br />
with the former. However, it should be borne in mind that T. mitsukurii is<br />
more abundant than T. chloropus in the southern coastal district <strong>of</strong> Fukuoka,<br />
whereas the reverse is true for the northern mountainous districts. The<br />
combined mortality caused by the two species amounted to 60 to 90% <strong>of</strong> the<br />
first spring generation N. viridula eggs.<br />
Three minor parasitoids have been bred from Nezara eggs in Fukuoka<br />
and several others are present elsewhere. The first, Telenomus gifuensis, is<br />
an effective parasitoid <strong>of</strong> Scotinophara lurida eggs, and also attacks a range<br />
<strong>of</strong> other pentatomids (Hidaka 1958; Hokyo et al. 1966b). The second,<br />
Ooencyrtus nezarae, the smallest <strong>of</strong> the three, is known from Nezara<br />
viridula, N. antennata and Anacanthocoris concoloratus. The third,
4.12 Nezara viridula 225<br />
Anastatus japonicus, is known as an egg parasitoid <strong>of</strong> the gypsy moth<br />
Lymantria dispar and other Lepidoptera (Hokyo et al. 1966b).<br />
KIRIBATI<br />
N. viridula was a pest on the islands <strong>of</strong> Betio and Tarawa in the 1970s.<br />
T. basalis was released in 1978 and, since 1984, this pest has not been<br />
recorded from Tarawa (E. Dharmaraju pers. comm. 1985).<br />
NEW CALEDONIA<br />
Trissolcus basalis was introduced in 1942Ð43 and became established<br />
(Lever 1943; Clausen 1978).<br />
NEW ZEALAND<br />
N. viridula was first recorded in 1944 and soon became a serious pest <strong>of</strong><br />
many crops. T. basalis was introduced from Australia in 1949 and rapidly<br />
became widely established. There followed a gradual decline in the severity<br />
<strong>of</strong> plant damage and, although populations continued to fluctuate seasonally,<br />
the situation became satisfactory (Cumber 1949, 1951, 1953, 1964). Over<br />
this period T. basalis extended its host range to other pentatomid bugs (e.g.<br />
Cuspicona simplex and Glaucias amyoti), thereby providing a source <strong>of</strong><br />
parasitoids to attack any eggs <strong>of</strong> Nezara that became available. Adaptation<br />
<strong>of</strong> T. basalis to Nezara under New Zealand conditions may also have been<br />
responsible for its improved performance (Cumber 1964).<br />
Trichopoda pennipes, originally from Florida, was obtained from<br />
Hawaii in 1965 and released over the next 3 summers. Evidence <strong>of</strong> a<br />
generation in the field was obtained in May 1967, but the fly did not become<br />
established. In addition to Nezara viridula, eggs were deposited on adults <strong>of</strong><br />
other pentatomids Antestia orbona, Cermatulus nasalis, Cuspicona simplex,<br />
Glaucias amyoti and Dictyotus caenosus, but a parasitoid was reared only<br />
from G. amyoti whose nymphs are readily parasitised (Cumber 1967).<br />
PAPUA NEW GUINEA<br />
N. viridula is a serious pest in the Markham Valley where T. basalis is<br />
present, but generally results in less than 30% parasitisation. In 1978 a strain<br />
<strong>of</strong> this parasite from Western Australia was released, but the level <strong>of</strong><br />
parasitisation did not increase (Young 1982). In Wau T. basalis and another<br />
scelionid egg parasite are generally effective, although N. viridula<br />
occasionally increases to pest proportions (Gagne 1979). The tachinid<br />
parasites Trichopoda pennipes and T. pilipes were introduced from Hawaii<br />
but failed to become established (J.W. Ismay pers. comm. 1985).<br />
T. giacomelli is being considered for release.<br />
PHILLIPINES<br />
Three decades ago it was reported that N. viridula was not a pest, apparently<br />
being controlled by a native egg parasitoid, Ooencyrtus sp. (Cendana, in<br />
Davis 1967) and it is interesting that Nezara was reported as present, but<br />
unimportant, in 1993 (Waterhouse 1993b). This may possibly be correlated
226 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
with the fact that soybean production in the Philippines is rather limited<br />
Ñmuch lower than in many other <strong>Southeast</strong> <strong>Asian</strong> countries, such as<br />
Thailand, Indonesia and Vietnam. Two egg parasitoids Telenomus comperei<br />
and T. pacificus have been reported from N. viridula eggs laid on<br />
groundnuts. Both species parasitised 100% <strong>of</strong> <strong>of</strong>fered eggs in 24 hours and<br />
adults emerged after 12 to 14 days. Adults lived up to 32 days when fed<br />
honey (Cadapan and Alba 1987).<br />
POHNPEI<br />
N. viridula became a major pest in the early 1990s on several islands in the<br />
Federated States <strong>of</strong> Micronesia. Following the introduction <strong>of</strong> T. basalis<br />
from Hawaii, the green vegetable bug population has become so low that it is<br />
rarely seen now on vegetables in Pohnpei (Esguerra et al. 1993; Suta and<br />
Esguerra 1993).<br />
SAMOA<br />
Trissolcus basalis was introduced in 1953 and became established (Clausen<br />
1978).<br />
SOLOMON IS<br />
Trissolcus basalis was introduced from Australia in 1940 against the<br />
coconut spathe bug Axiagastus campbelli. It is said to be established<br />
(CSIRO files) and a Trissolcus sp. has been recorded from pentatomid eggs<br />
(N. viridula or Plautia brunneipennis) on beans (R. Macfarlane pers. comm.<br />
1985).<br />
Trichopoda pennipes was introduced from Florida via Fiji in 1950 in<br />
order to control the coconut bug Amblypelta cocophaga and other<br />
phytophagous bugs (OÕConnor 1950), but it has not been collected since.<br />
SOUTH AFRICA<br />
Trichopoda pennipes was introduced from USA and Italy and liberated in<br />
1994 (Farinelli et al. 1994; van den Berg et al. 1994) but there is no<br />
information on establishment. T. giacomelli has also been imported for<br />
study (D.P.A. Sands pers. comm. 1997).<br />
TAIWAN<br />
T. basalis was introduced in 1983 and, two months after release,<br />
parasitisation rates <strong>of</strong> 90% and 60% respectively <strong>of</strong> N. viridula eggs at two<br />
sites was reported (Su and Tseng 1984).
THAILAND<br />
TONGA<br />
USA<br />
VANUATU<br />
4.12 Nezara viridula 227<br />
The most abundant egg parasitoid is Gryon fulviventris, which exists as a<br />
number <strong>of</strong> biotypes. Under a series <strong>of</strong> synonyms (Dissolcus fulviventris,<br />
Hadronotus fulviventris, H. antestiae and Gryon antestiae) it is known from<br />
Africa, India, Thailand, southern USSR and Malaysia. It parasitises the eggs<br />
<strong>of</strong> many species <strong>of</strong> Pentatomidae, Scutelleridae and Coreidae, but was<br />
reported by Jones (1988) for the first time in N. viridula eggs in Thailand,<br />
where it also breeds in the eggs <strong>of</strong> Piezodorus hybneri. In Africa larvae<br />
develop in Nezara viridula eggs, but adults do not emerge successfully.<br />
Other egg parasitoids are Telenomus chloropus, Ooencyrtus nezarae,<br />
Anastatus sp. (Jones et al. 1983b; Jones 1988), Telenomus sp. and Trissolcus<br />
basalis (Napompeth 1990).<br />
Trissolcus basalis was imported in 1941 and became established (Clausen<br />
1978).<br />
The influence <strong>of</strong> the host plant on which egg rafts <strong>of</strong> N. viridula are laid on<br />
the level <strong>of</strong> both parasitisation and predation was investigated in North<br />
Carolina by Shepard et al. (1994). Parasitisation was higher than predation<br />
on eggs on tomato and about equal in okra, soybean and cowpea. In one year,<br />
predation was higher than parasitisation in soybean towards the end <strong>of</strong> the<br />
growing season, but parasitisation was higher early in the season in okra and<br />
cowpea. Parasitisation <strong>of</strong> egg masses in wild radish reached a peak <strong>of</strong> nearly<br />
100% during spring and declined to about 30% in autumn. The major<br />
parasitoid from all crops was Trissolcus basalis, although Ooencyrtus<br />
submetallicus occured in low numbers. The conclusion was reached that<br />
both parasioids and predators may play an important role in regulating<br />
populations <strong>of</strong> N. viridula and that their combined action <strong>of</strong>ten resulted in<br />
the attack <strong>of</strong> 100% <strong>of</strong> egg masses in some crops.<br />
N. viridula occurs in Vila where its eggs are heavily parasitised by a wasp<br />
Trissolcus sp. (not T. basalis) (R. Weller pers. comm. 1986).
228 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Biology <strong>of</strong> the major species<br />
Scelionidae: Hymenoptera<br />
This is the most important family <strong>of</strong> hymenopterous parasitoids emerging<br />
from the eggs <strong>of</strong> N. viridula and is dealt with, amongst others, by Nixon<br />
(1935, 1936, 1937, 1966).<br />
Trissolcus basalis<br />
Kamal (1937) was the first <strong>of</strong> many to make a detailed study <strong>of</strong> the impact <strong>of</strong><br />
the egg parasite Trissolcus basalis on the abundance <strong>of</strong> Nezara viridula: no<br />
control measures are needed in Egypt. His work on the biology <strong>of</strong> the wasp<br />
has been supplemented by studies by many later workers (e.g. Wilson 1961;<br />
Cumber 1964; Powell and Shepherd 1982; Correa and Moscardi 1993, 1994;<br />
Awadalla 1996; Colazza et al. 1996b). The minute female <strong>of</strong> T. basalis<br />
oviposits in the side <strong>of</strong> the bug egg, after which she marks the egg by rubbing<br />
an abdominal secretion over it with the ovipositor, as a deterrent to other<br />
females from laying in the same egg. The length <strong>of</strong> the life cycle ranges from<br />
9 to 24 days depending on temperature, the entire egg, larval and pupal<br />
stages being passed inside the same eggshell. The adult wasps chew their<br />
way out through the lid <strong>of</strong> the eggshell, males usually emerging first and<br />
disputing with one another for possession <strong>of</strong> the egg batch and thereby the<br />
right to fertilise the later-emerging females. In hot weather the females live 4<br />
to 15 days and have considerable dispersive powers, as shown by the<br />
rapidity with which they spread through newly colonised areas. The adult<br />
wasps overwinter among leaves and litter.<br />
As indicated earlier, there is good evidence for the existence <strong>of</strong> several<br />
different strains <strong>of</strong> T. basalis. Experimental work using strains from three<br />
widely separated regions <strong>of</strong> Australia and from Florida showed that they<br />
were not reproductively isolated, that the Australian strains had a higher<br />
fecundity, but that adults <strong>of</strong> the Florida strain lived longer (Powell and<br />
Shepherd 1982).<br />
T. basalis is frequently recorded from several other pentatomids, but has<br />
a special preference for N. viridula (Jones 1988). It appears to be most<br />
effective in coastal and subcoastal areas and has been established in<br />
Argentina, Australia, Fiji, Hawaii, Kiribati, Papua New Guinea, New<br />
Caledonia, Samoa, Solomon Is, South Africa and Tonga and possibly in<br />
several other countries (Table 4.12.2).<br />
In Australia two species <strong>of</strong> the pteromalid Acroclissodes are parasitic on<br />
T. basalis (Clarke and Seymour 1992).
4.12 Nezara viridula 229<br />
Gryon sp.<br />
Some aspects <strong>of</strong> mating and reproduction <strong>of</strong> Gryon sp. in India have been<br />
investigated by Velayudhan and Senrayan (1989).<br />
Trissolcus mitsukurii<br />
This important egg parasitoid <strong>of</strong> Nezara in Japan also attacks the eggs <strong>of</strong><br />
several other pentatomids (Kishino and Teixeira 1994) , preferring species<br />
that deposit their eggs in small masses. It is bisexual and the first egg<br />
deposited by a mated female always produces a male. Both sexes have<br />
aggressive behaviour and females drive Telenomus chloropus females <strong>of</strong>f a<br />
pentatomid egg mass (Hokyo et al. 1966b). The fecundity and longevity <strong>of</strong><br />
T. mitsukurii were found to be less than those <strong>of</strong> T. basalis in laboratory<br />
trials at 26¡C and 65% RH. T. basalis parasitised 82.2% <strong>of</strong> eggs on the<br />
second day <strong>of</strong> adult life, whereas T. mitsukurii parasitised only 51.3%. On<br />
average, the former laid 250 eggs and the latter 80, and the longevity <strong>of</strong><br />
T. basalis was 80.1 days and <strong>of</strong> T. mitsukurii 42.6 days (Ferreira and<br />
Zamataro 1989).<br />
Too little is known about the dozen other Trissolcus species in Table<br />
4.12.1 to form an opinion <strong>of</strong> their value in biological control.<br />
Telenomus chloropus (= T. nakagawai)<br />
This is one <strong>of</strong> the most important egg parasitoids <strong>of</strong> N. viridula in Japan and<br />
also attacks eggs <strong>of</strong> N. antennata. Females are parthenogenic, lay about 100<br />
eggs and live for 11 days. They have a high searching ability and can<br />
parasitise all egss in a raft (Nakasuji et al. 1966). The species is oligophagous<br />
and prefers large egg rafts <strong>of</strong> pentatomids, such as those <strong>of</strong> N. viridula or<br />
N. antennata, to smaller egg masses <strong>of</strong> many other pentatomids. When<br />
introduced to the laboratory in Louisiana, USA. females lived for 8 days at<br />
24¡C, laid on average 60 eggs and developed from oviposition to emergence<br />
in 18 days. When reared from eggs <strong>of</strong> N. viridula which had been reared on<br />
resistant soybean its fecundity was half that <strong>of</strong> parasitoids reared on<br />
susceptible soybean and its mortality within Nezara eggs was higher (Orr et<br />
al. 1985b).<br />
Although morphological differences have not been found, it appears that<br />
T. chloropus exists as a series <strong>of</strong> biotypes. It is a widespread polyphagous<br />
parasitoid <strong>of</strong> pentatomid eggs throughout the Palaearctic Region, but it is<br />
recorded from Nezara only in Japan, Korea and Thailand. The Japanese<br />
biotype rarely produces males, although males occur elsewhere (Jones<br />
1988). T. chloropus from Japan has been released in Australia (Callan 1963;<br />
Field 1984), Brazil (Kobayashi and Cosenza 1987) Hawaii (Davis and<br />
Chong 1968) and the USA (Jones 1988), but has not become established,<br />
possibly due to its requirement for high humidity (85% RH or higher) for<br />
successful emergence (Orr et al. 1985a).
230 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Telenomus cyrus<br />
This parasitoid is known from Indonesia, the Philippines and Taiwan. In<br />
Taiwan it parasitises up to 19% <strong>of</strong> eggs in soybean and is the most important<br />
egg parasitoid <strong>of</strong> N. viridula in soybean, rice and jute (Taiwan Agricultural<br />
Research Institute 1984).<br />
Encyrtidae: Hymenoptera<br />
Species <strong>of</strong> Ooencyrtus have been recorded from N. viridula eggs from many<br />
parts <strong>of</strong> the world (Table 4.12.1), but are never a major component <strong>of</strong> the<br />
suite <strong>of</strong> parasitoids.<br />
Ooencyrtus submetallicus<br />
This species ranges from Florida through to West Indies, Brazil and<br />
Argentina (Jones 1988). It was found to be inferior to T. basalis in host<br />
location and dispersal in soybeans in Louisiana (Lee 1979). It was<br />
introduced to Australia, Hawaii and New Zealand, but did not become<br />
established (Wilson 1960; Davis and Krauss 1963; Davis 1967).<br />
Tachinidae: Diptera<br />
These are clearly important parasitoids <strong>of</strong> adult N. viridula in the Americas<br />
and the Ethiopian region. Outside <strong>of</strong> Japan, where one species attacking the<br />
green vegetable bug is known, there appear to be no records <strong>of</strong> tachinids<br />
regularly attacking N. viridula in Eastern Asia.<br />
Trichopoda spp.<br />
Trichopoda pennipes<br />
T. pennipes in North America is a complex <strong>of</strong> biotypes or sibling species. In<br />
the east its native hosts are the squash bug Anasa tristis, other coreids and<br />
several pentatomids (Arnaud 1978). In the southeast it occurs regularly in<br />
the native pentatomid Acrosternum hilare, but seldom in other pentatomids<br />
(Jones 1988). In California it does not oviposit on A. tristis, but in the field<br />
breeds in a pyrrhocorid and a largid bug (Sabrosky 1955; Dietrick and van<br />
den Bosch 1957). Salles (1991, 1993) has studied T. pennipes in Florida.<br />
The female Trichopoda pennipes lays eggs singly on the cuticle, mainly<br />
<strong>of</strong> the undersurface <strong>of</strong> fourth and fifth instar nymphs and adult bugs. The<br />
eggs hatch in 3 to 4 days and the young larvae bore directly into the host, tap<br />
the respiratory system <strong>of</strong> the bug for air and feed on the body fluids and<br />
internal organs <strong>of</strong> the host. When fully fed (16 days), the third instar larvae<br />
forces its way out through an intersegmental membrane <strong>of</strong> the host abdomen<br />
and pupates in the soil. After about 14 days the adult fly emerges. Up to 232<br />
eggs are laid by a female and, unlike Trissolcus basalis, there is <strong>of</strong>ten great<br />
wastage, many eggs (up to 237) being laid by several females on one adult<br />
bug, although only one parasitoid larva survives (Shahjahan 1968). The
4.12 Nezara viridula 231<br />
reproductive organs <strong>of</strong> the host bug may or may not be aborted by the<br />
feeding <strong>of</strong> the parasite, and it ultimately dies from mechanical injury caused<br />
by the emerging larva. The parasitoid overwinters as a second instar larva<br />
inside the hibernating adult bug (Beard 1940; Clausen 1978). Rearing<br />
methods are discussed by Gianguiliani and Farinelli (1995). Male<br />
N. viridula receive more parasitoid eggs than females and are more heavily<br />
parasitised (Mitchell and Mau 1971; Todd and Lewis 1976).<br />
In Georgia, USA, female Nezara parasitised by T. pennipes live about<br />
half as long as normal females and lay about a quarter the number <strong>of</strong> fertile<br />
eggs (Harris and Todd 1980).<br />
Trichopoda pilipes<br />
In the West Indies, instead <strong>of</strong> T. pennipes, the closely related T. pilipes<br />
(sometimes regarded as a subspecies) occurs. Both species have been<br />
established in Hawaii and T. pilipes is the more important (Davis 1967).<br />
Unsuccessful attempts have been made to establish one or both <strong>of</strong> these<br />
species in Australia, Fiji, South Africa and a number <strong>of</strong> other places,<br />
although T. pennipes has been established in Italy (Table 4.12.2).<br />
Trichopoda giacomellii<br />
Parasitisation levels by T. giacomellii <strong>of</strong> N. viridula in Argentina were<br />
45.3% on sorghum, 42.1% on flax, 29.9% on wheat and 27.9% on soybean.<br />
Levels on males were higher than on females, except on soybeans, where<br />
there was no significant difference (La Porta 1990). Liljesthršm (1985,<br />
1995) observed that the highest densities <strong>of</strong> parasitoids and the highest rate<br />
<strong>of</strong> parasitisation occurred in areas with the highest densities <strong>of</strong> N. viridula.<br />
Many fly eggs are laid on some individual hosts and few or none on<br />
others. More eggs are deposited on adult N. viridula than on 4th or 5th instar<br />
nymphs and more on adult males than on females. Some bugs are attacked<br />
sufficiently late in their development that they are able to produce at least<br />
one normal egg batch which has unaffected egg viability. In one study less<br />
than 7% parasitised nymphs died in the 5th instar. This resulted in sufficient<br />
eggs being laid to enable N. viridula to persist in the environment<br />
(Liljesthršm 1992, 1993a,b). In the laboratory at 26¡C, 70% RH and a 16hour<br />
day, the egg, larval and pupal stages lasted 2.8, 33.0 and 13.3 days<br />
respectively and females laid an average <strong>of</strong> 29 eggs (La Porta 1987).<br />
On hatching, young larvae penetrate the host cuticle and, on moulting to<br />
the 2nd instar, attach their posterior spiracles to one <strong>of</strong> the hostÕs tracheal<br />
trunks. The fully grown 3rd instar larva emerges from the host to pupate in<br />
the soil. Most hosts die shortly after the parasitoid larva has left.<br />
In one large field sample the maximum number <strong>of</strong> living parasitoid<br />
larvae found per host was 2 (4% <strong>of</strong> hosts), whereas only 1 living parasitoid
232 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
larva was found in 66% <strong>of</strong> hosts. Small, dead, damaged parasitoid larvae in<br />
some hosts provided evidence that there was competition for survival<br />
(Liljesthršm 1993b).<br />
Because moulting led to loss <strong>of</strong> unhatched eggs with the discarded<br />
cuticle, eggs laid on nymphs less frequently led to successful parasitisation<br />
than eggs laid on adults. With 1 parasitoid egg per adult, success was greater<br />
on males than on females whereas, when more than 4 eggs were present, the<br />
success rate was higher with females (Liljesthršm 1991). T. giacomellii<br />
parasitised 100% <strong>of</strong> N. viridula adults for 3 consecutive generations in an<br />
uncultivated area near Buenos Aires (Liljesthršm 1981) and it was<br />
concluded that T. giacomellii could regulate the population <strong>of</strong> N. viridula<br />
(Liljesthršm and Bernstein 1990).<br />
T. giacomellii (at times referred to incorrectly as Eutrichopodopsis<br />
nitens) is also an important parasitoid <strong>of</strong> N. viridula in Brazil. When<br />
parasitisation occurred in nymphs or newly moulted adults, adults did not<br />
reproduce and longevity was greatly reduced. Female N. viridula,<br />
parasitised on the 7th day <strong>of</strong> the adult stage, had their fecundity reduced by<br />
58%, but neither egg fertility nor size was affected (Ferreira et al. 1991).<br />
Parasitisation by T. giacomellii collected in the field in Brazil from soybean<br />
and other crops ranged from 27.1% to 52.7%. More parasitised eggs were<br />
found on males than on females and most eggs were on the thorax (Ferreira<br />
1984). High rates <strong>of</strong> parasitisation <strong>of</strong> N. viridula by T. giacomellii were<br />
observed on the weed Leonurus sibericus, but populations transferred to<br />
nearby soybean when this entered the reproductive phase. On the other hand,<br />
bugs living on castor, Ricinus communis, stayed on this plant all year round.<br />
This weed is <strong>of</strong> low nutritional value to them, but on it they are less liable to<br />
attack by the tachinid (Panizzi 1989).<br />
A recent laboratory study <strong>of</strong> the reproductive attributes <strong>of</strong> T. giacomelli<br />
determined the influence <strong>of</strong> adult food availability and body size on<br />
fecundity, and longevity, both relevant to any introduction program<br />
(Coombs 1997).<br />
Little is known about other Trichopoda species attacking pentatomid<br />
bugs: T. lanipes in Florida (Drake 1920), Trichopoda sp. in Uruguay (Guido<br />
and Ruffinelli 1956) and possibly other species in Brazil (Jones 1988).<br />
Bogosia antinorii<br />
This species is widespread in eastern and southern Africa and is known only<br />
from N. viridula (van Emden 1945; Barraclough 1985). There was<br />
apparently an unsuccessful attempt to establish it in Australia from material<br />
from Kenya (Greathead 1971).
4.12 Nezara viridula 233<br />
Ectophasiopsis arctuata<br />
This tachinid is common on N. viridula adults in Chile. Following its<br />
introduction to Easter Is it brought this bug under successful biological<br />
control (Ripa and Rojas 1989).<br />
Gymnosoma rotundata<br />
This tachinid parasitises N. viridula in Japan where up to 5% parasitisation is<br />
recorded (Kiritani et al. 1963). It is widespread in Palaearctic regions and<br />
attacks many hosts, including Nezara antennata, in Japan and Korea.<br />
Gymnosoma clavata has been recorded once from N. viridula in Europe<br />
(Herting 1960).<br />
One other tachinid has been reported once from N. viridula,<br />
Cylindromyia rufifemur from Australia (Cantrell 1984).<br />
Comments<br />
Although predators are undoubtedly important natural enemies <strong>of</strong><br />
N. viridula, particularly <strong>of</strong> its early stages, most are generalists which are<br />
unlikely to be approved nowadays by quarantine authorities for introduction<br />
as biological control agents. This situation is likely to be partly <strong>of</strong>fset by the<br />
fact that most countries possess a suite <strong>of</strong> generalist predators, some <strong>of</strong><br />
which are likely to attack N. viridula.<br />
The species <strong>of</strong> egg parasitoid most closely associated with N. viridula<br />
are concentrated in Africa and Japan. Elsewhere its eggs are parasitised by<br />
introduced species or by native species that have expanded their activities<br />
from native bugs. Indeed, it seems likely that the complex <strong>of</strong> Japanese<br />
parasitoids has probably expanded its host range from the oriental stink bug<br />
N. antennata <strong>of</strong> Japanese origin, just as the complex <strong>of</strong> Central and South<br />
American tachinids has clearly expanded to N. viridula adults from adults <strong>of</strong><br />
native bugs.<br />
Successful biological control <strong>of</strong> N. viridula has been achieved in many<br />
countries to which it has spread this century. These include a vast area (but<br />
not all) <strong>of</strong> Australia, also New Zealand, Hawaii and several other Pacific<br />
islands. The prospects are good for reducing its pest status in many other<br />
areas where effective parasitoids are not yet present. This might involve the<br />
introduction <strong>of</strong> additional species or strains <strong>of</strong> Trissolcus. In addition several<br />
<strong>of</strong> the many other parasitoids known to attack eggs (species in the genera<br />
Telenomus and Gryon) are well worth considering. In the Ethiopian region a<br />
tachinid (Bogosia antinorii) is an important parasitoid <strong>of</strong> adults and large<br />
nymphs and in the Americas there are at least 4 tachinid species worthy <strong>of</strong><br />
consideration: Trichopoda pennipes: USA; T. pilipes: West Indies;
234 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
T. giacomelli Argentina; and Ectophasiopsis arcuata: Chile. These 4 are<br />
reported to be more abundant now on N. viridula than on the native bugs<br />
they parasitised before the arrival <strong>of</strong> N. viridula (Jones 1988). A problem in<br />
their effective use is that location <strong>of</strong> N. viridula hosts is dependent, in some<br />
species at least, upon the secretion by N. viridula <strong>of</strong> a specific attractive<br />
blend <strong>of</strong> chemicals. Some biotypes <strong>of</strong> N. viridula that do not produce the<br />
appropriate blend largely escape oviposition. There are no records <strong>of</strong><br />
tachinids regularly attacking N. viridula in the East <strong>Asian</strong> Region, except for<br />
the widespread, polyphagous Gymnosoma rotundata, which was found to<br />
cause up to 5% parasitisation <strong>of</strong> N. viridula in Japan and also to attack<br />
N. antennata in Japan and Korea (Kiritani et al. 1963).<br />
The main areas where N. viridula continues to be an economically<br />
important pest in spite <strong>of</strong> attempts to use natural enemies including<br />
T. basalis appear, with the exception <strong>of</strong> certain crops such as macadamia and<br />
pecan nuts, to be associated with extensive plantings <strong>of</strong> soybeans. A detailed<br />
re-examination is required <strong>of</strong> the behaviour <strong>of</strong> T. basalis (and perhaps other<br />
egg parasitoids) in relation to ability to parasitise N. viridula egg masses laid<br />
on soybean. If it is demonstrated that certain physical or chemical<br />
characteristics <strong>of</strong> soybeans are responsible for poorer than usual<br />
performance, serious consideration should be given to the selection <strong>of</strong><br />
varieties that have minimal adverse effects on the parasitoids. This, <strong>of</strong><br />
course, is different from selecting soybean cultivars that are resistant to<br />
N. viridula, some <strong>of</strong> which are known (e.g. Kester et al. 1984; Bowers 1990).<br />
In this context it is relevant that the biology <strong>of</strong> Telenomus chloropus, an<br />
egg parasite introduced into southern USA in 1982 from Japan, was studied<br />
on eggs <strong>of</strong> N. viridula that had been reared on the stink bug-resistant<br />
soybean, PI 717444, or on the susceptible cultivar, Davis. Time <strong>of</strong><br />
development <strong>of</strong> the parasite did not differ significantly in eggs from either<br />
source, but success <strong>of</strong> emergence was lower from eggs laid on resistant<br />
soybean and fecundity <strong>of</strong> those that did emerge was about half <strong>of</strong> that <strong>of</strong><br />
individuals reared from eggs laid on Davis. The authors (Orr et al. 1985b)<br />
point out that, with a marked reduction in emergence and fecundity,<br />
combined with decreased host availability, there is the potential for<br />
reduction or elimination <strong>of</strong> resident parasite populations in fields <strong>of</strong> resistant<br />
soybeans.<br />
Comparatively little is known <strong>of</strong> the range <strong>of</strong> Nezara parasitoids in its<br />
centre <strong>of</strong> origin, namely the Ethiopian region, although at least 6 Scelionidae<br />
including T. basalis have been recorded, together with the apparentlyspecific,<br />
widespread tachinid Bogosia antinorii, whose effectiveness<br />
deserves study. It is probable that a thorough investigation in the Ethiopian<br />
region would disclose an additional range <strong>of</strong> potentially valuable species.
4.13 Ophiomyia phaseoli<br />
India<br />
Myanmar<br />
+<br />
20°<br />
Laos<br />
++<br />
0°<br />
20°<br />
China<br />
P<br />
Thailand<br />
+<br />
Cambodia<br />
Vietnam<br />
++<br />
P<br />
++ Brunei<br />
Malaysia<br />
+<br />
Singapore<br />
+++<br />
Indonesia<br />
Taiwan<br />
P<br />
++<br />
Philippines<br />
Australia<br />
Papua<br />
New Guinea<br />
++<br />
235<br />
It appears that the bean fly Ophiomyia phaseoli originated in Asia. Its most effective<br />
natural enemy, the braconid Opius phaseoli,<br />
is known from eastern Africa, India and the<br />
Philippines and has been introduced to Hawaii and Taiwan. This species is capable <strong>of</strong><br />
parasitisation levels <strong>of</strong> up to 90% or more and, when introduced to Hawaii, it and the related<br />
O. importatus resulted in successful biological control <strong>of</strong> bean fly.<br />
There are good reasons for countries where bean fly is a problem and where<br />
parasitisation levels are low, to consider introducing these and other parasitoids to assist in<br />
reducing bean fly populations.<br />
20°<br />
0°<br />
20°
236 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Ophiomyia phaseoli (Tryon)<br />
Rating<br />
Origin<br />
Distribution<br />
Diptera: Agromyzidae (this species was earlier included in the<br />
genus Agromyza or Melanagromyza)<br />
bean fly<br />
<strong>Southeast</strong> Asia China Southern and Western Pacific<br />
+++ Indo +++ Guam<br />
14 ++ Laos, Viet, Msia,<br />
Phil<br />
9 ++ Fiji, PNG<br />
+ Myan, Thai, Sing + Sam, Sol Is<br />
P Brun P P FSM<br />
Unknown, but presumably in association with one <strong>of</strong> its current legume host<br />
genera in India or possibly <strong>Southeast</strong> Asia. It was described by Tryon (1895)<br />
from specimens causing damage to beans in 1888 in Queensland, Australia<br />
and, shortly after, reported to cause similar damage in New South Wales<br />
(Froggatt 1899).<br />
This was given (CIE 1974a) as Africa:<br />
Burundi, Congo (Zaire), Egypt,<br />
Ethiopia, Kenya, Madagascar, Mauritius, Malawi, Mali, Nigeria, RŽunion,<br />
Rwanda, Senegal, South Africa, Sudan, Tanzania, Uganda, Zambia,<br />
Zimbabwe; Asia:<br />
Bangladesh, China, Hong Kong, India, Indonesia, Iraq,<br />
Israel, Jordan, Malaysia, Myanmar, Nepal, Pakistan, Philippines, Ryukyu Is,<br />
Singapore, Sri Lanka, Taiwan, Thailand; Australia and Pacific Islands:<br />
Australia, Caroline Is, Fiji, Hawaii, Irian Jaya, Mariana Is, Papua New<br />
Guinea and Samoa. To the above must be added Israel (Spencer 1990)<br />
Brunei, Laos, Singapore (Waterhouse 1993b) and Japan (Makino et al.<br />
1990). It has not been recorded from Europe or the Americas.<br />
A morphologically very similar species, O. spencerella,<br />
only readily<br />
distinguishable from O. phaseoli by the male genitalia, occurs in association<br />
with it in Kenya, Uganda, Tanzania and Nigeria on Phaseolus vulgaris and,<br />
less commonly, on several other legumes. There are 3 economically<br />
important agromyzid miners other than Ophiomyia phaseoli that attack<br />
much the same legumes in Asia. Melanagromyza (= Agromyza)<br />
obtusa is<br />
widely distributed in India as a pest <strong>of</strong> the developing seeds <strong>of</strong> chick and<br />
pigeon peas. The stem miners M. sojae and M. dolichostigma are pests <strong>of</strong><br />
soybean in Indonesia, Japan and Taiwan and damage French beans and<br />
cowpeas in Sri Lanka and East Africa (Singh and van Emden 1979).
Biology<br />
Host plants<br />
4.13<br />
Ophiomyia phaseoli<br />
237<br />
The adult O. phaseoli is a small fly (females 2.2 mm and males 1.9 mm in<br />
length), shiny black in colour except for legs, antennae and wing veins,<br />
which are light brown (Abul-Nasser and Assem 1966). Females generally<br />
oviposit in bright sunlight in the upper surface <strong>of</strong> the cotyledons (soybeans)<br />
or young leaves <strong>of</strong> its many hosts, laying from 100 to 300 eggs during a<br />
2Ðweek period (Otanes y Quesales 1918). Not all ovipositor punctures<br />
receive an egg, many provide sap which the females ingest (Goot 1930). On<br />
hatching from the egg after 2 to 4 days, the young larva forms a short leaf<br />
mine before tunneling into the nearest vein. Next the petiole is mined and the<br />
larva then moves down the stem (Taylor 1958). In young plants the main<br />
feeding takes place in the lower layers <strong>of</strong> the stem and the tap root may be<br />
penetrated. When larvae are numerous, some feed more deeply inside the<br />
stem and higher up in the plant.<br />
The larval and pupal stages occupy 7 to 10 days and 9 to 10 days<br />
respectively, resulting in a life cycle <strong>of</strong> about 3 weeks (Taylor 1958; Ooi<br />
1988). However, the life cycle may be as short as 17 days in the field in<br />
Malaysia (Khoo et al. 1991) and in the laboratory in India at 24¡ to 31¡C as<br />
short as 11 days (Singh et al. 1991). At the other end <strong>of</strong> the scale, at higher<br />
altitudes in Java, the larval stage can be extended from 17 to 22 days and the<br />
pupal stage from 13 to 20 days (Goot 1930). Pupation occurs head upwards<br />
beneath the epidermis and generally near the base <strong>of</strong> the stem (Greathead<br />
1969). In older plants, larvae may pupate at the base <strong>of</strong> the petioles.<br />
Talekar and Lee (1989) have developed a method for mass rearing bean<br />
fly on newly-germinated soybean cotyledons, permitting one person to<br />
produce 2 000 adults per day.<br />
Bean fly is known to attack at least 40 plant species. Most <strong>of</strong> its important<br />
hosts belong to the legume tribe Phaseoleae and particularly to the genus<br />
Phaseolus.<br />
The very susceptible P. vulgaris (French, kidney, haricot, runner<br />
or snap bean) is <strong>of</strong> Central American origin as are several other economic<br />
species <strong>of</strong> Phaseolus.<br />
However, from the point <strong>of</strong> view <strong>of</strong> the possible origin<br />
<strong>of</strong> O. phaseoli in Asia, all <strong>of</strong> the Asiatic species formerly placed in the genus<br />
Phaseolus have now been placed in the genus Vigna (Verdcourt 1970) and it<br />
is relevant that a number <strong>of</strong> important Vigna species are believed to have<br />
originated in India or nearby (Purseglove 1968). These include<br />
V. aconitifolia (moth bean), V. aurea (green or golden gram, mung bean)<br />
V. calcarata (rice bean) and V. mungo (black gram, urd bean). Other<br />
important hosts include Cajanus cajan (pigeon pea: origin Africa); Glycine
238 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Damage<br />
max (soybean: southern China); Lablab niger (= Dolichos lablab)<br />
(hyacinth<br />
bean: India); Pisum sativum (pea: southwestern Asia); and Vigna<br />
unguiculata (cowpea: Africa) (Purseglove 1968; Spencer 1973). Wild hosts<br />
include Canavalia ensiformis,<br />
Crotalaria juncea,<br />
C. laburnifolia,<br />
C. mucronata,<br />
Macroptilium atropurpureum,<br />
M. lathryoides,<br />
Phaseolus<br />
panduratus,<br />
P. semierectus and Vigna radiata (Goot 1930; Kleinschmidt<br />
1970; Spencer 1973; Abate 1991).<br />
There is a large variation in susceptibility between different cultivars <strong>of</strong><br />
susceptible species. This variation affords an important opportunity to select<br />
cultivars that suffer comparatively little damage and is being extensively<br />
investigated (e.g. Annappan et al. 1984, Gill and Singh 1988; AVRDC 1990,<br />
Talekar and Hu 1993; Talekar and Tengkano 1993; Gupta et al. 1995). Both<br />
morphological and chemical characteristics are involved (Chiang and Norris<br />
1983).<br />
O. phaseoli can be a limiting factor in the cultivation <strong>of</strong> susceptible legumes<br />
in <strong>Southeast</strong> Asia and most other regions where it occurs. Spencer (1973)<br />
and many other authors consider it to be one <strong>of</strong> the most serious <strong>of</strong> all<br />
agromyzid pests. Losses <strong>of</strong> 50% to 100% <strong>of</strong> crops are reported from many<br />
parts <strong>of</strong> the world, and are particularly heavy under dry conditions.<br />
Although some leaves may wilt as a result <strong>of</strong> mining and petiole<br />
tunnelling, most damage results from destruction <strong>of</strong> tissue at the junction <strong>of</strong><br />
the stem and root. When damage is limited, plants may survive by forming<br />
adventitious roots (e.g. with P. vulgaris and soybeans) and produce a limited<br />
crop. However, seedlings most frequently die. Plants that do not respond<br />
rapidly to root damage and develop adventitious roots are liable to break <strong>of</strong>f<br />
at ground level during windy periods. When infestations are heavy, the<br />
aggregation <strong>of</strong> puparia within the stem results in it swelling, splitting open<br />
and rotting. De Meijere (1922) reported that young plants in Sumatra<br />
generally died when they contained 10 to 20 larvae. In Egypt, 25 larvae and<br />
pupae have been found in a single bean plant (Hassan 1947) and as many as<br />
320 in a cowpea plant (Abul-Nasr and Assem 1966).<br />
Seed treatments or post emergence sprays with broad spectrum<br />
insecticides have been used to control O. phaseoli but, inter alia,<br />
they<br />
undoubtedly have serious adverse effects on its parasitoids. Useful control<br />
has been obtained by intercropping, the use <strong>of</strong> resistant varieties, adjusting<br />
planting dates, crop rotation and other cultural methods such as planting into<br />
rice stubble or covering the newly sown areas with rice straw. It is clear,<br />
however, that, in areas where effective parasitoids are already present, these<br />
can play an important role in minimising bean fly damage if not interfered<br />
with by insecticides.
Natural enemies<br />
4.13<br />
Ophiomyia phaseoli<br />
More than 50 parasitic Hymenoptera have been reported from bean fly<br />
(Table 4.13.1), almost all emerging from pupae, arising from eggs laid in<br />
host larvae. No egg parasitoids are known and no dipterous parasitoids have<br />
been recorded. Although figures for percent parasitisation are unavailable<br />
from many countries where bean fly occurs, the levels recorded are<br />
generally unimpressive, <strong>of</strong>ten less than 30%. The outstanding exception is<br />
the 90% or more, <strong>of</strong>ten produced by the braconid Opius phaseoli in East<br />
Africa and Ethiopia (Greathead 1969; Abate 1991) and similar levels from<br />
Opius phaseoli and Eurytoma sp. in the Agra region <strong>of</strong> India (Singh 1982).<br />
AUSTRALIA<br />
Although Tryon (1895), who described Ophiomyia phaseoli from<br />
Queensland specimens, believed it to be native to Australia, it has never been<br />
recorded extensively from indigenous plants (Kleinschmidt 1970), so is<br />
unlikely to have evolved there. Twelve parasitoids (two <strong>of</strong> them possibly<br />
hyperparasitoids) (Table 4.13.1) emerged from O. phaseoli pupae taken<br />
mainly from cowpea ( Vigna unguiculata).<br />
A later re-examination <strong>of</strong><br />
Australasian Chalcidoidea by Bou‹ek (1988) indicates that only 11 species<br />
were actually involved and none were hyperparasitoids. Bou‹ek (1988) lists<br />
one additional parasitoid, the eurytomid Plutarchia bicarinativentris, which<br />
also occurs in Papua New Guinea. The braconid Opius oleracei, which<br />
attacks bean fly, has also been recorded from the widespread agromyzid<br />
Chromatomyia horticola (= Phytomyza atricornis).<br />
EAST AFRICA<br />
Ophiomyia phaseoli was not reported in Uganda until the 1920s, Tanzania<br />
until 1937 and Kenya until 1939. It occurs in close association with<br />
Ophiomyia spencerella and also with O. centrosematis,<br />
a species which is<br />
known also from Indonesia.<br />
The bean fly is very heavily attacked in all climatic zones by the braconid<br />
Opius phaseoli (usually <strong>of</strong> the order <strong>of</strong> 70% to 90% and even up to 94.4%<br />
parasitisation), and relatively lightly (3% to 9%) by Opius importatus,<br />
by a<br />
polyphagous pteromalid near Herbertia sp. (less than 1%) and by an<br />
assemblage <strong>of</strong> chalcidoids. Puparia are also consumed by ants. Three wasps<br />
found in small numbers were considered to be hyperparasitoids, namely<br />
Norbanus sp. (Pteromalidae), Eupelmus sp. nr australiensis (= E. sp. nr<br />
popa)<br />
(Eupelmidae) and Pediobius sp. (Eulophidae). In spite <strong>of</strong> the heavy<br />
parasitisation by Opius phaseoli,<br />
sufficient bean flies survive in some<br />
seasons to cause heavy infestations <strong>of</strong> plantings. These parasitoids are thus,<br />
at times, unable to prevent economic damage (Greathead 1969).<br />
239
Table 4.13.1<br />
Natural enemies <strong>of</strong> Ophiomyia phaseoli<br />
Species<br />
HYMENOPTERA<br />
BRACONIDAE<br />
Country Reference<br />
Fopius sp. Thailand Burikam 1978; Burikam & Napompeth 1979,<br />
Napompeth 1994<br />
Opius importatus<br />
East Africa<br />
Hawaii<br />
Opius ?liogaster* Mauritius<br />
Zimbabwe<br />
Greathead 1969, 1975<br />
Raros 1975<br />
Moutia 1932<br />
Jack 1913; Taylor 1958<br />
Opius oleracei Australia Kleinschmidt 1970<br />
Opius phaseoli Botswana<br />
East Africa<br />
Ethiopia<br />
Hawaii<br />
India<br />
Madagascar<br />
Malawi<br />
Mauritius<br />
Philippines<br />
Taiwan<br />
Zimbabwe<br />
Opius sp.<br />
CHALCIDIDAE<br />
Taiwan Chu & Chou 1965<br />
1. East Africa Greathead 1969<br />
2. Sri Lanka Rutherford 1914b<br />
3.<br />
* probably Opius phaseoli (Fischer 1971a)<br />
Zimbabwe Jack 1913<br />
Greathead 1969<br />
Greathead 1969<br />
Abate 1991; Greathead 1969<br />
Davis 1971, 1972; Fischer 1971a; Greathead 1975;<br />
Raros 1975<br />
Fischer 1963; Ipe 1987<br />
Greathead 1969<br />
Letourneau 1994<br />
Greathead 1969<br />
Fischer 1971b<br />
N.S. Talekar pers. comm. 1994<br />
Greathead 1969<br />
240 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong>
Table 4.13.1 (contÕd) Natural enemies <strong>of</strong> Ophiomyia phaseoli<br />
Species<br />
HYMENOPTERA<br />
Country Reference<br />
CYNIPIDAE<br />
Cynipoide sp. Indonesia Goot 1930<br />
Eucoilidea sp. Taiwan Chu & Chou 1965<br />
Unidentified Thailand Burikam 1978<br />
EULOPHIDAE<br />
Aprostocetus sp. Ethiopia Abate 1991<br />
Chrysonotomyia douglasi Australia Bou‹ek 1988; Kleinschmidt 1970<br />
Chrysonotomyia sp. nr erythraea Ethiopia Abate 1991<br />
Chrysonotomyia formosa Ethiopia Abate 1991<br />
Chrysonotomyia sp. Australia Dodd 1917<br />
Cirrospilus sp. Ethiopia Abate 1991<br />
Euderus sp. Thailand Burikam 1978; Napompeth 1994<br />
Hemiptarsenus varicornis Australia Dodd 1917; Kleinschmidt 1970; Bou‹ek 1988<br />
Hemiptarsenus sp. Australia<br />
Philippines<br />
Kleinschmidt 1970<br />
Litsinger 1987<br />
Meruana liriomyzae Ethiopia Abate 1991<br />
Pediobius acantha Ethiopia Abate 1991<br />
Pediobius sp. East Africa Greathead 1969<br />
Tetrastichus sp. India Gangrade 1974; Ipe 1987<br />
Unidentified Hawaii Raros 1975<br />
4.13 Ophiomyia phaseoli 241
Table 4.13.1 (contÕd) Natural enemies <strong>of</strong> Ophiomyia phaseoli<br />
Species<br />
HYMENOPTERA<br />
Country Reference<br />
EUPELMIDAE<br />
Eupelmus ?australiensis Ethiopia, East Africa Greathead 1969; Abate 1991<br />
Eupelmus grayi Australia Kleinschmidt 1990<br />
Eupelmus sp. nr urozonus Egypt<br />
Ethiopia<br />
Eupelmus sp. Australia<br />
Ethiopia<br />
EURYTOMIDAE<br />
Eurytoma larvicola Australia<br />
Egypt<br />
Eurytoma poloni Indonesia<br />
Malaysia<br />
Philippines<br />
Eurytoma spp. 2 ´<br />
1 ´<br />
1 ´<br />
1 ´<br />
1 ´<br />
1 ´<br />
Australia<br />
Egypt<br />
Ethiopia<br />
India<br />
Indonesia<br />
Taiwan<br />
Plutarchia sp. Malaysia<br />
Philippines<br />
Abul-Nasr & Assem 1968<br />
Abate 1991<br />
Dodd 1917<br />
Abate 1991<br />
Kleinschmidt 1970; Bou‹ek 1988<br />
Hassan 1947<br />
Goot 1930<br />
Ho 1967; Yunus & Ho 1980<br />
Otanes y Quesales 1918<br />
Dodd 1917; Kleinschmidt 1970<br />
Abul-Nasr & Assem 1968<br />
Abate 1991<br />
Singh 1982; Ipe 1987<br />
Goot 1930<br />
Chu & Chou 1965<br />
Ooi 1973<br />
Litsinger 1987<br />
Plutarchia bicarinativentris Australia, PNG Dodd 1917; Bou‹ek 1988<br />
Plutarchia indefensa Thailand Burikam 1978<br />
242 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong>
Table 4.13.1 (contÕd) Natural enemies <strong>of</strong> Ophiomyia phaseoli<br />
Species<br />
HYMENOPTERA<br />
Country Reference<br />
PTEROMALIDAE<br />
Callitula filicornis Ethiopia Abate 1991<br />
Callitula viridicoxa Australia, PNG Bou‹ek 1988; Kleinschmidt 1970<br />
Callitula yasudi Japan Yasuda 1982<br />
Chlorocytus sp. India Kundu 1985<br />
Cryptoprymna sp. Egypt<br />
Taiwan<br />
Abul-Nasr & Assem 1968<br />
Chu & Chou 1965<br />
Halticoptera ?circulus Ethiopia Abate 1991<br />
Halticoptera patellana Hawaii Greathead 1975; Raros 1975<br />
Halticoptera sp. Egypt<br />
Taiwan<br />
Herbertia sp. Ethiopia<br />
East Africa<br />
Abul-Nasr & Assem 1968<br />
Chu & Chou 1965<br />
Abate 1991<br />
Greathead 1969<br />
Oxyharma subaenea Australia Dodd 1917; Kleinschmidt 1970<br />
Polycystus propinquus Sri Lanka Waterston 1915<br />
Polycystus sp. India Babu 1977<br />
Sphegigaster brunneicornis Ethiopia<br />
India<br />
Sphegigaster hamygurivara Japan Yasuda 1982<br />
Sphegigaster rugosa India<br />
Sri Lanka<br />
Abate 1991<br />
Peter & Balasubramanian 1984<br />
Ipe 1987<br />
Waterston 1915<br />
4.13 Ophiomyia phaseoli 243
Table 4.13.1 (contÕd) Natural enemies <strong>of</strong> Ophiomyia phaseoli<br />
Species<br />
HYMENOPTERA<br />
Country Reference<br />
PTEROMALIDAE (contÕd)<br />
Sphegigaster stella Malaysia<br />
Philippines<br />
Sphegigaster stepicola Ethiopia Abate 1991<br />
Sphegigaster voltairei Australia, PNG<br />
Egypt<br />
Indonesia<br />
Sphegigaster sp. Japan<br />
Philippines<br />
Taiwan<br />
Ho 1967<br />
Otanes y Quesales 1918<br />
Syntomopus shakespearei Australia Kleinschmidt 1970<br />
Syntomopus sp. Japan Yasuda 1982<br />
Unidentified Thailand Burikam 1978<br />
TETRACAMPIDAE<br />
Epiclerus sp. nr nomocerus Ethiopia Abate 1991<br />
A NEMATODE East Africa<br />
Thailand<br />
Dodd 1917; Kleinschmidt 1970; Bou‹ek 1988<br />
Hassan 1947<br />
Goot 1930<br />
Yasuda 1982<br />
Litsinger 1987<br />
Chu & Chou 1965<br />
Greathead 1969<br />
Burikam 1978<br />
244 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong>
EGYPT<br />
ETHIOPIA<br />
4.13 Ophiomyia phaseoli 245<br />
The bean fly was first reported in 1922, together with a parasitoid, later<br />
identified as Sphegigaster voltairei (= Trigonogastra agromyzae)<br />
(Pteromalidae). The eurytomid Eurytoma larvicola was also recorded<br />
(Hassan 1947). Later, Abul-Nasr and Assem (1968) recorded 5 parasitoid<br />
species emerging from bean fly puparia in the laboratory (Table 4.13.1), but<br />
no information was provided on their effectiveness.<br />
O. phaseoli was first reported in the early 1970s from Phaseolus vulgaris<br />
(French bean), Vigna unguiculata (cowpea) and soybean (Glycine max),<br />
although economic damage occurred only on French bean. The leguminous<br />
bush, Crotalaria laburnifolia was the only wild host which supported bean<br />
fly and its 17 species <strong>of</strong> parasitoids throughout the year (Table 4.13.1). Of<br />
these parasitoids, the pteromalids Sphegigaster stepicola and<br />
S. brunneicornis were the commonest species, accounting for up to 44.5%<br />
(average 26.2%) <strong>of</strong> total parasitisation on Crotalaria. Parasitisation by the<br />
braconid Opius phaseoli averaged a low 5.6% (range 0% to 23.2%) on<br />
Crotalaria, but it was the major parasitoid on French bean, accounting for<br />
over 87% <strong>of</strong> the total parasitisation. Abate (1991) concluded from this that<br />
the host plant plays an important role in Ophiomyia phaseoli population<br />
dynamics, although it remains to be shown whether this applies equally to<br />
the range <strong>of</strong> food legumes. It is, perhaps, relevant that the bean fly acts as a<br />
true leaf miner on Crotalaria, the larvae mining and eventually pupating<br />
within the leaf. This behaviour renders both stages more accessible to<br />
smaller parasitoids than when larvae are in the deeper tissues <strong>of</strong> a bean stem.<br />
The remaining 14 parasitoids recorded from Ethiopia (Table 4.13.1) were<br />
classified as very rare, (defined by Abate (1991) as equal to or less than 10%<br />
<strong>of</strong> total insect emergence) and together on average caused 9.6% mortality.<br />
Observations suggested that mortality <strong>of</strong> French bean seedlings caused by<br />
bean fly was much less severe in areas where the wild host, Crotalaria,<br />
occurred (14.8% in 1987 and 3.8% in 1988) than in its absence (39.1% in<br />
1987 and 36.1% in 1988 (Abate 1991)). This is circumstantial evidence that<br />
natural enemies play an important part in regulating bean fly populations.<br />
Negasi and Abate (1986) recorded a pteromalid Cyrtogaster sp. from<br />
O. phaseoli on French beans but, as this was not mentioned in the later, much<br />
more detailed paper on parasitoids by Abate (1991), it has not been included<br />
in Table 4.13.1.
246 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
HAWAII<br />
Damaging populations <strong>of</strong> O. phaseoli built up rapidly and caused<br />
widespread damage to cultivated legumes after it was first recorded in 1968.<br />
The only parasitoid found attacking it at that stage was the pteromalid<br />
Halticoptera patellana, a polyphagous European parasite <strong>of</strong> agromyzids<br />
(Greathead 1975).<br />
INDIA<br />
Ophiomyia phaseoli is said not to cause as serious damage to food legumes<br />
in India as it does elsewhere, especially in Indonesia and East Africa<br />
(Talekar 1990).<br />
The pteromalid Chlorocytus sp. was reported to parasitise 8% to 10% <strong>of</strong><br />
O. phaseoli puparia infesting stems <strong>of</strong> soybean (Glycine max) in the New<br />
Delhi area (Kundu 1985). In Bangalore, the pteromalid Sphegigaster<br />
brunneicornis, which emerged from bean fly puparia in cowpea, was the<br />
only parasitoid recorded. The extent <strong>of</strong> parasitisation ranged from 16.7% in<br />
July to 85.5% in September, but this did not achieve adequate control<br />
because bean fly infestation rose from 12% in July to 68% in September<br />
(Peter and Balasubramanian 1984). Ipe (1987) recorded four parasitoids (a<br />
eurytomid, Opius phaseoli, Tetrastichus sp. and Sphegigaster rugosa) from<br />
Agra. He commented that Ophiomyia phaseoli infestations are kept under<br />
control by parasitoids and that the percentage <strong>of</strong> parasitisation reaches<br />
appreciable levels each season. Babu (1977) reported Polycystus sp.<br />
emerging from bean fly puparia and causing parasitisation ranging from<br />
3.4% during February to 61% in August. However, the highest levels <strong>of</strong><br />
attack are those recorded at Agra by Singh (1982) for Opius phaseoli and<br />
Eurytoma sp.. The parasitisation <strong>of</strong> bean fly infesting cowpea reached 46.2%<br />
during early October, rising to 94% by the end <strong>of</strong> November. From<br />
December to March these parasites effectively controlled Ophiomyia<br />
phaseoli populations infesting cowpea, garden pea and Lablab niger (Singh<br />
1982).<br />
It appears that the different species <strong>of</strong> bean fly parasitoids in India may<br />
not occur at all widely. Whether this is due to climatic limitations, sampling<br />
from different hosts, or overall inadequate sampling <strong>of</strong> populations remains<br />
to be determined.<br />
INDONESIA<br />
The most comprehensive account <strong>of</strong> the biology, hosts and parasitoids <strong>of</strong><br />
bean fly in Indonesia was published in Dutch by Goot (1930). This was<br />
translated into English and republished in 1984 by the <strong>Asian</strong> Vegetable<br />
Research and Development Center Taiwan. A summary appears in<br />
Kalshoven (1981).
4.13 Ophiomyia phaseoli 247<br />
Three agromyzids are pests <strong>of</strong> soybean and some other economically<br />
important legumes in Indonesia: Ophiomyia phaseoli (by far the most<br />
important), the soybean stem borer Melanagromyza sojae (which bores into<br />
the pith <strong>of</strong> the stem, but seldom kills the plant) and the soybean top borer<br />
M. dolichostigma (which bores into the tops and causes stunting). The first<br />
two species have been known in Java since 1900.<br />
Four bean fly parasitoids are known (Goot 1930: Table 4.13.1), but their<br />
combined effect is generally low, parasitisation averaging 5.1%, with a<br />
maximum <strong>of</strong> 42.4%. All species emerge from the host pupa. The most<br />
effective species is the pteromalid Sphegigaster voltairei, which comprised<br />
59.1% <strong>of</strong> the parasitoids. It is also the most important parasitoid <strong>of</strong><br />
Melanagromyza sojae and occasionally attacks M. dolichostigma. Next in<br />
importance is the cynipid Cynipoide sp., contributing 10.5%: it also attacks<br />
the two other agromyzids. Finally, Eurytoma poloni and Eurytoma sp. each<br />
contribute 0.2% to the total. Both are more frequently bred from the two<br />
other agromyzids.<br />
MALAYSIA<br />
The bean fly was first reported in Peninsular Malaysia in 1924 and is<br />
regarded as the most important pest <strong>of</strong> green gram (Phaseolus aureus). It can<br />
also cause serious damage to French bean and other legume crops. Two<br />
parasitoids, which also occur in the Philippines, are Eurytoma poloni<br />
(Eurytomidae) and Sphegigaster (= Paratrigonogastra) stella (Ho 1967).<br />
Ooi (1973) recorded the eurytomid Plutarchia sp.<br />
PHILIPPINES<br />
The bean fly was first noticed in 1912, but was thought at the time to have<br />
been present for some years. Two parasitoids are known, a more abundant<br />
Eurytoma poloni (Eurytomidae) and a less abundant Sphegigaster stella, in<br />
the ratio 60:47. Their joint parasitisation averaged 17% with a range from<br />
6% to 49% (Otanes y Quesales 1918). Opius phaseoli was not recorded in<br />
this study.<br />
SRI LANKA<br />
Ophiomyia phaseoli was first recorded in 1901. Several unidentified species<br />
<strong>of</strong> hymenopterous parasitoids were bred by Rutherford (1914b) but,<br />
although they Ôno doubt do a considerable amount <strong>of</strong> goodÕ, they were<br />
unable to keep the fly in check.<br />
TAIWAN<br />
Chu and Chou (1965) reported a braconid (Opius sp.), 4 pteromalids<br />
(Cryptoprymna sp., Halticoptera sp., Sphegigaster sp. and Eucoilidea sp.)<br />
and a eurytomid (Eurytoma sp.) parasitoid attacking bean fly infesting<br />
soybean and Rose et al. (1976) added another eurytomid (Plutarchia sp.).
248 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Chiang et al. (1978) surveyed the parasitoids <strong>of</strong> three agromyzids<br />
infesting mungbean, namely Ophiomyia phaseoli, O. centrosematis and<br />
Melanagromyza sojae. Parasitisation fluctuated considerably, surpassing<br />
60% during July, but declining to nearly 0% in December and January. Since<br />
there was a negative correlation between agromyzid populations and percent<br />
parasitisation it was concluded that parasitoids played a role in controlling<br />
agromyzid populations.<br />
THAILAND<br />
Surveys in several regions <strong>of</strong> Thailand for natural enemies <strong>of</strong> the major pest<br />
<strong>of</strong> soybean (Ophiomyia phaseoli) revealed 5 species <strong>of</strong> hymenopterous<br />
parasitoid and a nematode (Burikam 1978). The most important species<br />
were Plutarchia indefensa (Eurytomidae) and Fopius sp. (Braconidae). A<br />
eulophid, a pteromalid and a cynipid were less important (Burikam and<br />
Napompeth 1979; Napompeth 1994). Parasitisation by P. indefensa<br />
averaged 52.8% and two samples gave 7.5% and 5.9% for Fopius sp.<br />
(Burikam 1978), although Napompeth (1994) later considered the two<br />
species to be <strong>of</strong> equal importance. The cynipid wasp also attacked the bean<br />
stem miner Melanagromyza sojae. On one occasion pupae containing 20 to<br />
50 nematodes were recorded, with a pupal parasitisation rate <strong>of</strong> 4.6%<br />
(Burikam 1978). A life table analysis showed that there was a densitydependent<br />
factor regulating bean fly populations (Burikam and Napompeth<br />
1979).<br />
Unless Fopius sp. (Braconidae) proves to be Opius phaseoli<br />
(Braconidae) (which is present in both India and the Philippines), it appears<br />
that O. phaseoli may not be widespread throughout <strong>Southeast</strong> Asia and may<br />
be well worth distributing more widely. The identity <strong>of</strong> the Fopius sp. is to<br />
be investigated (B. Napompeth pers. comm. 1994).<br />
ZIMBABWE<br />
Phaseolus spp. are the principal host crops damaged, but cowpeas (Vigna<br />
sinensis) and soybeans (Glycine soja) are also attacked. Plantings in late<br />
summer are usually only lightly infested and a braconid, identified as Opius<br />
liogaster (but quite possibly O. phaseoli), exercises effective control in most<br />
years (Taylor 1958). Earlier Jack (1913) had reported that a braconid larval<br />
parasitoid was ineffective in controlling bean fly although it was bred freely<br />
from O. phaseoli late in the season.
Attempts at biological control<br />
4.13 Ophiomyia phaseoli 249<br />
The only introductions for biological control <strong>of</strong> bean fly (Table 4.13.2) have<br />
been <strong>of</strong> the two braconid parasitoids Opius phaseoli and O. importatus from<br />
Uganda to Hawaii in 1969 (Davis 1971, 1972; Greathead 1975; Funasaki et<br />
al. 1988) and <strong>of</strong> O. phaseoli from Hawaii to Taiwan in 1974Ð75 (N.S.<br />
Talekar pers comm. 1994). In Hawaii both species rapidly became<br />
established on Oahu and host density was soon markedly reduced. They<br />
were introduced to other islands and, by 1971, on Kauai 100% <strong>of</strong> bean flies<br />
sampled produced parasitoids: on Maui rates ranged from 25% to 83%. No<br />
differences in the incidence <strong>of</strong> parasitisation were detected when<br />
infestations on French bean and cowpea were compared. By 1973<br />
O. importatus had become the dominant parasitoid and the polyphagous<br />
Halticoptera patellana (which was already present) was only rarely<br />
encountered (Greathead 1975). Surveys by Raros (1975) in 1973 and 1974<br />
<strong>of</strong> three locations on Oahu revealed average parasitisation ranging from<br />
8.3% to 23.5%, a result Talekar (1990) suggested might have been due to the<br />
heavy use <strong>of</strong> insecticides diminishing the earlier effectiveness <strong>of</strong> the<br />
parasitoids. In 1994 the bean fly was reported to be still a problem on young<br />
seedlings, so that farmers usually apply one or two insecticide sprays after<br />
seedlings emerge above the ground. Once the bean plant has developed a<br />
couple <strong>of</strong> leaves the fly is no longer a problem (W.C. Mitchell pers. comm.).<br />
Opius phaseoli was not recovered in Taiwan for the first two years after its<br />
introduction in 1974Ð75 from Hawaii, but there have been recent reports <strong>of</strong><br />
its presence, in spite <strong>of</strong> the current excessive use <strong>of</strong> insecticides against bean<br />
fly (N.S. Talekar pers. comm. 1994).
Table 4.13.2 Introductions for the biological control <strong>of</strong> Ophiomyia phaseoli<br />
Species<br />
BRACONIDAE<br />
Origin Liberated Year Result Reference<br />
Opius importatus Uganda Hawaii 1969 + Fischer 1971a; Davis 1971, 1972; Funasaki et<br />
al. 1988; Greathead 1975; Raros 1975<br />
Opius phaseoli Uganda<br />
Hawaii<br />
Hawaii<br />
Taiwan<br />
1969<br />
1974Ð75<br />
+<br />
+<br />
Fischer 1971a; Raros 1975<br />
N.S. Talekar pers. comm. 1994<br />
250 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong>
The more important parasitoids<br />
4.13 Ophiomyia phaseoli 251<br />
The names <strong>of</strong> a number <strong>of</strong> species are now different from those used by<br />
earlier authors. To enable cross referencing with those used in Table 4.13.1,<br />
the older names are shown below, together with a summary <strong>of</strong> information<br />
available on the biology <strong>of</strong> the more important species.<br />
Callitula viridicoxa (= Eurydinotellus viridicoxa = Polycystomyia<br />
beneficia) Hym.: Eurytomidae<br />
Chrysonotomyia (= Achrysocharis) douglasi Hym.: Eulophidae<br />
Chrysonotomyia ?erythraea Hym.: Eulophidae<br />
Chrysonotomyia formosa Hym.: Eulophidae<br />
The two latter species are widely distributed primary parasitoids attacking<br />
bean fly infesting Crotalaria in Ethiopia, parasitisation ranging from 0% to<br />
8.7% (average 2.6%). C. formosa has also been recorded from Liriomyza<br />
trifolii infesting beans in Guam (Schreiner et al. 1986; Abate 1991).<br />
Cynipoide sp. Hym.: Cynipoidea<br />
This parasitoid has only been reported from Java, where Goot (1930) found,<br />
from 90 samplings between 1919 and 1923, that it constituted 40% <strong>of</strong> the<br />
parasitoids reared, although the level <strong>of</strong> parasitisation varied greatly. It also<br />
emerged from puparia <strong>of</strong> Melanagromyza sojae and M. dolichostigma. It<br />
occurred in both tropical lowland and cool highland conditions.<br />
Euderus sp. Hym.: Eulophidae<br />
A tentative assignation as Euderus ?sp. was made by Napompeth (1994) <strong>of</strong><br />
the tiny eulophid recorded by Burikam (1978) and Burikam and Napompeth<br />
(1979). More than one parasitoid could be produced per bean fly host. The<br />
female parasitoid oviposited in the first instar host larva and pupation<br />
occurred during the hostÕs third instar, either within or alongside the host.<br />
The pupal stage averaged 7 days (Burikam 1978).<br />
Eurytoma poloni Hym.: Eurytomidae<br />
This parasitoid has been recorded from Indonesia, Malaysia and the<br />
Philippines, but little is known <strong>of</strong> its biology. It was recorded only once in<br />
Java from Ophiomyia phaseoli so it is clearly not an important parasitoid <strong>of</strong><br />
the bean fly there. In fact, it generally emerges from Melanagromyza<br />
dolichostigma. Adults live 22 to 28 days (Goot 1930).<br />
Fopius sp. Hym.: Braconidae<br />
This was referred to as Biosteres sp. by Burikam (1978) and Burikam and<br />
Napompeth (1979), but altered to Fopius sp. by Napompeth (1994). Adults<br />
emerged from bean fly puparia (one per host) and mated on the first day.<br />
Within two days 27 mature and immature eggs could be counted per female.
252 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Two samples in August revealed parasitisation levels <strong>of</strong> 5.9% and 7.5%<br />
(Burikam 1978).<br />
Halticoptera ?circulus Hym.: Pteromalidae<br />
This is a widespread primary parasitoid <strong>of</strong> agromyzid leafminers in many<br />
parts <strong>of</strong> the world. It was recorded as very rare (² 10% parasitisation) on<br />
bean fly in Ethiopia (Abate 1991).<br />
Hemiptarsenus varicornis (= Neodimmockia agromyzae and<br />
probably = Hemiptarsenus semialbicornis) Hym.: Eulophidae<br />
This is a very widespread parasitoid <strong>of</strong> dipterous leaf miners, including<br />
O. phaseoli and Liriomyza sativae. It occurs throughout tropical and<br />
southern temperate countries <strong>of</strong> the eastern hemisphere. In Australia it is<br />
common in many places along the eastern and southeastern coast. It also<br />
occurs in New Zealand, New Caledonia, Fiji and Vanuatu; Malaysia, Sri<br />
Lanka, India, Pakistan and Saudi Arabia; Senegal, Ghana, Sudan, Ethiopia,<br />
Kenya and Tanzania (Bou‹ek 1988).<br />
Meruana liriomyzae Hym.: Eulophidae<br />
This species was recorded as rare (³10% parasitisation) on bean fly infesting<br />
Crotalaria in Ethiopia (Abate 1991). It is also known from Liriomyza<br />
brassicae in Mauritius, L. sativae in Mauritius and RŽunion, from L. trifolii<br />
in Kenya, Chromatomyia horticola (= Phytomyza atricornis) in Ethiopia<br />
and South Africa, and from unidentified hosts in Australia and Zimbabwe<br />
(Bou‹ek 1988; Abate 1991).<br />
Opius importatus Hym.: Braconidae<br />
This species was first recorded from East Africa as Opius sp. by Greathead<br />
(1975) and later described as O. importatus by Fischer (1971b). In nature, it<br />
is known only from East Africa and only from Ophiomyia phaseoli. When<br />
first taken between November 1967 and April 1968 it attained parasitisation<br />
levels <strong>of</strong> 3% to 9% (Greathead 1969). However, later samples taken in<br />
Uganda in 1971 contained nearly 50% <strong>of</strong> O. importatus (Greathead 1975).<br />
The first instar larva was found in the third instar host larva and developed<br />
rapidly once the host pupated. Adults, that are similar in appearance to dark<br />
specimens <strong>of</strong> Opius phaseoli, emerge about 33 days after the appearance <strong>of</strong><br />
the host plant above the soil. O. importatus was inadvertently included in<br />
shipments <strong>of</strong> Opius phaseoli to Hawaii, where it soon became the dominant<br />
parasitoid <strong>of</strong> Ophiomyia phaseoli (Greathead 1975).<br />
Opius phaseoli (= O. melanagromyzae) Hym.: Braconidae<br />
O. phaseoli was originally described from the Philippines (Manila) without<br />
a host by Ashmead (1904) as Eurytenes nanus. However, as this name was<br />
preoccupied, it was redescribed as O. phaseoli by Fischer (1963), who listed
4.13 Ophiomyia phaseoli 253<br />
its host as Ophiomyia phaseoli and its distribution as India (Nagpur) and the<br />
Philippines. Later, Fischer (1966) listed it as a parasitoid <strong>of</strong> the leaf miner<br />
Melanagromyza atomella in India and Singh (1982) from Melanagromyza<br />
sojae. Both Singh (1982) and Ipe (1987), working in the area <strong>of</strong> Agra,<br />
considered it to be important in regulating bean fly populations there. No<br />
information is available on its presence or effectiveness elsewhere in Asia or<br />
<strong>Southeast</strong> Asia, except for the original record <strong>of</strong> a single female wasp from<br />
Manila (Ashmead 1904).<br />
In East Africa, Greathead (1969, 1975) reported parasitisation levels <strong>of</strong><br />
Ophiomyia phaseoli by Opius phaseoli that were frequently above 50%, and<br />
sometimes reached 94.4%. Levels <strong>of</strong> up to 10% on Ophiomyia spencerella<br />
were also recorded and Opius phaseoli was twice reared from Ophiomyia<br />
centrosematis. He concluded that Opius phaseoli was the chief biotic factor<br />
limiting the population <strong>of</strong> bean fly in East Africa. Nevertheless, he reported<br />
levels <strong>of</strong> only 38% parasitisation from 3 bean fly samples taken in<br />
Madagascar and only about 20% in Mauritius. He suggested that the latter<br />
result might be due to a different strain <strong>of</strong> the parasitoid or, perhaps, even a<br />
different species. In Ethiopia Abate (1991) reported over 93% parasitisation<br />
<strong>of</strong> bean fly on French bean, but much lower levels (average 5.6%) on a wild<br />
host Crotalaria.<br />
One to five (but generally two or three) eggs are laid at a time, usually in<br />
first instar host larvae. Hatching occurs about 2 days later, soon after the host<br />
larva has moulted to the second instar. The first instar parasitoid larva grows,<br />
but does not moult until its host has pupated. Meanwhile, all except one<br />
Opius larva are suppressed. Development is rapid in the host pupa, the entire<br />
larval period lasting 9 to 10 days, leading to a prepupal period <strong>of</strong> 1 to 2 days<br />
and a pupal period (within the host puparium) <strong>of</strong> about 3 days (Raros 1975).<br />
In East Africa, the pupal period lasts a minimum <strong>of</strong> 4 days and adults<br />
commence emerging about 30 days after the appearance <strong>of</strong> the host plant<br />
above the ground (Greathead 1969). On average, males live 20 and females<br />
23 days. Females mate a day after emergence and, following a<br />
preoviposition period <strong>of</strong> 1 to 2 days, may lay up to 358 eggs throughout life.<br />
First instar host larvae are preferred to second instar larvae (62.9:32.8)<br />
(Raros 1975). Other host stages are not attacked. Adult parasitoids feed on<br />
water droplets and host plant sap resulting from oviposition punctures made<br />
either by host adults or by female parasitoids. The male:female sex ratio was<br />
about 1:1.4 in Hawaii (Raros 1975).
254 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Oxyharma (= Pterosema) subaenea Hym.: Pteromalidae<br />
Plutarchia indefensa Hym.: Eurytomidae<br />
This wasp was referred to by Burikam (1978) and Burikam and Napompeth<br />
(1979) as Plutarchia sp. Eggs are laid in third instar larvae <strong>of</strong> Ophiomyia<br />
phaseoli, usually at the posterior end. One or two eggs are laid per host and,<br />
on hatching after 2 to 3 days, one surviving larva remains in the first instar<br />
until the host has pupated. The larval stage lasts 5 to 7 days and the pupal<br />
stage (inside the host puparium) 7 to 8 days, giving a development period<br />
from egg to adult <strong>of</strong> 16 to 19 days. Adult males lived 4 to 19 days (average<br />
11.5) and females 10 to 25 days (average 16.9), during which 6 to 14 eggs<br />
developed per day (Burikam 1978).<br />
Pteromalid Hym.: Pteromalidae<br />
The unidentified pteromalid recorded from Thailand (Burikam 1978;<br />
Burikam and Napompeth 1979; Napompeth 1994) was found parasitising<br />
pupae <strong>of</strong> O. phaseoli. The female laid 1 or 2 eggs in the host puparium. These<br />
hatched in 2 days and, after 4 days larval development, pupation occurred<br />
within the host puparium. The pupal stage lasted 7 to 8 days, resulting in a<br />
life cycle <strong>of</strong> 12 to 14 days. Female wasps lived more than 2 weeks (Burikam<br />
1978).<br />
Sphegigaster brunneicornis Hym.: Pteromalidae<br />
This species has been reported from Ethiopia (Abate 1991), India and Sri<br />
Lanka (Peter and Balasubramanian 1984). O. phaseoli is its only recorded<br />
host.<br />
Sphegigaster (= Trigonogastra) rugosa Hym.: Pteromalidae<br />
Sphegigaster (= Paratrigonogastra) stella Hym.: Pteromalidae<br />
Sphegigaster stepicola Hym.: Pteromalidae<br />
This species is known from Ophiomyia phaseoli in Ethiopia (Abate 1991)<br />
and from Phytomyza albiceps in southern Europe and India (Abate 1991).<br />
Combined parasitisation with S. brunneicornis <strong>of</strong> bean fly on Crotalaria in<br />
Ethiopia ranged from 3.1% to 44.4% (average 26.2%), <strong>of</strong> which S. stepicola<br />
accounted for nearly 72% (Abate 1991).<br />
Sphegigaster voltairei (= Sphegigaster agromyzae<br />
= Trigonogastra agromyzae) Hym.: Pteromalidae<br />
This species is recorded from Australia, Papua New Guinea, Egypt and<br />
especially Indonesia where Goot (1930) reported that, on average, it<br />
comprised 60% <strong>of</strong> emergences from parasitised puparia and that it could be<br />
kept alive from 30 to 48 days. It was also the most important parasitoid <strong>of</strong><br />
Melanagromyza sojae and was bred several times from M. dolichostigma.<br />
Syntomopus (= Merismorella) shakespearei Hym.: Pteromalidae
Comment<br />
4.13 Ophiomyia phaseoli 255<br />
Although O. phaseoli occurs in Africa, it now seems unlikely that it evolved<br />
there. It was reported in Uganda as recently as the 1920s, in Tanzania in<br />
1937 and in Kenya in 1939. In East Africa, it occurs in close association with<br />
the very similar O. spencerella, which is generally the dominant species<br />
(Greathead 1969). Spencer (1973) postulated that O. spencerella evolved in<br />
Africa and O. phaseoli in Asia. The latter subsequently arrived in Africa to<br />
occupy a similar niche in much the same host plants, but isolated<br />
reproductively from O. spencerella. It is interesting that the major natural<br />
enemy <strong>of</strong> Ophiomyia phaseoli in East Africa is the braconid Opius phaseoli,<br />
which also occurs in India, whereas that <strong>of</strong> Ophiomyia spencerella is a<br />
cynipid Eucoilidea sp.. Although Eucoilidea sp. is not restricted to<br />
Ophiomyia spencerella, this species nevertheless does not attack Ophiomyia<br />
phaseoli.<br />
In spite <strong>of</strong> Asia (and probably India) being nominated as the probable<br />
centre <strong>of</strong> origin <strong>of</strong> Ophiomyia phaseoli, it is interesting that bean fly is even<br />
more heavily parasitised in East Africa and Ethiopia than anywhere else Ñ<br />
and by 2 braconid parasitoids, one <strong>of</strong> which is native to the latter regions.<br />
These are Opius phaseoli and O. importatus. Opius phaseoli was described<br />
from a single female from Manila, but the absence <strong>of</strong> records <strong>of</strong> it attacking<br />
Ophiomyia phaseoli there or elsewhere in <strong>Southeast</strong> Asia raises doubts that<br />
it is native to <strong>Southeast</strong> Asia.<br />
The rapid control <strong>of</strong> Ophiomyia phaseoli in Hawaii following the<br />
introduction <strong>of</strong> Opius phaseoli and O. importatus (the latter soon becoming<br />
the dominant species) demonstrates that, under favourable circumstances,<br />
biological control alone can produce valuable results. It seems probable that<br />
this success would be repeated, at least in other Pacific island nations.<br />
In the far more complex ecological environment in <strong>Southeast</strong> Asia<br />
extrapolation from experience in Hawaii is more risky. It is probable that the<br />
other components <strong>of</strong> integrated pest management (varietal resistance,<br />
cultural methods, rational pesticide use, etc.) will all be required to<br />
supplement the reduction in bean fly density that can be brought about by<br />
parasitoids. Nevertheless, any substantial decrease in bean fly populations<br />
that can be achieved by introducing additional effective parasitoids is likely<br />
to be a valuable contribution towards reduced crop losses.
Table 14<br />
Table 14 shows for weeds what Table 4 did for<br />
invertebrates in relation to the top 10 entries.<br />
Since no information was available on the<br />
relative rating for the five weeds nominated by<br />
Tokelau (shown by an asterisk), each was<br />
70<br />
D.F. Waterhouse<br />
allocated the median value <strong>of</strong> 5. The ranking<br />
order is only given for species that attain an<br />
aggregated value <strong>of</strong> 10 or more. These are<br />
arranged in descending order <strong>of</strong> importance in<br />
Table 15.
The Major Invertebrate <strong>Pests</strong> and Weeds <strong>of</strong> Agriculture and Plantation Forestry in the Southern and Western Pacific<br />
71<br />
Table 14<br />
The<br />
relative importance given to the top 10 weeds <strong>of</strong> agriculture (72 species) <strong>of</strong> each country in the southern and western Pacific .<br />
Name Family CoI Fij FrP FSM Gua Kir Mar NCa Niu PNG ASa WSa SoI Tok Ton Tuv Van W.F. No. * Rating Order<br />
Acacia farnesiana<br />
Acacia nilotica<br />
Achyranthes aspera<br />
Agave americana<br />
Ageratum conyzoides<br />
Alternanthera sessilis<br />
Amaranthus interruptus<br />
Amaranthus spinosus<br />
Amaranthus viridis<br />
( = A. gracilis)<br />
Antigonon leptopus<br />
Argemone mexicana<br />
( = A. americana)<br />
Bidens alba, Bidens<br />
pilosa<br />
Blechum pyrimidatum<br />
( = B. brownei)<br />
Brachiaria mutica<br />
Brachiaria reptans<br />
Brachiaria<br />
subquadripara<br />
Broussonetia papyrifera<br />
Canavalia rosea<br />
Cardiospermum<br />
halicacabum<br />
Cassytha filiformis<br />
Cecropia peltata<br />
Cenchrus echinatus<br />
Chamaesyce<br />
( = Euphorbia) hirta<br />
Chloris barbata<br />
Mimosaceae 4 1 7<br />
Mimosaceae<br />
Amaranthaceae<br />
Agavaceae<br />
Asteraceae 2 1 9<br />
Amaranthaceae<br />
Amaranthaceae<br />
Amaranthaceae<br />
Amaranthaceae<br />
Polygonaceae<br />
Papaveraceae<br />
2 1 9<br />
Asteraceae 7 9 3 8 1 9 6 29 6 =<br />
Acanthaceae 8 1 3<br />
Poaceae 8 1 3<br />
Poaceae<br />
Poaceae<br />
Urticaceae 4 1 7<br />
Fabaceae 10 1 1<br />
Sapindaceae 2 1 9<br />
Lauraceae 7 2 6 3 18 13<br />
Euphorbiaceae 5 1 6<br />
Poaceae 3 * 3 3 21 12<br />
Euphorbiaceae 10 10 * 3 7 45=<br />
Poaceae
72<br />
D.F. Waterhouse<br />
Table 14<br />
Chromolaena odorata<br />
Clerodendrum chinense<br />
( = C. philippinum)<br />
Clidemia hirta<br />
Coccinia grandis<br />
Commelina<br />
benghalensis<br />
Commelina diffusa<br />
Cordia subcordata<br />
Crassocephalum<br />
crepidoides<br />
Crotolaria pallida<br />
Crotolaria retusa<br />
Cuphea carthagenensis<br />
Cynodon dactylon<br />
Cyperus rotundus<br />
Dactyloctenium<br />
aegyptium<br />
Desmodium incanum<br />
Digitaria ciliaris<br />
Digitaria eriantha<br />
( = D. decumbens)<br />
Digitaria insularis<br />
Digitaria setigera<br />
Echinochloa colona<br />
Echinochloa crus-galli<br />
Eichhornia crassipes<br />
Eleocharis geniculata<br />
Elephantopus mollis<br />
( = E. scaber)<br />
Eleusine indica<br />
(cont’d)<br />
Asteraceae 3 7 2 12 21=<br />
Verbenaceae 2 3 1 3 27 9<br />
Melastomataceae 4 5 2 13 20<br />
Cucurbitaceae 5 1 6<br />
Commelinaceae 2 1 9<br />
Commelinaceae<br />
Boraginaceae<br />
9 6 2 7<br />
Asteraceae 10 1 1<br />
Fabaceae<br />
Fabaceae<br />
Lythraceae<br />
Poaceae<br />
Cyperaceae<br />
Poaceae<br />
7 6 8 7 1 1 5 4 1 1 1 9 1 13 98 1<br />
Fabaceae<br />
Poaceae<br />
Poaceae<br />
Eleutheranthera ruderalis Asteraceae<br />
The<br />
relative importance given to the top 10 weeds <strong>of</strong> agriculture (72 species) <strong>of</strong> each country in the southern and western Pacific .<br />
Name Family CoI Fij FrP FSM Gua Kir Mar NCa Niu PNG ASa WSa SoI Tok Ton Tuv Van W.F. No. * Rating Order<br />
Poaceae<br />
Poaceae<br />
Poaceae<br />
10 1 1<br />
Poaceae 6 1 5<br />
Pontederiaceae<br />
Cyperaceae<br />
7 1 2 8 4 26 10<br />
Asteraceae 9 3 2 10 26=<br />
Poaceae 8 9 6 10 * 10 7 7 21 11
The Major Invertebrate <strong>Pests</strong> and Weeds <strong>of</strong> Agriculture and Plantation Forestry in the Southern and Western Pacific<br />
73<br />
Table 14<br />
Emilia sonchifolia<br />
Eragrostis tenella<br />
Euphorbia heterophylla<br />
( = E. geniculata)<br />
Fimbristylis cymosa<br />
( = F. atollensis)<br />
Fimbristylis dichotoma<br />
Fimbristylis miliacea<br />
Guettarda speciosa<br />
Hydrilla verticillata<br />
Hyptis pectinata<br />
Imperata conferta<br />
( = I. cylindrica)<br />
Indig<strong>of</strong>era suffruticosa<br />
Ipomoea macrantha<br />
Ischaemum spp.<br />
Jatropha gossypifolia<br />
Kyllinga brevifolia<br />
Kyllinga nemoralis<br />
Kyllinga polyphylla<br />
Lantana camara<br />
Leucaena leucocephala<br />
Ludwigia octovalvis<br />
( = Jussiaea suffruticosa)<br />
Asteraceae<br />
Poaceae<br />
Euphorbiaceae 8 1 3<br />
Cyperaceae 7 1 4<br />
Cyperaceae<br />
Cyperaceae<br />
Cyperaceae 4 * 2 12 21=<br />
Hydrocharitaceae<br />
Lamiaceae 3 1 8<br />
Poaceae 10 1 1<br />
Fabaceae<br />
Convolvulaceae * 5 2 11 23 =<br />
Poaceae 10 1 1<br />
Euphorbiaceae<br />
Cyperaceae<br />
Cyperaceae<br />
Macroptilium lathyroides Fabaceae<br />
Melaleuca<br />
quinquenervia<br />
Merremia peltata<br />
Miconia calvescens<br />
Mikania micrantha<br />
Mimosa invisa<br />
(cont’d)<br />
The<br />
relative importance given to the top 10 weeds <strong>of</strong> agriculture (72 species) <strong>of</strong> each country in the southern and western Pacific .<br />
Name Family CoI Fij FrP FSM Gua Kir Mar NCa Niu PNG ASa WSa SoI Tok Ton Tuv Van W.F. No. * Rating Order<br />
Cyperaceae 3 9 2 10 26 =<br />
Verbenaceae 8 6 6 7 1 6 3 7 40 4<br />
Mimosaceae 2 8 10 3 13 19<br />
Onagraceae 5 2 2 15 15 =<br />
Myrtaceae<br />
Convolvulaceae 1 10 2 11 23 =<br />
Melastomataceae 1 1 10 28 =<br />
Asteraceae 6 9 4 8 6 2 6 7 8 5 10 49 3<br />
Mimosaceae 1 1 3 4 7 4 3 7 5 2 10 73 2
74<br />
D.F. Waterhouse<br />
Table 14 (cont’d) The relative importance given to the top 10 weeds <strong>of</strong> agriculture (72 species) <strong>of</strong> each country in the southern and western Pacific .<br />
Mimosa pigra<br />
Mimosa pudica<br />
Miscanthus floridulus<br />
Momordica charantia<br />
Monochoria hastata<br />
Nephrolepis hirsutula<br />
Ocimum gratissimum<br />
Oxalis corniculata<br />
Panicum maximum<br />
Parthenium<br />
hysterophorus<br />
Name Family CoI Fij FrP FSM Gua Kir Mar NCa Niu PNG ASa WSa SoI Tok Ton Tuv Van W.F. No. * Rating Order<br />
Paspalum conjugatum<br />
Paspalum dilatatum<br />
Paspalum paniculatum<br />
Paspalum vaginatum<br />
Passiflora foetida<br />
Passiflora maliformis<br />
Mimosaceae 3 1 8<br />
Mimosaceae 5 6 4 6 9 2 6 40 5<br />
Poaceae<br />
Cucurbitaceae<br />
Pontederiaceae<br />
Davalliaceae<br />
Lamiaceae 5 1 6<br />
Oxalidaceae<br />
Poaceae 3 1 5<br />
Asteraceae 7 1 4<br />
Poaceae 5 8 2 9<br />
Poaceae<br />
Poaceae<br />
Poaceae 5 1 6<br />
Passifloraceae<br />
Passifloraceae<br />
Pennisetum polystachion Poaceae 2 1 9<br />
Pennisetum purpureum Poaceae 2 1 9<br />
Phyllanthus amarus Euphorbiaceae<br />
Physalis angulata Solanaceae<br />
Pistia stratiotes Araceae<br />
Pluchea indica Asteraceae<br />
Portulaca oleracea Portulacaceae 5 3 2 14 18<br />
Premna obtusifolia<br />
( = P. serratifolia)<br />
Verbenaceae 1 1 10 28 =<br />
Pseudelephantopus<br />
spicatus<br />
Asteraceae<br />
Psidium guajava Myrtaceae 3 1 8<br />
Ricinus communis Euphorbiaceae 6 1 5
The Major Invertebrate <strong>Pests</strong> and Weeds <strong>of</strong> Agriculture and Plantation Forestry in the Southern and Western Pacific 75<br />
Table 14 (cont’d) The relative importance given to the top 10 weeds <strong>of</strong> agriculture (72 species) <strong>of</strong> each country in the southern and western Pacific .<br />
Rottboellia<br />
cochinchinensis<br />
Name Family CoI Fij FrP FSM Gua Kir Mar NCa Niu PNG ASa WSa SoI Tok Ton Tuv Van W.F. No. * Rating Order<br />
Ruellia prostrata Acanthaceae<br />
Poaceae 5 2 2 15 15 =<br />
Salvinia molesta Salviniaceae<br />
Scaveola sericea<br />
( = S. taccada)<br />
Goodeniaceae 7 4 2 11 23 =<br />
Schinus terebinthifolius Anacardiaceae<br />
Senna ( = Cassia)<br />
occidentalis<br />
Caesalpinaceae<br />
Senna ( = Cassia) tora Caesalpinaceae 8 6 2 8<br />
Sida acuta Malvaceae 9 9 8 4 4 14 17<br />
Sida cordifolia Malvaceae<br />
Sida fallax Malvaceae 8 1 3<br />
Sida rhombifolia Malvaceae 8 9 7 3 9<br />
Solanum americanum<br />
( = S. nigrum)<br />
Solanaceae<br />
Solanum mauritianum Solanaceae<br />
Solanum torvum Solanaceae 2 9 5 1 4 27 8<br />
Sonchus oleraceus Asteraceae<br />
Sorghum arundinaceum<br />
( = S. verticilliflorum)<br />
Poaceae 4 1 7<br />
Sorghum halepense Poaceae 9 4 4 3 16 14<br />
Sorghum sudanense Poaceae 10 1 1<br />
Spathodea<br />
companulata<br />
Bignoniaceae 4 1 7<br />
Sphaerostaphanos<br />
invisus<br />
Thelypteridaceae<br />
Sphaerostephanos<br />
unitus<br />
Thelypteridaceae<br />
Stachytarpheta<br />
cayennensis<br />
Verbenaceae<br />
Stachytarpheta<br />
jamaicensis<br />
Verbenaceae 10 6 2 6
76 D.F. Waterhouse<br />
Table 14 (cont’d) The relative importance given to the top 10 weeds <strong>of</strong> agriculture (72 species) <strong>of</strong> each country in the southern and western Pacific .<br />
Name Family CoI Fij FrP FSM Gua Kir Mar NCa Niu PNG ASa WSa SoI Tok Ton Tuv Van W.F. No. * Rating Order<br />
Stachytarpheta urticifolia Verbenaceae 10 5 7 10 4 12 20<br />
Stictocardia tiliifolia Convolvulaceae<br />
Syndrella nodiflora Asteraceae<br />
Tecoma stans Bignoniaceae 4 1 7<br />
Themeda quadrivalis Poaceae<br />
Tournefortia<br />
( = Messerschmidia)<br />
argentea<br />
Boraginaceae 9 1 2<br />
Tribulus cistoides Zygophyllaceae<br />
Tridax procumbens Asteraceae<br />
Triumfetta rhomboidea Tiliaceae<br />
Urena lobata Malvaceae<br />
Vernonia cinerea Asteraceae 9 1 2<br />
Vigna marina Fabaceae 10 1 1<br />
Vitex trifolia Verbenaceae<br />
Wedelia trilobata Asteraceae<br />
Xanthium pungens Asteraceae 3 1 8
4.15 Planococcus citri<br />
India<br />
20°<br />
Myanmar<br />
P Laos<br />
0°<br />
20°<br />
China<br />
++<br />
Thailand<br />
+<br />
Cambodia<br />
Vietnam<br />
+++<br />
+<br />
+ Brunei<br />
Malaysia<br />
Singapore<br />
+<br />
Indonesia<br />
Taiwan<br />
++<br />
P<br />
Philippines<br />
Australia<br />
Papua<br />
New Guinea<br />
+<br />
287<br />
It is speculated that the citrus mealybug Planococcus citri is <strong>of</strong> south China origin,<br />
although it now occurs very widely in tropical, subtropical and temperate regions wherever<br />
citrus is grown. Like other mealybugs, it is attacked by a large number <strong>of</strong> non-specific<br />
predators, especially Coccinellidae, but also Chrysopidae. These consume vast numbers<br />
<strong>of</strong> prey when mealybugs are abundant, but <strong>of</strong>ten do not reduce host numbers to a level at<br />
which economic injury no longer occurs. There are several specific or near specific encyrtid<br />
parasitoids that are capable <strong>of</strong> lowering P. citri populations below the economic threshold.<br />
These are worth serious consideration for introduction to regions where they do not already<br />
occur: Leptomastix dactylopii (<strong>of</strong> Brazilian origin), Leptomastidea abnormis and Anagyrus<br />
pseudococci (<strong>of</strong> Mediterranean origin) and Coccidoxenoides peregrinus (<strong>of</strong> south China or<br />
Indian origin). In some situations (particularly cooler conditions) augmentative releases<br />
are necessary if the use <strong>of</strong> insecticides is to be avoided.<br />
20°<br />
0°<br />
20°
288 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Planococcus citri (Risso)<br />
Rating<br />
Origin<br />
Distribution<br />
Hemiptera, Pseudococcidae<br />
citrus mealybug<br />
This account draws heavily on that <strong>of</strong> Bartlett (1978) and CABI abstracts<br />
since then.<br />
.<br />
<strong>Southeast</strong> Asia China Southern and Western Pacific<br />
+++ Viet<br />
7 ++ ++ 4<br />
+ Thai, Msia, Brun,<br />
+ Cook Is, PNG, Tong,<br />
Indo<br />
Sam<br />
P Myan, Phil P Fr P, Niue<br />
P. citri was described from citrus in southern France (Risso 1813), but<br />
Bartlett (1978) speculates that it is <strong>of</strong> Chinese origin. However, the fact that<br />
it is a widespread and important pest <strong>of</strong> citrus in 11 <strong>of</strong> the 14 provinces <strong>of</strong><br />
southern China casts some doubt on this view (Li Li-ying et al. 1997).<br />
The citrus mealybug is extremely widespread, being present in almost all<br />
tropical, subtropical and temperate regions <strong>of</strong> the world and in many<br />
glasshouses in cooler parts. Other species <strong>of</strong> Planococcus have <strong>of</strong>ten been<br />
confused with it, including P. pacificus (which does not occur in the<br />
Mediterranean but is common in the Pacific: Cox 1981, 1989), P. ficus<br />
(restricted to fig, pomegranate and grape in the Mediterranean: Tranfaglia<br />
1979; Cox and Freeston 1985; Cox and Ben-Dov 1986) and P. kenyae (on<br />
C<strong>of</strong>fea in Kenya: Le Pelley 1943a,b). However, their hymenopterous<br />
parasitoids discriminate between P. citri and P. kenyae and biological<br />
control fails when the incorrect parasites are used (Rosen and De Bach<br />
1977). The collection data <strong>of</strong> Pacific specimens suggest that P. citri is a<br />
recent introduction there (Williams 1982). There may well be different<br />
strains <strong>of</strong> P. citri.<br />
For example, in Brazil it is rarely found on citrus, but is<br />
common on other plants (Compere 1939); and in South Africa it is common<br />
on citrus, but seldom found on grape vines (De Lotto 1975).
Biology<br />
Host plants<br />
4.15<br />
Planococcus citri<br />
289<br />
P. citri shows considerable morphological variation when reared under<br />
different environmental conditions. Small specimens, produced by rearing<br />
at high temperatures (32¡C), have smaller appendages and lower numbers <strong>of</strong><br />
cuticular structures than those reared between 17¡ and 25¡C (Cox 1981).<br />
Adult females are oval, flat and yellow to yellowish brown, with a barely<br />
visible dorsal line under their waxy covering. Along the edge <strong>of</strong> the wax<br />
cover there are short waxy protruberances, the longest <strong>of</strong> which are at the<br />
posterior end. Yellow eggs are deposited in an ovisac <strong>of</strong> wax threads and<br />
young nymphs are lemon-yellow in colour. Females are oviparous, and<br />
possibly parthenogenetic when males are not available.<br />
The development time varies from 20 to 40 days, depending upon the<br />
host plant and temperature. The pre-oviposition period is 7 to 10 days, eggs<br />
hatch in 3 to 6 days and, 300 to 500 eggs are laid per female. Additional<br />
details <strong>of</strong> developmental periods on c<strong>of</strong>fee and potato sprouts are provided<br />
by Bartlett and Lloyd (1958) and Martinez and Suris (1987a, b). Eggs,<br />
nymphs and adults are capable <strong>of</strong> overwintering. There are 3 instars for<br />
female nymphs and 2 for male nymphs which then form a waxy puparium. In<br />
warm areas there are normally 4 or 5 overlapping generations per year<br />
(Bartlett 1978; Kalshoven 1981), but about double this number under<br />
laboratory conditions (Gray 1954).<br />
P. citri can be readily reared in the laboratory on potato sprouts (Fisher<br />
1963; Martinez and Suris 1987b), lemons and butternut pumpkins<br />
(Samways and Mapp 1983) and a crawler-pro<strong>of</strong> cage has been described<br />
(Rao 1989). Dispersal <strong>of</strong> crawlers is brought about by wind, rain and ants.<br />
Until mated, females secrete a pheromone continously to attract males:<br />
(1Rcis) (3-isopropenyl-2-2-dimethylcyclobutyl) methyl acetate (Dunkelblum<br />
et al. 1986). Synthetic analogues have been tested to reduce<br />
populations (Rotundo and Tremblay 1974, 1980, 1982; Rotundo et al. 1979).<br />
P. citri<br />
has an extremely wide host range, attacking almost every flowering<br />
plant and some grasses as well. It is most frequently reported from citrus, and<br />
occurs commonly on other fruit trees and grape vines. Other important crops<br />
include banana, tobacco, c<strong>of</strong>fee, fig, mango, cocoa, date, casssava,<br />
macadamia, passionfruit and cut flowers.
290 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Damage<br />
The citrus mealybug is a widespread and severe pest <strong>of</strong> citrus; also, in<br />
temperate areas, <strong>of</strong> grape vines and, in tropical areas, <strong>of</strong> c<strong>of</strong>fee and mango.<br />
P. citri is the most injurious <strong>of</strong> 6 species <strong>of</strong> mealybug in the Mediterranean<br />
basin, the damage caused, especially to fruit, being so severe in France that<br />
the economic threshold is 2% infestation (Panis 1979). In glasshouses its<br />
hosts include bulbs, ferns, gardenias and other ornamentals. It is generally<br />
found on the aerial parts <strong>of</strong> plants, although a root form occurs. Tender<br />
growing tips, flower buds and young fruit clusters are favoured aggregating<br />
points. Excessive removal <strong>of</strong> sap by large numbers <strong>of</strong> mealybugs leads to<br />
wilting and death <strong>of</strong> shoots and flower buds and to drop <strong>of</strong> fruit.<br />
Furthermore, there is abundant growth <strong>of</strong> sooty moulds on the large amounts<br />
<strong>of</strong> honeydew produced, which may render produce unmarketable. P. citri<br />
becomes most abundant during the dry season.<br />
P. citri has been incriminated in the transmission <strong>of</strong> viruses <strong>of</strong> grape<br />
vines, cocoa, cucumber, taro and tobacco (Carpenter et al. 1976; Kenten and<br />
Woods 1976; Bartlett 1978; Legg and Lockwood 1981; Dufour 1988; Agran<br />
et al. 1990; Pedroso et al. 1991).<br />
Natural enemies<br />
The citrus mealybug, like other mealybugs, is attacked by a wide range <strong>of</strong><br />
naturally occurring predators, particularly coccinellid beetles and lacewings<br />
(Table 4.15.1). Many <strong>of</strong> these have a wide to very wide host range and thus<br />
are less likely nowadays than in the past to be regarded as suitable to<br />
introduce to new environments. Nevertheless, those reported in recent<br />
literature are recorded, but no attempt has been made to provide an<br />
exhaustive list.<br />
The coccinellid predator Cryplolaemus mountrouzieri in particular has<br />
been very extensively distributed in the past, <strong>of</strong>ten with moderate to good<br />
success, against a variety <strong>of</strong> mealybug, scale and aphid pests, including<br />
P. citri.<br />
C. montrouzieri and some <strong>of</strong> the other predators play an important<br />
role in greatly reducing the abundance <strong>of</strong> high populations. In general,<br />
however, their searching ability is poor and prey are missed when population<br />
densities fall, so their action must generally be supplemented by parasitoids<br />
or other means for effective control.
Table 4.15.1<br />
Predators <strong>of</strong> Planococcus citri<br />
Species<br />
HEMIPTERA<br />
ANTHOCORIDAE<br />
Country Reference<br />
Orius minutus<br />
NEUROPTERA<br />
CHRYSOPIDAE<br />
Turkey Soylu & Urel 1977<br />
Brinckochrysa (= Chrysopa)<br />
scelestes India Krishnamoorthy 1984<br />
Chrysopa sp. Australia<br />
USA<br />
Chrysoperla carnea<br />
India<br />
Israel<br />
USSR<br />
Mallada (= Anisochrysa) basalis<br />
Mallada boninensis<br />
Odontochrysa (= Chrysopa =Plesiochrysa )<br />
lacciperda<br />
Oligochrysa lutea<br />
Sympherobius barberi<br />
Sympherobius sanctus<br />
COLEOPTERA<br />
COCCINELLIDAE<br />
Brumoides lineatus<br />
Brumus suturalis<br />
Murray 1978<br />
Meyerdirk et al. 1982<br />
Krishnamoorthy & Mani 1989b<br />
Berlinger et al. 1979<br />
Niyazov 1969<br />
India Krishnamoorthy & Mani 1989b<br />
India Krishnamoorthy & Mani 1989b<br />
India Mani & Krishnamoorthy 1990<br />
Australia Murray 1978, 1982<br />
USA Dean et al. 1971; Meyerdirk et al. 1979, 1982<br />
Turkey Soylu & Urel 1977<br />
China Weng & Huang 1988<br />
Indonesia Kalshoven 1981<br />
4.15<br />
Planococcus citri<br />
291
Table 4.15.1<br />
Species<br />
COLEOPTERA<br />
Country Reference<br />
COCCINELLIDAE (contÕd)<br />
Chilocorus bipustulatus<br />
(contÕd) Predators <strong>of</strong> Planococcus citri<br />
Israel<br />
Turkey<br />
Berlinger et al. 1979<br />
Soylu & Urel 1977<br />
Coccinella californica<br />
California Bartlett & Lloyd 1958<br />
Coccinella transversalis (= C. repanda)<br />
Indonesia Kalshoven 1981<br />
Coccinella semipunctata<br />
USSR Niyazov 1969<br />
Cryptolaemus affinis<br />
USA<br />
Meyerdirk et al. 1979, 1982<br />
Papua New Guinea Szent-Ivany 1963<br />
Cryptolaemus montrouzieri<br />
Easter Is<br />
Ripa et al. 1995<br />
India<br />
Chacko et al. 1978<br />
Mediterranean<br />
Panis 1977<br />
Diomus pumilio<br />
Australia Meyerdirk 1983<br />
Exochomus flavipes<br />
South Africa Samways 1983<br />
Exochomus flaviventris<br />
Mediterranean Kanika et al. 1993<br />
Harmonia octomaculata<br />
Australia Murray 1978<br />
Hyperaspis lateralis<br />
California Bartlett & Lloyd 1958<br />
Hyperaspis polita<br />
Turkey Soylu & Urel 1977<br />
Hyperaspis 2 ´ spp. USSR Niyazov 1969<br />
Bennett & Hughes 1959<br />
Nephus (= Scymnus) bipunctatus<br />
USSR Niyazov 1969<br />
Nephus (= Scymnus)<br />
reunioni<br />
East Africa Ershova & Orlinskii 1982<br />
Pullus pallidicollis<br />
India Prakasan 1987<br />
Scymnus (= Nephus)<br />
includens<br />
Italy Tranfaglia & Viggiani 1973<br />
Scymnus apetzi<br />
USSR Niyazov 1969<br />
Scymnus apiciflavus<br />
Indonesia Kalshoven 1981<br />
Scymnus biguttatus<br />
USSR Niyazov 1969<br />
292 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong>
Table 4.15.1 (contÕd) Predators <strong>of</strong> Planococcus citri<br />
Species<br />
COLEOPTERA<br />
Country Reference<br />
COCCINELLIDAE (contÕd)<br />
Scymnus binaevatus South Africa Smith 1923<br />
Scymnus spp. South Africa<br />
Turkey<br />
Samways 1983<br />
Soylu & Urel 1977<br />
Scymnus roepkei Indonesia Kalshoven 1981<br />
Scymnus sordidus California Bartlett & Lloyd 1958;<br />
Bennett & Hughes 1959<br />
Scymnus subvillosus USSR Niyazov 1969<br />
DIPTERA<br />
CECIDOMYIIDAE<br />
Coccidodiplosis smithi Indonesia Kalshoven 1981<br />
Diadiplosis hirticornis Japan Yukawa 1978<br />
Dicrodiplosis sp. India Chacko et al. 1977<br />
Triommata coccidivora India Prakasan 1987<br />
CHAMAEMYIIDAE<br />
Leucopis alticeps USSR<br />
Italy<br />
Niyazov 1969<br />
Raspi & Bertolini 1993<br />
Leucopis bella California Bartlett & Lloyd 1958<br />
Leucopis silesiaca Italy Raspi & Bertolini 1993<br />
CRYPTOCHETIDAE<br />
Cryptochetum sp. India Chacko et al. 1977<br />
SYRPHIDAE<br />
Syrphus sp. Australia Murray 1982<br />
4.15<br />
Planococcus citri 293
Table 4.15.1 (contÕd) Predators <strong>of</strong> Planococcus citri<br />
Species<br />
LEPIDOPTERA<br />
Country Reference<br />
LYCAENIDAE<br />
Spalgis epius<br />
HYMENOPTERA<br />
ENCYRTIDAE<br />
India Chacko et al. 1977;<br />
Mani & Krishnamoorthy 1990<br />
Achrysophagus sp. Turkey Soylu & Urel 1977<br />
Anagyrus bohemani Spain<br />
India<br />
Carrero 1980a<br />
Varma 1977<br />
Anagyrus greeni Indonesia Kalshoven 1981<br />
Anagyrus pseudococci Italy<br />
Turkey<br />
USA<br />
USSR<br />
Viggiani 1975a<br />
Soylu & Urel 1977<br />
Meyerdirk et al. 1982<br />
Niyazov 1969<br />
Anagyrus sp. nr sawadai USA Meyerdirk et al. 1982<br />
Blepyrus insularis India Chacko et al. 1977<br />
Blepyrus saccharicola California Bennett & Hughes 1959<br />
Chrysoplatycerus splendens USA Bartlett & Lloyd 1958;<br />
Summy et al. 1986<br />
Clausenia josefi Mediterranean Niyazov 1969<br />
Coccidoxenoides (= Pauridia) peregrinus China<br />
India<br />
Italy<br />
USA<br />
Bartlett 1978<br />
Krishnamoorthy & Mani 1989a; Mani 1994<br />
Viggiani & Maresca 1973<br />
Meyerdirk et al. 1978, 1979, 1982<br />
294 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong>
Table 4.15.1 (contÕd) Predators <strong>of</strong> Planococcus citri<br />
Species<br />
HYMENOPTERA<br />
Country Reference<br />
ENCYRTIDAE (contÕd)<br />
Leptomastidea abnormis Spain<br />
Turkey<br />
USA<br />
Leptomastix dactylopii Brazil<br />
India<br />
USA<br />
Carrero 1980a<br />
Soylu & Urel 1977<br />
Meyerdirk et al. 1979, 1982<br />
Mani 1995<br />
Mani 1995<br />
Meyerdirk et al. 1978, 1979, 1982<br />
Leptomastix nigrocoxalis India Prakasan & Kumar 1985<br />
Leptomastix trilongifasciatus India Kalshoven 1981<br />
Ophelosia crawfordi Bartlett & Lloyd 1958<br />
Pseudaphycus angelicus California Bartlett & Lloyd 1958<br />
Pseudaphycus maculipennis USSR Sinadskii & Kozarzhevskaya 1980<br />
Pseudaphycus perdignus Bennett & Hughes 1959<br />
Sympherobius barberi USA Meyerdirk et al. 1982<br />
Timberlakia gilva South Africa Prinsloo 1982<br />
PLATYGASTERIDAE<br />
Allotropa citri China Bartlett & Lloyd 1958<br />
Allotropa kamburovi South Africa Annecke & Prinsloo 1977<br />
Allotropa mecrida USSR Niyazov 1969<br />
4.15 Planococcus citri 295
296 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
The citrus mealybug is also attacked in most regions by encyrtid and<br />
sometimes by platygasterid parasitoids (Table 4.15.2), several <strong>of</strong> which are<br />
capable <strong>of</strong> having a significant effect in warm climates. If the origin <strong>of</strong><br />
P. citri is really China, it is surprising that there are not reports <strong>of</strong> a number<br />
<strong>of</strong> specific or near specific parasitoids from that region. Indeed the species<br />
most commonly employed for biological control is Leptomastix dactylopii<br />
which is believed to be native to Brazil.<br />
A few fungi attack P. citri under humid conditions (Table 4.15.3).<br />
Attempts at biological control<br />
Any account <strong>of</strong> the biological control <strong>of</strong> P. citri is complicated by the facts that<br />
(i) it has been confused with other species on a number <strong>of</strong> occasions, so that<br />
early records are <strong>of</strong>ten unreliable, (ii) documentation <strong>of</strong> some introductions is<br />
poor or lacking, and (iii) natural enemies (that also attack it) have <strong>of</strong>ten been<br />
introduced in programs aimed at other mealybugs. Table 4.15.4 summarises<br />
the main introductions for biological control <strong>of</strong> P. citri.<br />
CALIFORNIA<br />
It is convenient to outline, first, the prolonged attempts against P. citri in<br />
California, since programs elsewhere almost always draw extensively on<br />
experience there. Furthermore, since the first introduction <strong>of</strong> Cryptolaemus<br />
montrouzieri from Australia to California in 1891Ð92, there have been few<br />
parasites or predators used in the control <strong>of</strong> any economically important<br />
mealybug anywhere in the world that have not also been tested against<br />
P. citri in California, in the hope that they might attack it also (Bartlett<br />
1978).<br />
C. montrouzieri was mass reared and released in California against<br />
P. citri with some success, but repeated releases were required to achieve<br />
satisfactory control. Another coccinellid Nephus (= Scymnus) bipunctatus<br />
(under the name <strong>of</strong> Cryptogonus orbiculus) was introduced in 1910 from the<br />
Philippines, but did not become established (Bartlett 1978). However,<br />
establishment followed the introduction from Sicily in 1914 <strong>of</strong> the parasitoid<br />
Leptomastidea abnormis, although control was only partly successful<br />
(Viereck 1915; Smith 1917). The coccinellid Scymnus binaevatus from<br />
South Africa was established in 1921, but persists only in small numbers.<br />
Unsuccessful attempts were made to establish the encyrtid<br />
Coccidoxenoides peregrinus from Hawaii where it was having a major<br />
impact on P. citri, misidentified at the time as Planococcus kraunhiae.<br />
However, progeny <strong>of</strong> a single female from South China in 1950 allowed the<br />
species to become established, although at a low level (Flanders 1957).
Table 4.15.2 Parasitoids <strong>of</strong> Planococcus citri<br />
Species<br />
HYMENOPTERA<br />
ENCYRTIDAE<br />
Country Reference<br />
Achrysophagus sp. Turkey Soylu & Urel 1977<br />
Anagyrus bohemani Spain<br />
India<br />
Carrero 1980a<br />
Varma 1977<br />
Anagyrus greeni Indonesia Kalshoven 1981<br />
Anagyrus pseudococci Italy<br />
Turkey<br />
USA<br />
USSR<br />
Viggiani 1975a<br />
Soylu & Urel 1977<br />
Meyerdirk et al. 1982<br />
Niyazov 1969<br />
Anagyrus sp. nr sawadai USA Meyerdirk et al. 1982<br />
Blepyrus insularis India Chacko et al. 1977<br />
Blepyrus saccharicola California Bennett & Hughes 1959<br />
Chrysoplatycerus splendens USA Bartlett & Lloyd 1958;<br />
Summy et al. 1986<br />
Clausenia josefi Mediterranean Niyazov 1969<br />
Coccidoxenoides (= Pauridia) peregrinus China<br />
India<br />
Italy<br />
USA<br />
Leptomastidea abnormis Spain<br />
Turkey<br />
USA<br />
Leptomastix dactylopii Brazil<br />
India<br />
USA<br />
Bartlett 1978<br />
Krishnamoorthy & Mani 1989a; Mani 1994<br />
Viggiani & Maresca 1973<br />
Meyerdirk et al. 1978, 1979, 1982<br />
Carrero 1980a<br />
Soylu & Urel 1977<br />
Meyerdirk et al. 1979, 1982<br />
Mani 1995<br />
Mani 1995<br />
Meyerdirk et al. 1978, 1979, 1982<br />
Leptomastix nigrocoxalis India Prakasan & Kumar 1985<br />
4.15 Planococcus citri 297
Table 4.15.2 (contÕd) Parasitoids <strong>of</strong> Planococcus citri<br />
Species<br />
HYMENOPTERA<br />
Country Reference<br />
ENCYRTIDAE (contÕd)<br />
Leptomastix trilongifasciatus India Kalshoven 1981<br />
Ophelosia crawfordi Bartlett & Lloyd 1958<br />
Pseudaphycus angelicus California Bartlett & Lloyd 1958<br />
Pseudaphycus maculipennis USSR Sinadskii & Kozarzhevskaya 1980<br />
Pseudaphycus perdignus Bennett & Hughes 1959<br />
Sympherobius barberi USA Meyerdirk et al. 1982<br />
Timberlakia gilva South Africa Prinsloo 1982<br />
PLATYGASTERIDAE<br />
Allotropa citri China Bartlett & Lloyd 1958<br />
Allotropa kamburovi South Africa Annecke & Prinsloo 1977<br />
Allotropa mecrida USSR Niyazov 1969<br />
Table 4.15.3 Pathogens attacking Planococcus citri<br />
Species Country Reference<br />
Aspergillus flavus Cuba Martinez & Bravo 1989<br />
Cladosporium oxysporum South Africa Samways 1983; Samways & Grech 1986<br />
Entomophthora fresenii Indonesia Kalshoven 1981<br />
Entomophthora fumosa Australia Murray 1978; Samal et al. 1978<br />
Fusarium sp. Easter Is Ripa et al. 1995<br />
298 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong>
4.15 Planococcus citri 299<br />
The encyrtid Leptomastix dactylopii, introduced in 1934 from Brazil,<br />
was mass reared and released. Recoveries were made over a number <strong>of</strong> years<br />
following each spring or summer release, but not following the ensuing<br />
winter. Eventually, a few managed to overwinter, resulting in a low level<br />
population (Compere 1939).<br />
The South China platygasterid, Allotropa citri, introduced in 1950,<br />
attacked 1st and 2nd instar P. citri, but mass releases over some 6 years<br />
resulted in few field recoveries and it is not thought to be established<br />
(Bartlett 1978).<br />
A somewhat polyphagous encyrtid, Anagyrus pseudococci, was<br />
unsuccessfully introduced from Brazil in 1934 and 1953. The same or a<br />
similar species, Anagyrus sp. nr pseudococci, was brought in from Italy in<br />
1955, but was established only briefly (Bartlett 1978).<br />
The coccinellid Exochomus metallicus from Eritrea was established<br />
from introductions in 1954 against citricola scale (Coccus pseudomagnoliarum)<br />
and black scales and attacks P. citri on plants other than citrus<br />
(Bartlett 1978).<br />
In spite <strong>of</strong> this series <strong>of</strong> introductions, the natural enemies <strong>of</strong> P. citri in<br />
California do not, unaided, maintain populations at sub-economic levels,<br />
mainly because, it is claimed, climatic conditions permit the overwintering<br />
<strong>of</strong>, at best, inadequate populations. Cryptolaemus montrouzieri has <strong>of</strong>ten<br />
provided spectacular control <strong>of</strong> high populations, but is unable to maintain<br />
its numbers on low prey populations and disappears, requiring<br />
reintroduction. Methods are available for its low cost mass production<br />
(Fisher 1963).<br />
The encyrtid Leptomastidea abnormis is <strong>of</strong> considerable value in<br />
attacking young mealybugs sheltering under citrus fruit sepals in spring and<br />
autumn, but is adversely affected by high temperatures. Both Leptomastix<br />
dactylopii and Coccidoxenoides peregrinus build up high numbers<br />
following mass releases, but crash over winter and require repeated mass<br />
releases to maintain effectiveness.<br />
AUSTRALIA<br />
P. citri can be an important pest <strong>of</strong> citrus, but also attacks passionfruit and<br />
custard apple in warmer regions. The earliest attempts at biological control<br />
were in Western Australia, where Cryptolaemus montrouzieri was<br />
introduced from New South Wales in 1902 and an unidentified coccinellid<br />
from Spain in 1903. Only the former was established and it rapidly became<br />
an important factor in the successful control <strong>of</strong> mealybugs there (Wilson<br />
1960).
300 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Before biological control was attempted in Queensland in 1980, six<br />
natural enemies were recorded, the coccinellids Cryptolaemus montrouzieri,<br />
and Harmonia octomaculata (= Coccinella arcuata) the chrysopids<br />
Chrysopa sp. and Oligochrysa lutea (all 4 native) and the exotic encyrtids<br />
Leptomastidea abnormis and Coccidoxenoides peregrinus. However, they<br />
were unable to maintain infestations consistently at acceptable commercial<br />
levels. Attack by a fungus similar to Entomophthora fumosa caused up to<br />
58.1% mortality <strong>of</strong> 3rd instar nymphs and adults on passionfruit during wet<br />
periods (Murray 1978, 1982; Smith et al. 1988).<br />
The Brazilian encyrtid Leptomastix dactylopii was imported from<br />
California and approximately 2.5 million adults released between 1980 and<br />
1987. It established readily and became the commonest natural enemy <strong>of</strong><br />
P. citri throughout southeast Queensland, reducing mealybug infestations,<br />
averaging 38% <strong>of</strong> fruit in early December, to an acceptable level <strong>of</strong> 5% or<br />
less at harvest in April. Parasitoid numbers were lowest during winter and<br />
spring and augmentative releases <strong>of</strong> 5 to 10 thousand parasitoids per hectare<br />
in spring to early summer advanced parasitoid activity by 6 weeks.<br />
When no releases were made, the parasitoid was first recorded in early<br />
February and was present in an average <strong>of</strong> 55% <strong>of</strong> mealybug-infested fruit<br />
by mid-March. The mealybug infestation peaked at an average <strong>of</strong> 47% in<br />
mid-December but, by late April, only dropped to 10% and the presence <strong>of</strong><br />
the mealybug on 25% or more fruit from December to March usually<br />
resulted in excessive amounts <strong>of</strong> sooty mould. C. montrouzieri was recorded<br />
on a maximum <strong>of</strong> 5% <strong>of</strong> mealybug-infested fruit, and augmentative release<br />
failed to increase this level. Augmentative release <strong>of</strong> L. dactylopii was found<br />
to be at least as cheap as pesticides and far more compatible with IPM <strong>of</strong><br />
other citrus pests (Smith et al. 1988).<br />
Placing sticky bands around the trunks <strong>of</strong> custard apple trees reduced the<br />
numbers <strong>of</strong> the ant, Pheidole megacephala, and lowered, somewhat, the<br />
numbers <strong>of</strong> P. citri. Parasitisation <strong>of</strong> P. citri by Leptomastidea abnormis was<br />
low and unaffected by banding, but there were more predators (especially<br />
Oligochrysa lutea, Cryptolaemus montrouzieri and Syrphus sp.).<br />
Nevertheless, natural enemies were still unable to maintain P. citri at<br />
acceptable levels (Murray 1982).<br />
BERMUDA<br />
Seven species <strong>of</strong> parasitoid and four predators were introduced between<br />
1951 and 1955, mainly from California, but originating elsewhere:<br />
Coccidoxenoides peregrinus from Hawaii and south China, Leptomastix<br />
dactylopii (South American race) Leptomastidea abnormis, Pseudaphycus<br />
perdignus, Anagyrus pseudococci, Blephyrus saccharicola and Allotropa<br />
citri. Only C. peregrinus, L. dactylopii and L. abnormis were established
BRAZIL<br />
CHILE<br />
CHINA<br />
CUBA<br />
CYPRUS<br />
FRANCE<br />
4.15 Planococcus citri 301<br />
and, <strong>of</strong> these, L. dactylopii may have been the local form. Of the predators,<br />
Cryptolaemus montrouzieri was established, but only briefly and two<br />
species <strong>of</strong> Hyperaspis and Scymnus sordidus failed to breed on P. citri<br />
(Simmonds 1957; Bennett and Hughes 1959).<br />
P. citri is common on a range <strong>of</strong> plants, but citrus is seldom infested. In<br />
1939 Leptomastidea abnormis and Leptomastix dactylopii were<br />
commonly reared from it, and it was attacked by numerous predators,<br />
including Hyperaspis c-nigrum, Nephus sp., Diomus sp., lacewings and<br />
cecidomyiids. Anagyrus pseudococci was not recorded, although it was<br />
present in Argentina (Compere 1939).<br />
Cryptolaemus montrouzieri was introduced in 1931, 1933 and 1939, but<br />
establishment is not recorded, although it is present on Easter Is (Ripa et al.<br />
1995). Extensive releases <strong>of</strong> two parasitoids from California resulted in<br />
establishment: Leptomastidea abnormis during 1931Ð36 and Leptomastix<br />
dactylopii in 1936 and 1938: both <strong>of</strong> these and Coccidoxenoides peregrinus<br />
were also established on Easter Is (Ripa et al. 1995). Attempts failed in 1954<br />
to establish from California: Allotropa citri, Anagyrus pseudococci,<br />
Coccidoxenoides peregrinus and Pseudaphycus perdignus (Duran 1944;<br />
Gonzalez and Rojas 1966).<br />
Four species <strong>of</strong> parasitic wasp were introduced from China to California in<br />
1950: a uniparental species <strong>of</strong> Coccidoxenoides and a biparental species <strong>of</strong><br />
Allotropa from Guangzhou, a biparental species <strong>of</strong> Pseudaphycus from<br />
Taiwan and a biparental species <strong>of</strong> Coccophagus from Hong Kong (Flanders<br />
1951).<br />
Seven natural enemies attack P. citri on c<strong>of</strong>fee including a Leptomastix sp..<br />
A cecidomyiid predator was commonest (Martinez et al. 1992). A fungus<br />
Aspergillus flavus was also detected (Martinez and Bravo 1989).<br />
Leptomastix dactylopii was introduced from Italy in 1977 and became<br />
established, attaining a parasitisation rate <strong>of</strong> 15% in 1979. At harvest there<br />
were far fewer P. citrus on the fruit than in a plot that had received 3<br />
applications <strong>of</strong> insecticide (Krambias and Kontzonis 1980).<br />
Cryptolaemus montrouzieri was introduced from California in 1918 and<br />
became established, but overwinter survival was low (Marchal 1921, 1922;<br />
Poutiers 1922; Marchal and Pussard 1938). Of the 6 species <strong>of</strong> mealybug on<br />
citrus in the Mediterranean basin, P. citri is the most injurious. Damage by it<br />
in France is such that an economic injury level <strong>of</strong> 2% infested fruit has been
302 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
GREECE<br />
INDIA<br />
set. Mass rearing and release <strong>of</strong> Leptomastix dactylopii, which requires the<br />
use <strong>of</strong> fewer mealybugs for laboratory rearing than the coccinellid<br />
C. montruzieri and is cheaper, is preferred to that <strong>of</strong> the coccinellid if only<br />
one natural enemy is to be used. However, it is preferable to employ both,<br />
with C. moutrouzieri destroying high populations <strong>of</strong> females and eggs and<br />
L. dactylopii parasitising nymphs, even if populations are scattered (Panis<br />
1977, 1979).<br />
Cryptolaemus montrouzieri was liberated against P. citri on potted orange<br />
trees in a glasshouse at 25 to 30¡C and 55 to 70% RH and the results compared<br />
with the release <strong>of</strong> Nephus (= Scymnus) reunioni or the application <strong>of</strong><br />
insecticide. C. montrouzieri significantly reduced mealybug populations<br />
and was as effective as treatment with methidathion (Hamid and Michelakis<br />
1994).<br />
Leptomastix dactylopii was introduced in 1983 to Bangalore and rapidly<br />
became established on P. citri on mandarins and c<strong>of</strong>fee, causing up to 100%<br />
parasitisation (Nagarkatti et al. 1992). Seven years later in 1991, P. citri was<br />
being attacked on lemon and lime by L. dactylopii and the more abundant<br />
indigenous Coccidoxenoides peregrinus, which was causing 10 to 30%<br />
parasitisation. C. peregrinus attacks preferentially the early and Leptomastix<br />
dactylopii the later instars (Krishnamoorthy and Mani 1989a; Mani 1994).<br />
In 1984 L. dactylopii was released in a lime orchard in Karnataka. Prior to<br />
release, infestation by P. citri ranged from 38 to 65%, but establishment <strong>of</strong><br />
the parasitoid led to excellent control within 4 months and no insecticides<br />
were required in following seasons. The parasitoid was shown to have<br />
migrated about 0.5 km in a 2-year period and a mean <strong>of</strong> 2.3 adult parasitoids<br />
were recovered from each infested fruit in an orange orchard<br />
(Krishnamoorthy 1990). Parasitisation <strong>of</strong> P. citri and other mealybugs on<br />
c<strong>of</strong>fee ranged, in different years, between 19 and 45% (Reddy et al. 1988)<br />
and 0 and 85% (Reddy et al. 1992). Although L. dactylopii greatly reduced<br />
the population <strong>of</strong> P. citri, augmentative releases were required for<br />
continuing control in a c<strong>of</strong>fee plantation (Reddy and Bhat 1993).<br />
Native predators <strong>of</strong> P. citri on c<strong>of</strong>fee include the lycaenid, Spalgis epius,<br />
the coccinellid, Pullus pallidicollis, and the cecidomyiid Triommata<br />
coccidivora (Prakasan 1987). On citrus the chrysopids Mallada boninensis,<br />
Odontochrysa (= Chrysopa) lacciperda, Mallada basalis and Chrysoperla<br />
carnea were found (Krishnamoorthy and Mani 1989b); and on guava<br />
Odontochrysa lacciperda, Spalgis epius and Cryptolaemus montrouzieri<br />
(Mani and Krishnamoorthy 1990).
ISRAEL<br />
4.15 Planococcus citri 303<br />
Releases <strong>of</strong> Cryptolaemus montrouzieri on a c<strong>of</strong>fee estate in Kerala<br />
virtually eliminated P. citri, but the coccinellid could not be found thereafter<br />
for some 6 months, when it reappeared about 10 km distant and virtually<br />
eliminated mealybugs from an infestation (Chacko et al. 1978). The<br />
requirement that each female consume at least 8 P. citri for normal egg<br />
production (192 eggs/female) was suggested as the reason for the poor<br />
establishment <strong>of</strong> C. montrouzieri when only low populations <strong>of</strong> P. citri are<br />
available (Reddy et al. 1991).<br />
The citrus mealybug is a serious pest <strong>of</strong> citrus and ornamentals <strong>of</strong> tropical<br />
origin. It develops on the fruit and roots <strong>of</strong> young trees, the main damage<br />
being caused by individuals settling beneath the sepals <strong>of</strong> citrus fruit and<br />
injuring the fruit: honeydew produced also attracts fruit-piercing moths<br />
(Mendel et al. 1992).<br />
Two native encyrtids, Anagyrus pseudococci and Leptomastidea<br />
abnormis, attack P. citri which is, nevertheless, an important pest <strong>of</strong> citrus<br />
and some other crops (Rosen and Ršssler 1966).<br />
Unsuccessful attempts were made to establish Cryptolaemus<br />
montrouzieri in 1924 from Egypt (Bodenheimer and Guttfeld 1929; Mason<br />
1941) and 1958 (Rosen 1967a), but a Spanish strain introduced in the 1980s<br />
finally established (Mendel et al. 1992). Leptomastix dactylopii imported<br />
from Canada in 1941 was only briefly established (Rivnay 1960). Clausenia<br />
purpurea, a parasitoid <strong>of</strong> Pseudococcus citriculus introduced from Japan in<br />
1940, attacks P. citri, but has little effect on its density (Rosen 1964).<br />
P. citrus became a serious pest on recently established grapefruit in<br />
southern Negev. The encyrtid Anagyrus pseudococci was present in very<br />
low numbers and Chrysoperla carnea was active, but only in spring. The<br />
coccinellid Chilocorus bipustulatus was more abundant and was considered<br />
to have potential as a biological control agent (Berlinger et al. 1979). Seven<br />
parasitoids and 10 predators have been recorded, <strong>of</strong> which Anagyrus<br />
pseudococci (Encyrtidae) and Sympherobius sanctus (Chrysopidae) were<br />
found in significant numbers by Mendel et al. (1992). However,<br />
development <strong>of</strong> A. pseudococci is restricted by winter temperatures. Its<br />
development threshold is 13¡C and no eggs are laid below 15¡C, whereas<br />
P. citri still lays eggs at 13¡C and its development threshold is 8.4¡C. By the<br />
time in late spring when the population <strong>of</strong> A. pseudococci starts to built up,<br />
P. citri has already settled under the sepals, where it is well protected from<br />
the parasitoid. Climatic conditions were thus regarded as unfavourable for<br />
existing natural enemies (Mendel et al. 1992).
304 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
ITALY<br />
Infestations <strong>of</strong> P. citri develop on citrus in Sicily, Procida, Sardinia and parts<br />
<strong>of</strong> mainland Italy, especially where it is protected by ants from its native<br />
parasitoids, Leptomastidea abnormis and Anagyrus pseudococci (Zinna<br />
1960). Cryptolaemus montrouzieri has been imported a number <strong>of</strong> times<br />
since 1908 and has become established in some warmer areas, but is so<br />
reduced in numbers during winter that satisfactory control is not obtained<br />
without supplementations (Constantino 1935; Liotta 1965; Liotta and Mineo<br />
1965). Coccidoxenoides peregrinus and Leptomastix dactylopii were<br />
introduced to Procida in 1956Ð57, but did not survive the winter (Bartlett<br />
1978). L. dactylopii was reintroduced in 1974 to this island, to the mainland<br />
(Campania, Calabria), and to Sardinia and Sicily. At almost all 15 release<br />
sites it afforded, initially, a high level <strong>of</strong> parasitisation (Viggiani 1975a,b).<br />
Small overwintering populations, reduced further by hyperparasitoids and<br />
fungus diseases, persisted in some areas, but required supplementation for<br />
control (Mineo and Viggiani 1975a,b). L. dactylopii was again released in<br />
1979 on Sicily and gave good results (5% <strong>of</strong> fruit infested), but it is unclear<br />
whether or not it is able to overwinter (Barbagallo et al. 1982; Longo and<br />
Benfatto 1982). Release <strong>of</strong> L. dactylopii in a mainland orange orchard in<br />
Calabria led to a reduction in infested fruit from 80.9% to 12% and only<br />
7.3% <strong>of</strong> the fruit was unmarketable (Luppino 1979). It now seems that<br />
L. dactylopii and Cryptolaemus montrouzieri are mass reared and released<br />
each year against P. citri (Raciti et al. 1995).<br />
<strong>Insect</strong>icides are seldom required against P. citri in Sardinia, where<br />
L. dactylopii may cause 96% cumulative parasitisation and where<br />
Cryptolaemus montrouzieri and Nephus (= Scymnus) reunioni may increase<br />
to more than 100 individuals per tree (Ortu 1982; Ortu and Prota 1983).<br />
PAPUA NEW GUINEA<br />
Mealybugs, including P. citri, were causing up to 75% reduction <strong>of</strong> c<strong>of</strong>fee<br />
yield in the highlands near Wau in the mid fifties. Introduction <strong>of</strong><br />
Cryptolaemus affinis in 1957 from the lower Markham Valley resulted in its<br />
rapid spread and a substantial reduction <strong>of</strong> the infestations within one season<br />
(Szent-Ivany 1963).<br />
PERU<br />
Very good control <strong>of</strong> P. citri is given in some areas by the encyrtid<br />
Coccidoxenoides peregrinus, which is restricted to the citrus mealybug and<br />
appears to have arrived accidentally in 1963 (Salazar 1972).<br />
SOUTH AFRICA<br />
The history <strong>of</strong> introductions to South Africa against P. citri is confused<br />
because <strong>of</strong> misidentifications <strong>of</strong> other mealybugs for this species. However,<br />
Bedford (1976) reports that P. citri is under biological control.
4.15 Planococcus citri 305<br />
When the ant Anoplolepis custodiens was excluded from guava trees<br />
bearing P. citri at Nelspruit, the population <strong>of</strong> both ants and mealybugs<br />
dropped to half. Without ants, the mealybugs were heavily preyed upon by<br />
the ant-intolerant Exochomus flavipes and the ant-tolerant Scymnus spp.<br />
Later, the mealybugs were almost eliminated by the fungus Cladosporium<br />
sp. nr oxysporum (Samways 1983).<br />
SPAIN<br />
P. citri is parasitised by two indigenous encyrtids, Leptomastidea abnormis<br />
and Anagyrus bohemani, but at a low level (Carrero 1980a). Cryptolaemus<br />
montrouzieri was introduced and established before 1928 and produces<br />
control in the warmer months (Gomez 1951; Carrero 1980b). Leptomastix<br />
dactylopii was introduced in 1948 from California (Gomez 1951) and in<br />
1977 from Italy, but did not become established (Carrero 1980b).<br />
USA (OTHER THAN CALIFORNIA)<br />
Florida<br />
Cryptolaemus montrouzieri was introduced to Florida in 1930 for the control<br />
<strong>of</strong> P. citri on citrus and bulbs. It became established but failed to overwinter<br />
in sufficient numbers to achieve adequate control (Bartlett 1978; Muma<br />
1954, Watson 1932).<br />
Hawaii<br />
P. citri (originally misidentified as P. kraunhiae) was the target <strong>of</strong> many<br />
introductions (Swezey 1931), although not as severe a pest as on the<br />
mainland. Cryptolaemus montrouzieri was introduced from Australia in<br />
1894, Leptomastidea abnormis from California in 1915 and Leptomastix<br />
dactylopii also from California in 1946. All became established.<br />
Texas<br />
In 1970, Coccidoxenoides peregrinus was the dominant parasitoid <strong>of</strong> 3<br />
present on P. citri attacking grapefruit and Sympherobius barberi the<br />
commonest predator (Dean et al. 1971). In 1977 Leptomastix dactylopii,<br />
which had been introduced from California in 1970, and Anagyrus sp. nr<br />
sawadai were found for the first time on P. citri in Texas. L. dactylopii was<br />
the most abundant parasite in mid-August (parasitising 20.7% <strong>of</strong> P. citri),<br />
the reputedly indigenous Coccidoxenoides peregrinus (with 48.5%) the<br />
most abundant in late August, whereas Anagyrus sp. (with 4.3%) was the<br />
only parasitoid recovered in mid-September. A hyperparasitoid<br />
Prochiloneurus dactylopii attacked 1% <strong>of</strong> the primary parasitoids in mid-<br />
August (Meyerdirk et al. 1978). Release <strong>of</strong> 5 encyrtid parasitoids on<br />
glasshouse citrus resulted in the rapid suppression <strong>of</strong> P. citri. Leptomastidea<br />
abnormis, Anagyrus pseudococci and Leptomastix dactylopii persisted for<br />
periods between 24 and 32 weeks and maintained the host at low densities,<br />
whereas Chrysoplatycerus splendens and Coccidoxenoides peregrinus
306 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
USSR<br />
persisted for only 20 weeks (Summy et al. 1986). The coccinellid Diomus<br />
pumilio, whose biology is described, was introduced from South Australia<br />
and is a potentially valuable predator (Meyerdirk 1983).<br />
P. citri can be a serious pest <strong>of</strong> grape vines, citrus, fig and pomegranate.<br />
Anagyrus pseudococci from Surkham Dalya and Leptomastix dactylopii and<br />
Leptomastidea abnormis from California were introduced to Uzbekistan<br />
commencing in 1959 and resulted in establishment (Roxanova and Loseva<br />
1963). Anagyrus pseudococci destroys up to 75% <strong>of</strong> P. citri in areas not<br />
treated with insecticides in the south <strong>of</strong> European Russia and in Soviet<br />
Central Asia. The next most important parasitoid, Allotropa mecrida<br />
attacked up to 20% in Turkmenia in 1967 and in Georgia. In 1960,<br />
Leptomastidea abnormis and Leptomastix dactylopii were introduced from<br />
USA into Georgia and Turkmenia. In Transcaucasia and Soviet Central Asia<br />
the hyperparasitoid Thysanus subaeneus attacks 18 to 20% <strong>of</strong> Allotropa<br />
mecrida. Other hyperparasitoids are Pachyneuron solitarius and<br />
Neoprochiloneurus bolivari.<br />
One <strong>of</strong> the most effective predators <strong>of</strong> P. citri is Cryptolaemus<br />
montrouzieri, introduced from Egypt in 1932 to the Black Sea area. Others<br />
are Coccinella septempunctata, Hyperaspis polita, Nephus bipunctatus,<br />
Scymnus apetzi, S. subvillosus, and S. biguttatus which were recorded in<br />
Turkmenia. The larvae <strong>of</strong> the fly Leucopis alticeps and <strong>of</strong> the lacewing<br />
Chrysoperla carnea are able to devastate all stages <strong>of</strong> P. citri. The<br />
coccinellids were parasitised by Homalotylus sp. and the lacewing by<br />
Telenomus acrobates (Niyazov 1969).<br />
The coccinellid Nephus reunioni was introduced into southern areas in<br />
1978 and has reduced P. citri on grape vines. It is capable <strong>of</strong> overwintering,<br />
but with high mortality, and is more tolerant <strong>of</strong> moisture conditions than<br />
Cryptolaemus montrouzieri (Orlinskii et al. 1989).
Table 4.15.4 Introductions for the biological control <strong>of</strong> Planococcus citri<br />
Species<br />
HYMENOPTERA<br />
ENCYRTIDAE<br />
From To Year Result Reference<br />
Anagyrus kivuensis Kenya California 1948 Ð Bartlett & Lloyd 1958<br />
Anagyrus pseudococci Brazil<br />
California<br />
California<br />
California<br />
Bermuda<br />
Chile<br />
1934<br />
1953<br />
1951Ð54<br />
1954<br />
Ð<br />
+<br />
Ð<br />
Ð<br />
Bartlett & Lloyd 1958<br />
Bartlett & Lloyd 1958<br />
Bennett & Hughes 1959<br />
Gonzalez & Rojas 1966<br />
Anagyrus sp. nr pseudococci Italy California 1955, 1965 Ð Bartlett 1978,<br />
Bartlett & Lloyd 1958<br />
Anagyrus sp. nr sawadai Texas + Meyerdirk et al. 1978<br />
Blepyrus saccharicola California Bermuda 1951Ð54 Ð Bennett & Hughes 1959<br />
Coccidoxenoides peregrinus Hawaii California Ð Armitage 1920<br />
China California + Flanders 1951<br />
China Bermuda 1951Ð54 + Bennett & Hughes 1959<br />
California Chile 1954 Ð Gonzalez & Rojas 1966<br />
Peru 1963 + Salazar 1972<br />
Leptomastidea abnormis Sicily<br />
California<br />
California<br />
USA<br />
California<br />
Chile<br />
Bermuda<br />
USSR<br />
1914<br />
1931<br />
1951Ð54<br />
1960<br />
+<br />
+<br />
+<br />
Viereck 1915; Smith 1917<br />
Gonzalez & Rojas 1966<br />
Bennett & Hughes 1959<br />
Niyazov 1969<br />
Leptomastidea sp. nr abnormis Mexico California 1956 Ð Bartlett & Lloyd 1958<br />
4.15 Planococcus citri 307
Table 4.15.4 (contÕd) Introductions for the biological control <strong>of</strong> Planococcus citri<br />
Species<br />
HYMENOPTERA<br />
From To Year Result Reference<br />
ENCYRTIDAE (contÕd)<br />
Leptomastix dactylopii Brazil<br />
California<br />
California<br />
California<br />
California<br />
Italy<br />
California<br />
Chile<br />
Chile<br />
Bermuda<br />
Turkey<br />
Texas<br />
Australia<br />
India<br />
India<br />
Israel<br />
Italy<br />
Cyprus<br />
Sardinia<br />
1934<br />
1936<br />
1958<br />
1951Ð54<br />
1970<br />
1980<br />
1983Ð85<br />
1984<br />
1956Ð7<br />
1977<br />
1974<br />
+<br />
Ð<br />
+<br />
?<br />
+<br />
+<br />
+<br />
+<br />
+<br />
Ð<br />
Ð (early)<br />
+ (later)<br />
+<br />
Ð<br />
Compere 1939<br />
Gonzalez & Rojas 1966<br />
Gonzalez & Rojas 1966<br />
Bennett & Hughes 1959<br />
Tuncyurek 1970<br />
Meyerdirk et al. 1978<br />
Smith et al. 1988<br />
Nagarkatti et al. 1992<br />
Krishnamoorthy 1990,<br />
Prakasan & Bhat 1985;<br />
Krishnamoorthy & Singh 1987;<br />
Prakasan 1987; Reddy et al. 1988;<br />
Mani 1994<br />
Rivnay 1960<br />
Luppino 1979; Longo & Benfatto<br />
1982<br />
Viggiani 1975a,b; Mineo &<br />
Viggiani 1976a<br />
Krambias & Kontzonis 1980<br />
Viggiani 1975a,b; Delrio et al.<br />
1981; Ortu & Prota 1981;<br />
Ortu 1982<br />
Sicily 1979 Ð Barbagallo et al. 1982<br />
Longo & Benfatto 1982<br />
Viggiani 1975a,b<br />
Italy Spain 1977 Ð Carrero 1980b<br />
USA USSR 1960 Niyazov 1969<br />
308 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong>
Table 4.15.4 (contÕd) Introductions for the biological control <strong>of</strong> Planococcus citri<br />
Species<br />
HYMENOPTERA<br />
From To Year Result Reference<br />
ENCYRTIDAE (contÕd)<br />
Pseudaphycus perdignus Eritrea California 1953 Ð Bartlett & Lloyd 1958<br />
California Bermuda 1951Ð54 Ð Bennett & Hughes 1959<br />
Tropidophryne melvillei Kenya California 1948 Ð Bartlett & Lloyd 1958<br />
HYMENOPTERA<br />
PLATYGASTERIDAE<br />
Allotropa citri China California ? Flanders 1951; Bartlett & Lloyd<br />
1958<br />
California Bermuda Ð Bennett & Hughes 1959<br />
California Chile 1954 Ð Gonzalez & Rojas 1966<br />
COLEOPTERA<br />
COCCINELLIDAE<br />
Brumus suturalis India California 1955 Ð Bartlett & Lloyd 1958<br />
Chilocorus angolensis Kenya California 1948 Ð Bartlett & Lloyd 1958<br />
Cryptolaemus montrouzieri Greece Ð Argyriou 1970<br />
Sardinia
Table 4.15.4 (contÕd) Introductions for the biological control <strong>of</strong> Planococcus citri<br />
Species<br />
COLEOPTERA<br />
From To Year Result Reference<br />
COCCINELLIDAE (contÕd)<br />
California Bermuda 1955 + Bennett & Hughes 1959<br />
California Chile<br />
India<br />
Indonesia<br />
1931<br />
1933<br />
1934<br />
Ð<br />
Ð<br />
?<br />
+<br />
Ð<br />
Gonzalez & Rojas 1966<br />
Gonzalez & Rojas 1966<br />
Gonzalez & Rojas 1966<br />
Prakasan 1987<br />
Kalshoven 1981<br />
Spain Israel 1980s + Mendel et al. 1992<br />
Diomus pumilio (= D. flavifrons) Australia Texas + Meyerdirk 1983<br />
Exochomus flavipes Kenya California 1948 Ð Bartlett & Lloyd 1958<br />
Exochomus metallicus Eritrea California 1954 + Bartlett 1978<br />
Hyperaspis jucunda Trinidad California 1955 Ð Bartlett & Lloyd 1958<br />
Hyperaspis sp. nr globula Mexico California 1954 Ð Bartlett & Lloyd 1958<br />
Hyperaspis sp. Eritrea California 1953 Ð Bartlett & Lloyd 1958<br />
Hyperaspis 2 ´ spp. California Bermuda 1958 Ð Bennett & Hughes 1959<br />
Nephus (=Scymnus) bipunctatus Philippines California 1910 Ð Bartlett 1978<br />
Nephus (=Scymnus) reunioni East Africa USSR 1978 + Ershova & Orlinskii 1982; Orlinskii<br />
et al. 1989<br />
Nephus sp. Trinidad California 1958 Ð Bartlett & Lloyd 1958<br />
Platynaspis (?) sp. Eritrea California 1953 Ð Bartlett & Lloyd 1958<br />
Scymnus binaevatus South Africa California + Smith 1923<br />
Scymnus quadrivittatus Kenya California 1948 Ð Bartlett & Lloyd 1958<br />
Scymnus sordidus California Bermuda 1955 Ð Bennett & Hughes 1959<br />
310 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong>
Biology <strong>of</strong> important natural enemies<br />
4.15 Planococcus citri 311<br />
Anagyrus pseudococci Hym.: Encyrtidae<br />
This is a solitary endoparasite <strong>of</strong> 2nd, 3rd and 4th instar mealybugs, but<br />
prefers 3rd instar and egg-laying females. It is believed to be native to the<br />
Mediterranean. A. pseudococci has been known since 1913 in Italy as a<br />
widespread parasitoid <strong>of</strong> P. citri and, in Israel, attacking both P. citri and<br />
Pseudococcus citriculus (Rivnay 1960; Rosen 1964). It is common on<br />
P. citri in Argentina (Compere 1939). In the laboratory it develops<br />
successfully on Pseudococcus fragilis, P. longispinus and P. obscurus.<br />
Females lay about 45 eggs at the rate <strong>of</strong> 3 to 4 per day. In the laboratory<br />
these hatch in 44 hours at 27¡C and the life cycle takes 17 to 18 days (Bartlett<br />
1978) or 15.5 days at 25.6¡C and 60% RH for a Californian strain, which<br />
also had a life span <strong>of</strong> 8.2 days for virgin and 6.9 days for mated females.<br />
Virgin females produce males. A. pseudococci is most active in the spring<br />
and autumn (Domenichini 1952; Avidov et al. 1967; Rivnay 1968; Chandler<br />
et al. 1980).<br />
Progeny production increased and longevity decreased with increase in<br />
temperature between 18¡ and 30¡C (Tingle and Copland 1989). Most<br />
progeny are produced between 27¡ and 30¡C and the threshold for<br />
development is 13.06¡C for males and 12.57¡C for females (Islam and Jahan<br />
1993). In the laboratory maximum egg production was achieved when 50%<br />
honey solution was provided (Islam and Jahan 1992, 1993). Oviposition<br />
behaviour is described by Islam (1992). About 40% <strong>of</strong> parasitoid eggs laid in<br />
P. citri may be lost due to encapsulation (Blumberg et al. 1995). In<br />
Argentina, larvae <strong>of</strong> A. pseudococci are attacked by the hyperparasitoid<br />
Coccophagus heteropneusticus (Compere 1939).<br />
Coccidoxenoides peregrinus Hym.: Encyrtidae<br />
This parasitoid, earlier widely known as Pauridia peregrina, is probably<br />
native to southern China (Bartlett 1978), although Meyerdirk et al. (1978)<br />
suggest that it is indigenous to Texas. It has been reported, inter alia, from<br />
India, Japan, Philippines, Fiji, Hawaii and Uganda. It is a solitary<br />
endoparasitoid <strong>of</strong> 1st, 2nd and 3rd instar female Planococcus citri and<br />
P. kenyae and 1st and 2nd instar males. It is normally parthenogenetic, but<br />
there are rare males. In Uganda it parasitises P. kenyae (Armitage 1920;<br />
Essig 1931) and, in Peru P. citri (Beingolea 1969). In India it completed its<br />
development only in P. citri, although it attacked other mealybugs<br />
(Krishnamoorthy and Mani 1989a).<br />
Females commence oviposition shortly after emergence, and continue<br />
for about 2 days. At 27¡C larval development takes 11 to 12 days and the<br />
pupal stage 16 to 18 days (Zinna 1960; Fisher 1963). However, in India,
312 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
development took 23 to 27 days and adults survived 4 to 9 days at 28 ± 2¡C<br />
(Krishnamoorthy and Mani 1989a). In 1991 in Karnataka, C. peregrinus was<br />
more abundant than Leptomastix dactylopii on P. citri on lemon and acid<br />
lime and was responsible for the decline <strong>of</strong> mealybug populations (Mani<br />
1994).<br />
C. peregrinus has been introduced into California (Flanders 1951), Italy<br />
(Bartlett 1978) and Bermuda (Bennett and Hughes 1959).<br />
Cryptolaemus montrouzieri Col.: Coccinellidae<br />
This general predator <strong>of</strong> mealybugs, which also feeds on some other scales<br />
(Eriococcus sp., Pulvinaria spp.) and aphids, is native to eastern Australia. It<br />
is the most widely distributed <strong>of</strong> all natural enemies <strong>of</strong> mealybugs, a count in<br />
1978, covering the past 80 years, listing more than 40 countries, geographic<br />
areas or islands into which it has been imported. In many instances it was<br />
introduced against mealybugs other than P. citri and sometimes against<br />
coccids such as Pulvinaria spp., which produce egg masses similar to those<br />
<strong>of</strong> mealybugs (Bartlett 1978).<br />
Both larvae and adults feed voraciously on all mealybug stages, for<br />
example a larva is recorded as consuming an average <strong>of</strong> 3331 host eggs<br />
(Oncuer & Bayhan 1982) and females need to consume at least 8 P. citri for<br />
normal egg production (Reddy et al. 1991). C. montrouzieri does not<br />
distinguish between unparasitised P. citri and mealybugs parasitised by<br />
Leptomastix dactylopii (Prakasan and Bhat 1985). Adults mate 1 or 2 days<br />
after emergence and, 5 to 6 days later, females begin ovipositing in or near<br />
host egg masses. About 100 eggs are deposited in 1 month. These hatch in 4<br />
to 8 days, and wax-covered larvae develop in 12 to 20 days, so that the life<br />
cycle can be completed in slightly less than a month (27.7 days at<br />
25.5¡ ± 1¡C: Oncuer and Bayhan 1982), although there are usually only 4<br />
generations a year. Development stops below 10¡C and freezing<br />
temperatures are lethal. Pupae, and occasionally adults, are capable <strong>of</strong><br />
hibernating. Hot dry climates are tolerated, but high humidities are said to be<br />
detrimental. C. montrouzieri thrives when host density is high and, under<br />
these conditions, is capable <strong>of</strong> providing spectacular control. However its<br />
searching ability and natural spread is poor, so it <strong>of</strong>ten dies out locally when<br />
hosts become scarce (Bodenheimer 1928; Cole 1933; Mineo 1967).<br />
Methods have been developed for the production <strong>of</strong> mealybugs and<br />
C. montrouzieri that permit the production and release <strong>of</strong> large numbers <strong>of</strong><br />
the predators at low cost (Branigan 1916, Smith and Armitage 1920, 1931;<br />
Fisher 1963; Chacko et al. 1978; Oncuer and Koldas 1981).
4.15 Planococcus citri 313<br />
Diomus pumilio Col.: Coccinellidae<br />
Details <strong>of</strong> the biology and voraciousness <strong>of</strong> this predator, which was<br />
introduced from South Australia to Texas for biological control <strong>of</strong> P. citri,<br />
are given by Meyerdirk (1983).<br />
Leptomastidea abnormis Hym.: Encyrtidae<br />
This solitary endoparasite is possibly native to the Mediterranean where it<br />
was first recognised attacking P. citri (Viereck 1915), although it is now<br />
widespread, occurring in eastern USA, Canada, Brazil (Compere 1939) and<br />
many other countries.<br />
L. abnormis strongly prefers small 2nd instar mealybugs for oviposition,<br />
but also attacks first and third instars. Females begin to search for hosts soon<br />
after emergence. The number <strong>of</strong> eggs laid varies from 57 to over 300,<br />
although it is reported that only about 33 survive to the adult stage. Fertilised<br />
eggs give rise to females and unfertilised eggs to males.<br />
The inconspicuously stalked eggs are laid free in the haemolymph and<br />
hatch in 36 to 72 hours. The larvae consume haemolymph at first but, in the<br />
last instar, consume the entire body contents. The tailed larvae complete<br />
development in about 8 days and the life cycle may be as short as 17 days in<br />
the laboratory at 26¡C (or 25 days at 25¡ to 27¡C and 50Ð70% RH). In the<br />
laboratory females attained their maximum progeny production at 24¡C and<br />
this remained constant up to 34¡C (Tingle and Copland 1989). In the field a<br />
generation in summer takes about 1 month. There may be 5 or 6 generations<br />
a year, adults living 11 days if provided with honey and water (Viereck<br />
1915; Smith 1916, 1917; Perez 1929; Rivnay and Perzelan 1943; Clancy<br />
1944; Viggiani and Maresca 1973).<br />
Leptomastix dactylopii Hym.: Encyrtidae<br />
This solitary endoparasitoid prefers 3rd instar and young (but not egglaying)<br />
females and occasionally attacks 1st and 2nd instars (Bartlett 1978;<br />
Mani 1995). It is presumed to be native to Brazil, although found also in the<br />
West Indies and parts <strong>of</strong> southern USA (Compere 1939). In the field it<br />
appears to be specific to P. citri (Bartlett 1978; Sinadskii and<br />
Kozarzhevskaya 1980; Nagarkatti et al. 1992), but it can be reared on<br />
Planococcus lilacinus (Mani 1995), P. pacificus (Nagarkatti et al. 1992),<br />
Phenacoccus solani (Lloyd 1964) and Pseudococcus comstocki (Clancy<br />
1944). Its reported attack on Dysmicoccus brevipes in Hawaii and on<br />
Pseudococcus vitis in USSR was probably on P. citri (Kobakhidze 1965;<br />
Bartlett 1978).<br />
It has been used in suppression <strong>of</strong> P. citri in USA (Fisher 1963), Procida<br />
island and mainland Italy (Luppino 1979), Cyprus (Krambias and Kontzonis<br />
1980) and India (Krishnamoorthy 1990).
314 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Adults live up to 35 days and longer at 15¡ than at 7¡ or 25¡C (Yigit et al.<br />
1994) although maximum progeny are produced at 30¡ (Tingle and Copland<br />
1989). Parasitised hosts are generally rejected after simple antennal contact<br />
but, if not then, also following defensive behaviour <strong>of</strong> the host or possibly<br />
after detection <strong>of</strong> the egg stalk emerging from the surface <strong>of</strong> the host. If not<br />
rejected earlier, they may be rejected after insertion <strong>of</strong> the ovipositor<br />
(Baaren and Nenon 1994). About 18 eggs are laid each day, up to a total <strong>of</strong><br />
300 per female. These hatch in 1.5 to 2 days at 28¡C and there are four larval<br />
instars, each <strong>of</strong> about 2 days. The pupal stage lasts 7 to 8 days. In Italy there<br />
are 6 (and a partial 7th) generations per year (Zinna 1959, 1960) and in<br />
Tashkent 5 generations (Roxanova and Loseva 1963).<br />
More males than females are produced from young than from old adult<br />
P. citri (Su and Li 1993; Mani 1995), more females from larger hosts and<br />
more males from smaller larval instars (Jong and Alphen 1988, 1989).<br />
Additional information on the biology <strong>of</strong> L. dactylopii is given by Lloyd<br />
(1958, 1964, 1966) and Tingle and Copland (1989).<br />
The original introduction <strong>of</strong> L. dactylopii from Brazil to California in<br />
1934 was based on a single pair (Compere 1939). The extent to which the<br />
progeny <strong>of</strong> this pair may have had genes from later introductions added to<br />
the gene pool is quite unclear. There may thus be good justification for<br />
obtaining fresh stock from matching climatic zones in Brazil if new<br />
introductions are to be made.<br />
Odontochrysa (= Chrysopa) lacciperda Neu.: Chrysopidae<br />
Details <strong>of</strong> the biology and voracity <strong>of</strong> this lacewing predator <strong>of</strong> P. citri are<br />
provided by Krishnamoorthy (1988).<br />
Pseudaphycus maculipennis Hym.: Encyrtidae<br />
This species, studied in Ukraine, for the biological control <strong>of</strong> P. citri, is said<br />
to be specific (Sinadskii and Kozarzkevskaya 1980).<br />
Scymnus includens Col.: Coccinellidae<br />
The life cycle and rearing details <strong>of</strong> this important predator <strong>of</strong> P. citri in Italy<br />
are described by Tranfaglia and Viggiani (1973).<br />
Spalgis epius Lep.: Lycaenidae<br />
The predatory larvae <strong>of</strong> this lycaenid butterfly are <strong>of</strong>ten the commonest<br />
natural enemies <strong>of</strong> P. citri in India. They also attack Planococcus lilacinus,<br />
Chloropulvinaria psidii and Ferrisia virgata (Chacko et al. 1977).
Comments<br />
4.15 Planococcus citri 315<br />
P. citri is frequently only one <strong>of</strong> several pests <strong>of</strong> importance on the economic<br />
crops that it infests and, if biological control is contemplated, it is advisable<br />
to have its identity confirmed by a competent taxonomist. Its abundance is<br />
increased when it is tended by ants for its honeydew and <strong>of</strong>ten, by the<br />
injudicious use <strong>of</strong> insecticides (against it or accompanying pests). This is<br />
mainly because <strong>of</strong> the adverse effects on natural enemies and sometimes<br />
because low levels <strong>of</strong> insecticide may stimulate egg-laying. There has thus<br />
been considerable effort, within an Integrated Pest Management framework<br />
and with varying degrees <strong>of</strong> success, to develop biological control <strong>of</strong> each <strong>of</strong><br />
the important pests in a complex, for example on citrus, grape vines and in<br />
glasshouses. This has also involved the careful selection <strong>of</strong> pesticides (if<br />
these are still required) that have the least possible adverse effect on the<br />
major natural enemies.<br />
Where natural enemies already present are not adequate, the almost<br />
universal response has been to introduce the encyrtid Leptomastix dactylopii<br />
and the coccinellid Cryptolaemus montrouzieri (if the latter is not already<br />
present as a result <strong>of</strong> introductions for other pests).<br />
Both species are affected by the winter in temperate zones and survive<br />
less well than P. citri in the Mediterranean region. Under these<br />
circumstances, classical biological control seldom provides economic<br />
control alone and requires augmentation <strong>of</strong> the natural enemies from time to<br />
time.<br />
Where P. citri is still a problem in the field in warm regions and<br />
Cryptolaemus montrouzieri is not present, serious thought should be given<br />
to introducing the latter. More importantly, however, if not already present<br />
Leptomastix dactylopii (<strong>of</strong> Brazilian origin) should be <strong>of</strong> highest priority,<br />
followed by Leptomastidea abnormis and Anagyrus pseudococci (both <strong>of</strong><br />
Mediterranean origin and capable <strong>of</strong> maintaining populations <strong>of</strong> low levels<br />
under slightly cooler conditions). Under <strong>Southeast</strong> <strong>Asian</strong> and Pacific<br />
conditions, Coccidoxenoides peregrinus (<strong>of</strong> south China or Indian origin)<br />
deserves special attention.<br />
If Planococcus citri proves to be <strong>of</strong> south China origin (and this<br />
hypothesis requires confirmation) it is surprising that only two parasitoids<br />
have been reportedÑthe encyrtid Coccidoxenoides peregrinus and the<br />
platygasterid Allotropa citri. A thorough survey in this region might well<br />
reveal additional valuable specific or near specific parasitoids. There are<br />
good grounds for optimism that biological control <strong>of</strong> P. citri can be<br />
improved in warmer regions by establishing, if missing, any one <strong>of</strong> the<br />
foregoing natural enemies.
316 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
P. citri is one <strong>of</strong> a group <strong>of</strong> pests that commonly cause problems in<br />
glasshouses in Europe and North America. The mass production and release<br />
from time to time, almost always <strong>of</strong> a predator (especially Cryptolaemus<br />
montrouzieri, but sometimes also Nephus reunioni) and one or more<br />
encyrtid parasitoids (especially Leptomastix dactylopii and Leptomastidea<br />
abnormis but, on occasion also, Coccidoxenoides peregrinus, Anagyrus<br />
pseudococci and Chrysoplatycerus splendens) has generally removed the<br />
need to use insecticides. Examples <strong>of</strong> control in glasshouses include those<br />
from Belgium (Ronse 1990), Netherlands (Heanekam et al. 1987), France<br />
(Panis and Brun 1971), U.K. (Copland 1983, Hussey and Scopes 1985;<br />
Tingle and Copland 1988), Israel (Rubin 1985) and USA (Summy et al.<br />
1986).
4.16 Trichoplusia ni<br />
India<br />
20°<br />
Myanmar<br />
++ Laos<br />
0°<br />
20°<br />
China<br />
+<br />
Thailand<br />
++<br />
Cambodia<br />
++<br />
Vietnam<br />
+<br />
Malaysia<br />
Singapore<br />
Brunei<br />
P<br />
Indonesia<br />
Taiwan<br />
+<br />
Philippines<br />
Australia<br />
Papua<br />
New Guinea<br />
317<br />
The cabbage looper, Trichoplusia ni,<br />
<strong>of</strong> North American origin, attacks cabbage (and other<br />
Brassicaceae), cotton, lettuce, tomatoes and a very wide range <strong>of</strong> other cultivated crops and<br />
wild hosts. In North America it is maintained for much <strong>of</strong> the time at sub-economic levels by a<br />
wide range <strong>of</strong> natural enemies, but damaging outbreaks do occur, particularly when its natural<br />
enemies are killed by insecticides applied against other pests in the same crop.<br />
The major predators, which together cause considerable mortality, are widely polyphagous,<br />
and hence are unlikely to be considered seriously as classical biological control agents. Several,<br />
among its 120 or so parasitoids are somewhat more host specific and are worth serious<br />
consideration. They include species <strong>of</strong> Trichogramma egg parasitoid; Copidosoma<br />
truncatellum (an egg-larval parasitoid); and the larval parasitoids Hyposoter exiguae,<br />
Microgaster brassicae and Voria ruralis.<br />
High larval mortality is frequently produced by a valuable, naturally occurring, nuclear<br />
polyhedral virus, particularly late in the season when T. ni populations are high and rainfall is<br />
adequate.<br />
There appear to be good reasons for optimism that the establishment <strong>of</strong> suitable missing<br />
natural enemies in regions into which T. ni has spread would assist in maintaining its populations<br />
at sub-economic levels.<br />
20°<br />
0°<br />
20°
318 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Trichoplusia ni (HŸbner)<br />
Rating<br />
Origin<br />
Distribution<br />
Biology<br />
Lepidoptera: Noctuidae<br />
cabbage looper<br />
Synonym: In North America T. ni has sometimes been referred to<br />
as Autographa brassicae (Riley).<br />
<strong>Southeast</strong> Asia China<br />
7 ++ Myan, Thai, Camb + (all 14 southern Provinces)<br />
+ Viet<br />
P Indo<br />
T. ni is native to the southern half <strong>of</strong> North America.<br />
Widespread in southern Europe, North, East and South Africa, extending<br />
eastwards through Pakistan, India and Bangladesh to much <strong>of</strong> <strong>Southeast</strong><br />
Asia, to China, Taiwan, Korea and Japan; not yet present in Papua New<br />
Guinea, Australia, New Zealand or the oceanic Pacific; present in South<br />
America in Argentina, Bolivia, Brazil, Chile, Colombia and Uruguay<br />
(Apablaza and Norero 1993; CIE 1974b). In North America, T. ni<br />
overwinters in the south, re-invading northern States each spring.<br />
Adult T. ni are mottled brownish in colour. The forewings, producing a span<br />
<strong>of</strong> about 3.8 cm, each bear an 8-shaped silvery mark near the middle. Adults<br />
are mostly active at night, but also on dull days. By day, they rest on the<br />
underside <strong>of</strong> host plants, in the debris at their base, or in vegetation bordering<br />
a cultivated crop. Adults are capable <strong>of</strong> flying long distancesÑ700 km<br />
northwards from southern Texas (Lingren et al. 1993) 161 km from land into<br />
the Gulf <strong>of</strong> Mexico, and up to 1500m in California (Kreasky et al. 1972).<br />
They move readily between cultivated and wild hosts. A female may lay<br />
her own bodyweight in eggs, but requires access to nectar and moisture to do<br />
so. After emerging from the pupa, there is a pre-ovipositional period <strong>of</strong> about<br />
4 days, after which mating begins and can occur up to 16 days. A female may<br />
produce well over 1000 viable eggs. Peak egg deposition is <strong>of</strong>ten correlated<br />
with the lunar cycle, a rapid rise in egg density on cotton occurring shortly
Host plants<br />
4.16<br />
Trichoplusia ni<br />
319<br />
after full moon. There are 3 generations a year in southern California, but<br />
breeding is continuous in the Caribbean (McKinney 1944; Kishaba et al.<br />
1967; Ehler and van den Bosch 1974; Ehler 1977a; Debolt et al. 1984;<br />
Mitchell and Chalfant 1984).<br />
In cotton, a single egg is laid on the underside <strong>of</strong> a mature leaf in the<br />
upper half <strong>of</strong> the plant, but seldom in a terminal. On hatching, the larva<br />
generally feeds on the underside <strong>of</strong> the leaf near the egg, later moving from<br />
leaf to leaf as it passes through 5 instars during 2 to 4 weeks (Ehler 1977a).<br />
Total development time ranges from 19.9 days at 30¡C to 40.4 days at 20¡C<br />
(Jackson et al. 1969). In India at 25¡C the egg stage lasted 2.06 days, the<br />
larval stage <strong>of</strong> 5 instars 12.38 days, the prepupal 1 day, the pupal 7.27 days<br />
and the adult 7.32 days (Gaikwad et al. 1983). Additional data are provided<br />
by Chi and Tang (1993) and Yadav et al. (1983). Flight and mating activity<br />
are diminished at temperatures less than 16¡C and the threshold for larval<br />
development lies between 10¡C and 13¡C. The larva has 3 pairs <strong>of</strong> true legs<br />
on the thorax and 3 pairs <strong>of</strong> fleshy abdominal prolegs near the posterior end.<br />
It crawls by doubling up to form a loop, thus projecting the body forward.<br />
Larvae are green with a white lateral line and 2 whitish lines along the<br />
middle <strong>of</strong> the dorsal surface. After a brief prepupal period, pupation occurs<br />
in a loosely spun cocoon either on the underside <strong>of</strong> a leaf or in plant debris at<br />
the soil surface (Ehler 1977a). There is no diapause (Fye 1979).<br />
Eggs and larvae, but also pupae, <strong>of</strong> T. ni are believed to be readily<br />
transhipped in vegetables and cut flowers (Poe and Workman 1984).<br />
Male T. ni are powerfully attracted to the sex pheromone emitted by<br />
virgin females and will fly long distances upwind under its influence. The<br />
pheromone is produced in a gland situated dorsally between the 8th and 9th<br />
abdominal segments. Six components have been identified and are required<br />
to ensure specificity to T. ni.<br />
The major component is (Z)-7-dodecenyl<br />
acetate, known as looplure, which also attracts males <strong>of</strong> other looper species.<br />
Looplure, with or without other components, has been used for trapping<br />
males and also in mating disruption experiments. Male T. ni also produce a<br />
pheromone (with at least 3 components) which attracts both females<br />
(especially when starved) and males. ( Bjšstad et al. 1984; McLaughlin<br />
1984; Heath et al. 1992; Dunkelblum and Mazor 1993; Landolt 1995;<br />
Landolt et al. 1996).<br />
In 1966, larvae <strong>of</strong> T. ni were recorded causing damage to at least 119 species,<br />
varieties and cultivars in 29 families <strong>of</strong> plants (Sutherland 1966) and that<br />
number has increased steadily over the years to over 160 species in 36
320 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Damage<br />
families, although cultivated brassicas are those most favoured when<br />
available (Martin et al. 1976a; Sutherland and Greene 1984). Brassicas and<br />
cotton are most frequently cited as being damaged, although the list <strong>of</strong><br />
economic crops affected also includes asparagus, beans, sugarbeet,<br />
cantaloupes, capsicum, carrot, celery, maize (silks), cucumber, lettuce,<br />
parsley, pea, potato, soybean, spinach, squash, tobacco, tomato and<br />
watermelon.<br />
At times, serious infestations occur, but T. ni is generally regarded as a<br />
secondary pest whose numbers increase late in the season (Ehler 1977a,b).<br />
Differences in susceptibility to T. ni have been found in cabbage and<br />
related brassicas, in cotton and in lettuce (Cuthbert and Kishaba 1984), in<br />
tomato (Sinha and McLaren 1989) and in soybeans (Luedders et al. 1978;<br />
Khan et al. 1986), but these largely remain to be exploited. Transgenic<br />
cotton lines containing Bacillus thuringiensis toxin genes limited damage to<br />
initial feeding sites, compared with more extensive skeletonisation in 2<br />
control cultivars (Flint et al. 1995). Transgenic Bt canola (rape) showed<br />
excellent resistance to T. ni (Stewart et al. 1996).<br />
Larvae are easily reared on an artificial diet (e.g. Shorey and Hale 1965;<br />
Honda et al. 1996).<br />
The cabbage looper is a major pest <strong>of</strong> commercial brassicas in North<br />
America and many other areas where it occurs and causes significant<br />
damage also, in particular, to lettuce, tomatoes, celery and cotton. Indeed,<br />
Schwartz (1983) claimed that, if uncontrolled, 92% loss would be sustained<br />
in the cotton yield in USA, compared with 30% if controlled. Larvae chew<br />
large irregular holes, leaving only main veins, in the outer leaves <strong>of</strong> cabbage,<br />
cauliflower and related plants, <strong>of</strong>ten leaving them riddled with holes. Later,<br />
the outer layers <strong>of</strong> cabbage heads are eaten and masses <strong>of</strong> faecal pellets<br />
contaminate the feeding sites. So much leaf tissue is eaten that heads <strong>of</strong><br />
cabbage and cauliflower are stunted and other leafy vegetables are rendered<br />
unfit to eat.<br />
Damage caused to cotton by the larvae consuming leaves is <strong>of</strong>ten<br />
considered less serious, since it frequently occurs late in the growth <strong>of</strong> the<br />
cotton plant, so that it may not have a major effect on yield. Cabbage looper<br />
larvae are essentially foliage feeders and cause their damage in this way.<br />
Natural enemies<br />
Over the last few decades T. ni has become a very widely used laboratory<br />
insect, particularly in North America. As a result, there are many papers
4.16<br />
Trichoplusia ni<br />
321<br />
describing laboratory experiments in which parasitoids, predators and/or<br />
pathogens have been tested on T. ni eggs or larvae. It is <strong>of</strong>ten not possible to<br />
determine from these accounts whether or not the natural enemy involved<br />
has been found attacking eggs or larvae <strong>of</strong> T. ni in the field and hence<br />
possibly a useful control agent. If T. ni is susceptible to the agent in the<br />
laboratory, such records have <strong>of</strong>ten been included, although behavioural or<br />
other factors might well influence its effectiveness under field conditions.<br />
No attempt has been made to include reports <strong>of</strong> all <strong>of</strong> the minor natural<br />
enemies, especially in the earlier literature.<br />
As will become clearer when the situation is discussed later, many<br />
parasitoids (Table 4.16.1), predators (Table 4.16.2) and pathogens (Table<br />
4.16.3) have been recorded attacking the cabbage looper in the field and/or<br />
in the laboratory and there is little doubt that looper populations are<br />
frequently kept at sub-economic levels by their action. A number <strong>of</strong> the<br />
natural enemies recorded are not widespread in distribution and appear to be<br />
incidental records.<br />
Many <strong>of</strong> the predators are widely polyphagous and attack insect pests in<br />
several orders, although a few are considerably more selective than that.<br />
Parasitoids, on the other hand, tend to be significantly less polyphagous than<br />
predators and some appear to be confined to T. ni,<br />
at least in certain crops.<br />
Where parasitoids have a relatively broad host range, available records<br />
indicate that this extends mainly to larvae <strong>of</strong> other Lepidoptera (generally<br />
pest species) in the same crop situation. The extent to which it extends also to<br />
non-pest, non-target species is not documented. Nevertheless, there are<br />
several species that merit serious consideration as candidates for classical<br />
biological control.<br />
All <strong>of</strong> the Trichogramma species oviposit and develop in the host egg. A<br />
few others (eg. Copidosoma truncatellum,<br />
Chelonus blackburni, Chelonus<br />
insularis)<br />
oviposit in the egg and develop in the host larva; and the remainder<br />
are larval and/or pupal parasitoids.<br />
Naturally occurring epizootics <strong>of</strong> nuclear polyhedrosis virus in medium<br />
to large T. ni larvae are considered to be the major mortality factor affecting<br />
them on cabbage in summer and autumn in southern California (Oatman and<br />
Platner 1969), on broccoli in Virginia (H<strong>of</strong>master 1961) and on cabbage in<br />
North Carolina (Elsey and Rabb 1970b). Polyhedrosis was seldom a major<br />
factor on cotton in southern California (Ehler 1977b), although outbreaks<br />
did occur late in the season or at times <strong>of</strong> high T. ni abundance (Ehler and van<br />
den Bosch 1974).<br />
Although the mortality produced is probably not significant, birds and<br />
bats are known to feed on moths in flight; and earwigs on adults resting<br />
beneath host plants (McKinney 1944; Sutherland 1966).
Table 4.16.1<br />
Parasitoids <strong>of</strong> Trichoplusia ni<br />
Species<br />
HYMENOPTERA<br />
BRACONIDAE<br />
Country Reference<br />
Cardiochiles nigriceps<br />
USA Harding 1976<br />
Chelonus blackburni<br />
USA Fye & Jackson 1973; Jackson et al. 1979<br />
Chelonus curvimaculatus<br />
USA Soldevila & Jones 1991, 1994<br />
Chelonus nr curvimaculatus<br />
USA Jones et al. 1981, 1990; Jones 1986<br />
Chelonus formosanus<br />
Taiwan Chou 1981<br />
Chelonus insularis<br />
USA Ehler et al. 1973; Ehler & van den Bosch 1974; Jones 1982; Henneberry et al.<br />
1991<br />
Chelonus sp. USA BŸhler et al. 1985<br />
Cotesia autographae<br />
USA Muesebeck & Krombein 1951<br />
Cotesia congregata<br />
USA Riley 1883<br />
Cotesia glomerata<br />
USA Muesebeck & Krombein 1951; van den Bosch & Hagen 1966; Ehler 1977a<br />
Cotesia laeviceps<br />
USA Oatman et al. 1983a<br />
Cotesia marginiventris<br />
USA van den Bosch & Hagen 1966; Ehler & van den Bosch 1974, Harding 1976,<br />
Latheef & Irwin 1983; Henneberry et al. 1991<br />
Cotesia plutellae<br />
India Manjunath 1972; Joshi & Sharma 1974<br />
Cotesia ruficrus<br />
USA<br />
India<br />
McCutcheon et al. 1983<br />
Manjunath 1972<br />
Cotesia spp. USA Harding 1976<br />
Cotesia yakutatensis<br />
Meteorus autographae<br />
USA<br />
India<br />
Miller & West 1987<br />
Manjunath 1972<br />
USA Muesebeck & Krombein 1951; Grant & Shepard 1984<br />
322 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong>
Table 4.16.1 (contÕd) Parasitoids <strong>of</strong> Trichoplusia ni<br />
Species<br />
HYMENOPTERA<br />
BRACONIDAE (contÕd)<br />
Country Reference<br />
Meteorus laphygmae<br />
USA Harding 1976<br />
Microgaster (= Microplitis)<br />
brassicae USA McKinney 1944; van den Bosch & Hagen 1966; Clancy 1969; Oatman & Platner<br />
1969; Ehler & van den Bosch 1974; Harding 1976; Ehler 1977a; Jones 1982;<br />
Oatman et al. 1983a; Henneberry et al. 1991<br />
Microgaster plutellae<br />
USA Oatman & Platner 1969; Oatman et al. 1983a<br />
Microplitis alaskensis<br />
USA Butler 1958a<br />
Rogas granulatus<br />
USA De Gant 1930<br />
Rogas molestus<br />
USA Butler 1958a<br />
Rogas rufocoxalis<br />
USA McKinney 1944<br />
Rogas sp. USA Wall & Berberet 1975<br />
Snellenius manilae<br />
CHALCIDIDAE<br />
Taiwan Chou 1981<br />
Brachymeria intermedia<br />
Italy<br />
Dindo 1993<br />
USA<br />
Thompson 1980<br />
Brachymeria lasus<br />
USA Thompson 1983a,b<br />
Brachymeria ovata<br />
USA Elsey & Rabb 1970b; Harding 1976; Patana et al. 1978; Chamberlin & Kok<br />
1986; Grant & Shepard 1987<br />
Spilochalcis flavopicta<br />
USA Harding 1976<br />
Spilochalcis side<br />
USA Harding 1976<br />
Spilochalcis sp. nr mariae<br />
USA Harding 1976<br />
4.16<br />
Trichoplusia ni<br />
323
Table 4.16.1 (contÕd) Parasitoids <strong>of</strong> Trichoplusia ni<br />
Species<br />
HYMENOPTERA<br />
Country Reference<br />
ENCYRTIDAE<br />
Copidosoma floridanum USA Strand et al. 1991; Baehrecke et al. 1993; Grbk et al. 1992; Ode & Strand 1995<br />
Copidosoma sp. USA Ehler 1977a<br />
Copidosoma truncatellum Canada<br />
USA<br />
Brazil<br />
Harcourt 1963<br />
Riley 1883; McKinney 1944; Pimentel 1961; Oatman 1966; van den Bosch &<br />
Hagen 1966; Clancy 1969; Oatman & Platner 1969; , Ehler & van den Bosch<br />
1974; Ehler 1977a; Latheef & Irwin 1983; Oatman et al. 1983a; Roltsch & Mayse<br />
1983; Chamberlin & Kok 1986<br />
Silva & Santos 1980<br />
EULOPHIDAE<br />
Baryscapus galactopus USA Peck 1963<br />
Euplectrus comstockii USA McKinney 1944; Harding 1976; Coudron et al. 1994<br />
Euplectrus platyhypenae USA Wall & Berberet 1974, 1975; Coudron et al. 1990; Kelly & Coudron 1990,<br />
Euplectrus sp. Santiago van Harten & Miranda 1985<br />
Pediobius facialis USA Oatman & Platner 1969<br />
Pediobius nr facialis USA Parkman et al. 1983<br />
ICHNEUMONIDAE<br />
Angitia insularis USA Hayslip et al. 1953; Harding 1976<br />
Campoletis flavicincta USA Krombein et al. 1979; Oatman et al. 1983a<br />
Campoletis sonorensis USA Cook et al. 1984<br />
Campoletis sp. USA Oatman et al. 1983a<br />
Campoletis websteri USA Harding 1976<br />
Casinaria infesta USA Harding 1976<br />
Cryptus rutovinctus USA Krombein et al. 1979<br />
324 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong>
Table 4.16.1 (contÕd) Parasitoids <strong>of</strong> Trichoplusia ni<br />
Species<br />
HYMENOPTERA<br />
Country Reference<br />
ICHNEUMONIDAE (contÕd)<br />
Diadegma insulare USA Martin et al. 1982<br />
Diadegma plutellae USA Harding 1976<br />
Diadegma spp. USA Sutherland 1966<br />
Echthromorpha punctum India Manjunath 1972<br />
Enicospilus sp. India Manjunath 1972<br />
Gambrus ultimus USA Chamberlin & Kok 1986<br />
Gelis tenellus USA Sutherland 1966<br />
Hyposoter exiguae USA Oatman 1966; van den Bosch & Hagen 1966; Clancy 1969; Oatman & Platner<br />
1969, Ehler & van den Bosch 1974; Ehler 1977a; Jones 1982; Oatman et al.<br />
1983a,<br />
Iseropus stercorator orgyiae USA Sutherland 1966<br />
Itoplectis conquisator USA<br />
Canada<br />
Muesebeck & Krombein 1951<br />
Harcourt 1963<br />
Microcharops bimaculata Brazil Silva & Santos 1980<br />
Microcharops tibialis USA Harding 1976<br />
Nepiera fuscifemora USA Clancy 1969; Oatman et al. 1983a<br />
Netelia sp. USA Watson et al. 1966<br />
Patrocloides montanus USA Clancy 1969; Ehler & van den Bosch 1974; Ehler 1977a; Fox et al. 1996<br />
Pimpla aequalis USA Sutherland 1966<br />
Pristomerus sp. USA Harding 1976<br />
Pristomerus spinator USA Harding 1976<br />
4.16 Trichoplusia ni 325
Table 4.16.1 (contÕd) Parasitoids <strong>of</strong> Trichoplusia ni<br />
Species<br />
HYMENOPTERA<br />
Country Reference<br />
ICHNEUMONIDAE (contÕd)<br />
Pterocormus gestuosus USA Mitchell 1961<br />
Stenichneumon culpator<br />
cincticornis<br />
Canada<br />
USA<br />
Harcourt 1963<br />
Chamberlin & Kok 1986<br />
Vulgichneumon brevicinctor USA Chamberlin & Kok 1986<br />
PTEROMALIDAE<br />
Pediobius nr sexdentatus USA Oatman & Platner 1969; Oatman et al. 1983a<br />
SCELIONIDAE<br />
Telenomus solitus Guatamala Johnson 1983; Navasero & Oatman 1989<br />
Telenomus sp. USA Martin et al. 1984<br />
TRICHOGRAMMATIDAE<br />
Trichogramma australicum India Manjunath 1972<br />
Trichogramma brevicapillum Brazil<br />
USA<br />
Hohmann et al. 1988b<br />
H<strong>of</strong>fmann et al. 1990<br />
Trichogramma chilotraeae India Manjunath 1972<br />
Trichogramma deion Brazil<br />
USA<br />
Hohmann et al. 1988a,b<br />
H<strong>of</strong>fmann et al. 1990<br />
Trichogramma evanescens USA Oatman et al. 1968<br />
Trichogramma exiguum USA Martin et al. 1982; Roltsch & Mayse 1983; Campbell et al. 1991<br />
Trichogramma japonicum India Manjunath 1972<br />
Trichogramma minutum USA McKinney 1944; Marston & Ertle 1973; Manweiler 1986<br />
Trichogramma platneri Brazil<br />
USA<br />
Hohmann et al. 1988a,b<br />
Manweiler 1986<br />
326 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong>
Table 4.16.1 (contÕd) Parasitoids <strong>of</strong> Trichoplusia ni<br />
Species<br />
HYMENOPTERA<br />
Country Reference<br />
TRICHOGRAMMATIDAE (contÕd)<br />
Trichogramma pretiosum USA Oatman 1966; H<strong>of</strong>fmann et al. 1975, 1990; Martin et al. 1976b; Ehler 1977a;<br />
Oatman & Platner 1978; Butler & Lopez 1980; Oatman et al. 1983a<br />
Trichogramma semifumatum USA Ehler & van den Bosch 1974<br />
Trichogramma spp. USA van den Bosch & Hagen 1966; Harding 1976; Jones 1982<br />
Trichogramma thalense Brazil<br />
USA<br />
DIPTERA<br />
Hohmann et al. 1988b<br />
H<strong>of</strong>fmann et al. 1990<br />
SARCOPHAGIDAE<br />
Sacrodexia sternodontis USA Aldrich 1927<br />
Senotainia sp. USA Manjunath 1972<br />
TACHINIDAE<br />
Achaetoneura archippivora USA Butler 1958b<br />
Aplomya theclarum USA Harding 1976<br />
Archytas californiae USA van den Bosch & Hagen 1966<br />
Bessa remota Malaysia Jayanth & Nagarkatti 1984<br />
Carcelia sp. USA<br />
India<br />
Harding 1976<br />
Manjunath 1972<br />
Chetogena sp. USA van den Bosch & Hagen 1966; Chamberlin & Kok 1986<br />
Compsilura concinnata Canada<br />
USA<br />
Harcourt 1963<br />
Schaffner & Griswold 1934<br />
Eucelatoria armigera USA Butler 1958b, Clancy 1969, Henneberry et al. 1991, Oatman 1966, Oatman &<br />
Platner 1969, van den Bosch & Hagen 1966<br />
Eucelatoria nr armigera USA Harding 1976, Henneberry et al. 1991<br />
4.16 Trichoplusia ni 327
Table 4.16.1 (contÕd) Parasitoids <strong>of</strong> Trichoplusia ni<br />
Species<br />
DIPTERA<br />
Country Reference<br />
TACHINIDAE (contÕd)<br />
Eucelatoria rubentis USA Watson et al. 1966<br />
Euphorocera spp. USA Harding 1976<br />
Euphorocera tachinomoides USA Oatman 1966; Harding 1976<br />
Hypantrophaga sp. USA Harding 1976<br />
Lespesia achaetoneura USA Oatman 1966<br />
Lespesia archippivora USA Watson et al. 1966; Oatman & Platner 1969; Henneberry et al. 1991<br />
Lespesia sp. USA Clancy 1969; Chamberlin & Kok 1986<br />
Madremyia saundersii USA Oatman 1966; Oatman & Platner 1969<br />
Metachaeta (=Periscepsia) helymus USA Clancy 1969<br />
Phorocera sp. USA Sutherland 1966<br />
Phryxe vulgaris USA Sutherland 1966<br />
Sarcophaga spp. USA van den Bosch & Hagen 1966<br />
Schizocerophaga leibyi USA Harding 1976<br />
Siphona plusiae USA Clancy 1969; Harding 1976; Henneberry et al. 1991<br />
Siphona sp. USA Oatman & Platner 1969; Oatman et al. 1983a<br />
Voria edentata India Manjunath 1972<br />
Voria ruralis USA McKinney 1944; Butler 1958b; Pimentel 1961; van den Bosch & Hagen 1966;<br />
Oatman 1966; Clancy 1969; Oatman & Platner 1969; Elsey & Rabb 1970a;<br />
Ehler & van den Bosch 1974; Wall & Berberet 1975; Harding 1976; Ehler 1977a;<br />
Jones 1982; Latheef & Irwin 1983; Oatman et al. 1983a; Chamberlin & Kok<br />
1986; Gordon et al. 1987; Henneberry et al. 1991; Biever et al. 1992<br />
Winthemia nr montana USA Harding 1976<br />
Winthemia quadripustulala USA Allen 1925<br />
328 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong>
Table 4.16.1 (contÕd) Parasitoids <strong>of</strong> Trichoplusia ni<br />
Species<br />
DIPTERA<br />
Country Reference<br />
TACHINIDAE (contÕd)<br />
Winthemia rufopicta USA Chamberlin & Kok 1986<br />
Winthemia spp. USA Elsey & Rabb 1970b<br />
Zenilla blanda blanda USA West 1925<br />
4.16 Trichoplusia ni 329
Table 4.16.2 Some predators <strong>of</strong> Trichoplusia ni in USA<br />
Species Reference<br />
DERMAPTERA<br />
LABIDURIDAE<br />
Labidura riparia Strandberg 1981a,b<br />
Tawfik et al. 1972<br />
HEMIPTERA<br />
ANTHOCORIDAE<br />
Orius insidiosus Lingren et al. 1978<br />
Orius tristicolor Ehler et al. 1973; Ehler & van den Bosch 1974; Ehler 1977a; Wilson & Gutierrez 1980;<br />
Jones 1982; Jones et al. 1983<br />
LYGAEIDAE<br />
Geocoris pallens van den Bosch & Hagen 1966; Ehler et al. 1973; Ehler & van den Bosch 1974; Ehler<br />
1977a; Wilson & Gutierrez 1980<br />
Geocoris punctipes van den Bosch & Hagen 1966; Barry 1973; Barry et al. 1974; Walker & Turnipseed<br />
1976; Wilson & Gutierrez 1980; Reed et al. 1984<br />
NABIDAE<br />
Nabis alternatus van den Bosch & Hagen 1966; Barry 1973; Barry et al. 1974<br />
Nabis americ<strong>of</strong>erus van den Bosch & Hagen 1966; Ehler et al. 1973; Ehler & van den Bosch 1974; Ehler<br />
1977a; Stoltz and Stern 1979; Wilson & Gutierrez 1980<br />
Nabis roseipennis Reed et al. 1984<br />
PENTATOMIDAE<br />
Alcaeorrhynchus grandis McLain 1979<br />
Euthyrhynchus floridanus McLain 1979<br />
330 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong>
Table 4.16.2 (contÕd) Some predators <strong>of</strong> Trichoplusia ni in USA<br />
Species Reference<br />
Podisus maculiventris Hayslip et al. 1953; Ign<strong>of</strong>fo et al. 1977; Marston et al. 1978; Richman & Whitcomb<br />
1978; McLain 1979; Biever et al. 1982<br />
Stiretrus anchorago Richman 1977<br />
HEMIPTERA<br />
REDUVIIDAE<br />
Sinea complexa van den Bosch & Hagen 1966<br />
Sinea confusa van den Bosch & Hagen 1966<br />
Sinea diadema van den Bosch & Hagen 1966<br />
Sycanus indagator Greene & Shepard 1974<br />
Zelus bilobus Hayslip et al. 1953<br />
Zelus exsaguis Whitcomb and Bell 1964<br />
Zelus renardii van den Bosch & Hagen 1966<br />
Zelus tetracanthus van den Bosch & Hagen 1966<br />
NEUROPTERA<br />
CHRYSOPIDAE<br />
Chrysopa lanata Ru et al. 1975<br />
Chrysopa nigricornis van den Bosch & Hagen 1966<br />
Chrysopa rufilabris Ru et al. 1976<br />
Chrysopa spp. Pimentel 1961<br />
Chrysoperla carnea van den Bosch & Hagen 1966; Barry 1973; Barry et al. 1974; Ehler & van den Bosch<br />
1974; Ehler 1977a; Wilson & Gutierrez 1980<br />
HEMEROBIIDAE<br />
Hemerobius spp. van den Bosch & Hagen 1966<br />
4.16 Trichoplusia ni 331
Table 4.16.2 (contÕd) Some predators <strong>of</strong> Trichoplusia ni in USA<br />
Species Reference<br />
COLEOPTERA<br />
CARABIDAE<br />
Calosoma affine van den Bosch & Hagen 1966<br />
Calosoma peregrinator McKinney 1944<br />
Labia analis Reed et al. 1984<br />
COLEOPTERA<br />
COCCINELLIDAE<br />
Ceratomegilla maculata fuscilabris Pimentel 1961<br />
Coccinella novemnotata franciscana van den Bosch & Hagen 1966<br />
Coccinella transversoguttata Pimentel 1961<br />
Cycloneda sanguinea van den Bosch & Hagen 1966; Jones 1982<br />
Hippodamia convergens van den Bosch & Hagen 1966; Jones 1982; Jones et al. 1983<br />
Hippodamia parenthesis van den Bosch & Hagen 1966<br />
Hippodamia quinquesignata punctulata van den Bosch & Hagen 1966<br />
Olla v-nigrum van den Bosch & Hagen 1966<br />
Paranaemia vittegera van den Bosch & Hagen 1966<br />
MELYRIDAE<br />
Collops marginellus van den Bosch & Hagen 1966<br />
Collops vittatus van den Bosch & Hagen 1966<br />
DIPTERA<br />
SYRPHIDAE<br />
Mesograpta marginata Pimentel 1961<br />
Sphaerophoria cylindrica Pimentel 1961<br />
Sphaerophoria menthastri Pimentel 1961<br />
332 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong>
Table 4.16.2 (contÕd) Some predators <strong>of</strong> Trichoplusia ni in USA<br />
Species Reference<br />
DIPTERA<br />
SYRPHIDAE (contÕd)<br />
Sphaerophoria robusta Pimentel 1961<br />
Several species van den Bosch & Hagen 1966<br />
HYMENOPTERA<br />
VESPIDAE<br />
Mischocyttarus flavitarsis Bernays & Montelor 1989<br />
Polistes apachus Cornelius 1993<br />
Polistes metricus van den Bosch & Hagen 1966; Hunt 1984; Greenstone & Hunt 1993<br />
Vespula pensylvanica Warren 1990<br />
SPIDERS<br />
Misumena ratia Lockley et al. 1989<br />
Oxyopes salticus Reed et al. 1984; Lockley & Young 1988<br />
Pardosa spp. Reed et al. 1984<br />
Phidippus regius Edwards & Jackson 1993, 1994<br />
Phidippus spp. Edwards & Jackson 1993, 1994<br />
BIRDS<br />
Dendroica palmarum Strandberg 1981a<br />
Passerculus sandwichensis Strandberg 1981a<br />
4.16 Trichoplusia ni 333
334 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
More than 20 species <strong>of</strong> microorganism (viruses, bacteria, protozoa and<br />
fungi) are associated with T. ni, most <strong>of</strong> them isolated from field<br />
populations, but some from laboratory cultures. All were initially isolated<br />
from larvae, although at least one species <strong>of</strong> each group has also been<br />
isolated from pupae, adults and even eggs (Table 4.16.3). Details concerning<br />
the causative agents and the symptoms they produce are given by Ign<strong>of</strong>fo<br />
and Hostetter (1984). Many papers have been published in recent years to<br />
add details to the records in Table 4.16.3, especially in the field <strong>of</strong> viruses,<br />
but also adding to the range <strong>of</strong> other pathogens, such as a rickettsia-like<br />
organism (Browning et al. 1982). Naturally-occurring virus infection has<br />
been found to be a major mortality factor <strong>of</strong> T. ni larvae in the field,<br />
particularly late in the season when populations are high. A singleembedded<br />
nuclear polyhedrosis has been mass produced and applied with<br />
excellent results by a number <strong>of</strong> authors to several crops (Ign<strong>of</strong>fo and<br />
Hostetter 1984).<br />
Table 4.16.3 Major pathogens <strong>of</strong> Trichoplusia ni (Ign<strong>of</strong>fo and Hostetter<br />
1984)<br />
VIRUSES single-embedded nuclear polyhedrosis<br />
multiple-embedded nuclear polyhedrosis<br />
granulosis<br />
cytoplasmic polyhedrosis<br />
BACTERIA Bacillus thuringiensis<br />
Serratia marcescens<br />
PROTOZOA Nosema trichoplusiae<br />
Thelohania sp. nr. diazoma<br />
FUNGI Aspergillus flavus<br />
Beauveria bassiana<br />
Entomopthora gammae<br />
Entomopthora sphaerosperma<br />
Metarrhizium anisopliae<br />
Metarrhizium brunneum<br />
Nomuraea rileyi
4.16 Trichoplusia ni 335<br />
Introductions for biological control <strong>of</strong> T. ni<br />
There do not appear to have been any introductions <strong>of</strong> natural enemies<br />
specifically for cabbage looper, but rather for the complex <strong>of</strong> lepidopterous<br />
larvae with which it is almost always associated. Examples <strong>of</strong> such<br />
introductions are shown in Table 4.16.4.<br />
BRAZIL<br />
The natural enemies <strong>of</strong> T. ni larvae on cotton at 3 sites in Paran‡ Province<br />
included the fungus, Nomuraea rileyi (which killed 76% <strong>of</strong> larvae at one<br />
site), a virus disease (that killed up to 47% at two sites), the parasitoids<br />
Copidosoma truncatellum (reared from about 5% <strong>of</strong> larvae at 2 sites) and<br />
Microcharops bimaculata (reared from 7.5% <strong>of</strong> larvae at 1 site) and the<br />
fungus Entomopthora sp. (which killed 2.5% <strong>of</strong> larvae at 1 site) (Silva and<br />
Santos 1980). The natural enemies <strong>of</strong> T. ni on cotton in Mato Grosso are<br />
discussed by Bleicher et al. (1985) and on tomato in Sao Paulo by Gravena<br />
(1984).<br />
CARIBBEAN<br />
T. ni is usually a minor pest <strong>of</strong> Brassicaceae, although outbreaks<br />
occasionally cause serious defoliation <strong>of</strong> crops. A large number <strong>of</strong> predators<br />
attack larvae, in addition to the parasitoids that are listed in Table 4.16.5.
Table 4.16.4 Introductions for the biological control <strong>of</strong> lepidopterous larvae including Trichoplusia ni<br />
Species<br />
HYMENOPTERA<br />
BRACONIDAE<br />
From To When Result Reference<br />
Cotesia marginiventris Cape Verde Is 1981 + Lima & van Harten 1985<br />
Cotesia plutellae India Barbados, Jamaica 1969 + Alam 1992<br />
Cotesia ruficrus Australia USA 1981 ? McCutcheon et al. 1983<br />
Microplitis (= Microgaster) demolitor Australia USA 1981 ? Shepard et al. 1983;<br />
Norlund & Lewis 1985<br />
Microgaster rufiventris<br />
ENCYRTIDAE<br />
Egypt USA 1983 Ð McCutcheon & Harrison 1987<br />
Copidosoma floridanum<br />
EULOPHIDAE<br />
India Barbados pre 1985 + Alam 1992<br />
Pediobius nr facialis Japan USA pre 1983 * Parkman et al. 1983<br />
* no indication <strong>of</strong> field release<br />
336 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong>
4.16 Trichoplusia ni 337<br />
Table 4.16.5 Parasitoids and a fungus attacking T. ni in the Caribbean<br />
(Alam 1992)<br />
Species % Parasitisation<br />
Jamaica Barbados<br />
BRACONIDAE<br />
Cotesia sp. (glomerata group) 20.0<br />
Cotesia plutellae 29.6Ð70.0 3.5<br />
Glyptapanteles sp. (vitripennis group)<br />
CHALCIDIDAE<br />
0.5Ð 2.0<br />
Brachymeria sp. 2.4<br />
Brachymeria ovata<br />
ENCYRTIDAE<br />
0.5<br />
Copidosoma sp. 12.5<br />
Copidosoma floridanum 0.5Ð5.0<br />
Copidosoma (truncatellum group)<br />
EULOPHIDAE<br />
25.8<br />
Euplectrus platyhypenae<br />
TACHINIDAE<br />
Winthemia nr pinguis<br />
4.2<br />
and Winthemia nr pyrrhopyga 20.2Ð35.8<br />
Winthemia sp. 1 specimen only<br />
ENTOMOPHTHORALES 9.5Ð80.0<br />
NORTH AMERICA<br />
The cabbage looper is a widespread and <strong>of</strong>ten highly destructive pest <strong>of</strong><br />
cabbage and other Brassicaceae southwards in North America, from about<br />
the level <strong>of</strong> Ontario in Canada. Throughout this range it is associated with up<br />
to about a dozen other species <strong>of</strong> Lepidoptera. It is third in importance to the<br />
cabbage white butterfly, Pieris rapae, and the diamondback moth Plutella<br />
xylostella in Canada (Harcourt 1963) and New York State (Pimentel 1961)<br />
and about as important as these in southwestern USA (Oatman and Platner<br />
1969; Reid and Cuthbert 1957).<br />
In Ontario the encyrtid wasp Copidosoma truncatellum is the most<br />
important parasitoid and populations are frequently destroyed by a<br />
polyhedral virus (Harcourt 1963). In New York State a polyhedral virus<br />
(40% mortality) was the major factor affecting T. ni populations in 1957 but<br />
less important (7%) in 1958 when predators (especially spiders), caused 2%<br />
to 3% mortality (Pimentel 1961). In northwestern USA up to 14% <strong>of</strong> T. ni<br />
larvae were parasitised on cabbage by the tachinid fly, Voria ruralis (Biever<br />
et al. 1992). In southern California up to 39% (av. 7.8%) <strong>of</strong> T. ni eggs were<br />
parasitised by Trichogramma pretiosum, which was also reared from<br />
Plutella xylostella eggs. Twelve species <strong>of</strong> parasitoid were reared from T. ni
338 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
larvae and pupae, 7 <strong>of</strong> which were Hymenoptera and 5 Diptera (Table<br />
4.16.1). These produced an average <strong>of</strong> 38.9% mortality, with a maximum <strong>of</strong><br />
66.7% in late autumn. The tachinid, Voria ruralis, was the dominant<br />
parasitoid, especially during autumn and winter months. The ichneumon,<br />
Hyposoter exiguae, and the encyrtid, Copidosoma truncatellum, occurred<br />
most commonly during the summer and autumn months, when the latter was<br />
associated with a nuclear polyhedrosis virus. Together (and particularly the<br />
virus) they were responsible for most <strong>of</strong> the 60% larval mortality. Pupal<br />
mortality was low (2.0%) and was due to the pteromalid Pediobius<br />
sexdentatus (Oatman and Platner 1969). Also in southern California, Clancy<br />
(1969) reared 5 wasps and 5 tachinid flies (Table 4.16.1) from T. ni larvae<br />
collected from annual weeds (Malva sp., Chenopodium sp.), mustard, tree<br />
tobacco (Nicotiana glauca) and lucerne. The fly, Voria ruralis, was the most<br />
abundant parasitoid in autumn and winter and Copidosoma truncatellum the<br />
most important wasp, the total parasitisation from all species ranging from<br />
29.5% to 41.1%. A nuclear polyhedrosis virus killed 70.9% <strong>of</strong> larvae<br />
collected in summer from weeds and 63.8% from lucerne (Table 4.16.6).<br />
The mortality caused by the various natural enemies varied according to the<br />
season and host plant.<br />
Henneberry et al. (1991) recorded 12 species <strong>of</strong> parasitoid from larvae <strong>of</strong><br />
the loopers T. ni and Autographa californica on lettuce, lucerne, sugarbeet<br />
and cotton in southern California. Of these, the braconid Microgaster<br />
brassicae (30%) Voria ruralis (23%) and Copidosoma truncatellum (23%)<br />
were the most abundant, with parasitisation rates ranging from 0% to 91.8%<br />
according to season and crop. Average mortality from viral infection ranged<br />
between 0.6% and 7.2%, also depending upon the crop.<br />
Thirteen parasitoid species reared from T. ni on tomatoes in southern<br />
California (Table 4.16.1) caused mean parasitisation rates <strong>of</strong> larvae <strong>of</strong> 51.4%<br />
and 70.5% and <strong>of</strong> eggs <strong>of</strong> 24.6% and 53.4% respectively in two successive<br />
years. Hyposoter exiguae and Copidosoma truncatellum were the most<br />
abundant larval parasitoids and Trichogramma pretiosum the most<br />
important species attacking eggs. The data on population trends and<br />
percentage parasitisation suggested that there was a density-dependent<br />
relationship between T. ni and its parasite complex on tomato (Oatman et al.<br />
1983a).<br />
In Arizona, the most abundant parasitoid <strong>of</strong> T. ni larvae collected from<br />
weeds and cultivated crops was Voria ruralis, which was present throughout<br />
the year, with peak abundance (up to 100%) in late autumn and winter<br />
(McKinney 1944; Butler 1958b; Brubaker 1968). Five wasps and 1 tachinid<br />
fly were reared from larvae, 1 tachinid from pupae and Trichogramma
Table 4.16.6 Natural enemies <strong>of</strong> Trichoplusia ni larvae in southern California from weed hosts, lucerne and tree tobacco<br />
(Nicotiana glauca) (from Clancy 1969)<br />
Month No reared % pupating % killed by<br />
virus all parasites Voria ruralis all other<br />
Tachinidae<br />
Copidosoma<br />
truncatellum<br />
all other<br />
Hymenoptera<br />
1966 Collections from annual weeds<br />
May 94 2.1 63.8 34.0 4.3 11.7 10.6 7.4<br />
June 286 12.6 57.3 30.1 7.7 9.8 4.9 7.0<br />
July 327 4.6 70.9 24.2 4.0 10.1 5.5 4.6<br />
Aug 280 6.1 44.3 49.6 16.1 6.4 25.4 1.8<br />
Sept 193 18.6 31.6 49.7 12.4 9.3 30.0<br />
Oct 267 41.9 37.8 20.2 6.4 3.4 10.5<br />
Nov 162 47.5 25.9 26.5 9.3 1.2 16.0<br />
Dec<br />
1967<br />
151 4.6 39.1 56.3 38.4 0.7 16.6 0.7<br />
Jan 138 31.9 16.7 51.4 34.1 0.7 16.7<br />
Feb 63 61.9 11.1 27.0 11.1 15.9<br />
March 7 14.3 28.6 57.1 14.3 42.9<br />
April 12 41.7 33.3 25.0 25.0<br />
May 120 60.0 15.8 24.2 16.7 1.7 5.8<br />
Totals 2100 22.0 42.5 35.2 13.1 5.9 13.3 2.8<br />
Collections from lucerne<br />
149 6.7 63.8 29.5 6.0 20.8 0.7 2.0<br />
Collections from tree tobacco, Nicotiana glauca<br />
479 47.0 11.9 41.1 21.5 1.0 11.7 6.9<br />
4.16 Trichoplusia ni 339
340 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
minutum from eggs collected from lettuce. In addition, the carabid beetle,<br />
Calosoma peregrinator, fed readily on T. ni larvae (McKinney 1944).<br />
A study in southern Texas <strong>of</strong> the loopers T. ni and Chrysodeixis<br />
includens on a range <strong>of</strong> host plants revealed high levels (58% to 71%) <strong>of</strong><br />
mortality <strong>of</strong> larvae and pupae by 29 parasitoid species (Table 4.16.1) during<br />
all but 4 months <strong>of</strong> the year, all <strong>of</strong> which existed at rather low levels (Harding<br />
1976).<br />
Turning to cotton in southern California, T. ni is a secondary pest and<br />
there is good evidence that natural enemies, in particular several predators,<br />
are mainly responsible for its generally low pest status. The 4 major<br />
predators there are larvae <strong>of</strong> the green lacewing, Chrysoperla carnea (which<br />
consume eggs and larvae) adults and nymphs <strong>of</strong> the bugs Geocoris pallens<br />
and Orius tristicolor (which prey upon eggs and small larvae) and adults and<br />
nymphs <strong>of</strong> the bug Nabis americ<strong>of</strong>erus (which prey upon larvae <strong>of</strong> all sizes).<br />
Spiders, mantids, carabids, vespids and reduviid bugs are among predators<br />
that are also present in smaller numbers (Ehler and van den Bosch 1974;<br />
Ehler 1977a).<br />
Eleven species <strong>of</strong> parasitoid have been reported from T. ni on cotton, <strong>of</strong><br />
which the following are most important, although their combined effect is<br />
far less than that <strong>of</strong> the predators: Trichogramma semifumatum (an egg<br />
parasitoid), Cotesia marginiventris, Hyposoter exiguae and Microgaster<br />
brassicae (which attack small larvae and kill hosts when <strong>of</strong> medium size),<br />
Copidosoma truncatellum (an egg-larval parasitoid, which emerges from<br />
large larvae or prepupae), Chelonus texanus (an egg-larval parasitoid which<br />
kills medium sized hosts), Voria ruralis (which lays eggs on medium sized<br />
larvae and kills large larvae or prepupae), and Patrocloides montanus (a<br />
larval-pupal parasitoid which usually oviposits in large larvae). Copidosoma<br />
truncatellum is polyembryonic and Voria ruralis is <strong>of</strong>ten gregarious (Ehler<br />
and van den Bosch 1974).<br />
A nuclear polyhedrosis virus was the only pathogen shown to cause T. ni<br />
mortality, particularly late in the season at peak density <strong>of</strong> T. ni larvae, when<br />
levels <strong>of</strong> 50 to 60% mortality have been observed.<br />
All <strong>of</strong> the predators mentioned above are widely polyphagous. The first 4<br />
parasitoids listed above had a restricted host range and the last 4 were host<br />
specific in the cotton ecosystem.<br />
Disappearance <strong>of</strong> eggs and small larvae, assumed to be due to predation,<br />
was consistently the major mortality factor. Parasitisation <strong>of</strong> larvae by any<br />
parasitoid seldom exceeded 30%, the exception being that by Copidosoma<br />
truncatellum which <strong>of</strong>ten reached 50 to 75%. Detailed life table studies <strong>of</strong><br />
T. ni on cotton were reported by Ehler (1977a).
4.16 Trichoplusia ni 341<br />
It was suggested that the temporary nature <strong>of</strong> the cotton crop, and<br />
sufficient time each season for only 3 generations <strong>of</strong> the host, left parasitoids<br />
insufficient time to build up adequate numbers to thoroughly exploit T. ni<br />
populations. Also, the low density <strong>of</strong> T. ni due to the intense activity <strong>of</strong><br />
predators early in each generation, impairs successful search by adult<br />
parasitoids, particularly those that are host specific (Ehler and van den<br />
Bosch 1974; Ehler 1977a).<br />
If the reader is perhaps, somewhat uncertain <strong>of</strong> what conclusions to draw<br />
from the extensive data in the foregoing accounts, the studies <strong>of</strong> Jones et al.<br />
(1983a), dealing with the impact <strong>of</strong> parasitoids and predators on T. ni<br />
populations on celery in California, provide valuable insights. He concludes<br />
that naturally-occuring entomophagous arthropods do, indeed cause<br />
irreplaceable mortality <strong>of</strong> T. ni and that they should be considered a key part<br />
<strong>of</strong> any Integrated Pest Management program for the crop. Although<br />
parasitoids (principally Trichogramma spp., Copidosoma truncatellum and<br />
Voria ruralis, but also Hyposoter exiguae, Microgaster brassicae, Cotesia<br />
marginiventris and Chelonus insularis) can explain, for this crop, most<br />
mortality <strong>of</strong> eggs and <strong>of</strong> both small and medium sized larvae, it should not be<br />
concluded that predators are unimportant. It is possible that the additional<br />
small amount <strong>of</strong> mortality due to parasitoids is that required to suppress pest<br />
density to just below damaging levels. The parasitoids involved have a more<br />
restricted host range than the 2 most important groups <strong>of</strong> egg predators in<br />
celery, namely Coccinellidae (Hippodamia convergens and Cycloneda<br />
sanguinea) and Anthocoridae (Orius tristicolor) which are both widely<br />
polyphagous.<br />
Major parasitoid species<br />
Laboratory and field studies have been published dealing with many <strong>of</strong> the<br />
parasitoids listed in Table 4.16.1. Several <strong>of</strong> these species that have emerged<br />
as worthy <strong>of</strong> serious consideration as biological control agents are dealt with<br />
below.<br />
Copidosoma truncatellum Hym.: Encyrtidae<br />
Females <strong>of</strong> this small wasp oviposit in T. ni eggs <strong>of</strong> all ages and<br />
polyembryonic development occurs after hatching <strong>of</strong> the host larva. The host<br />
is later killed in the mature larval or prepupal stage. Either one or two eggs<br />
are inserted in a host egg during oviposition but, if the latter, generally only<br />
one parasitoid egg is fertile. Offspring are unisexual although, when 2 fertile<br />
eggs are laid, both males and females may emerge, an average <strong>of</strong> 1526 wasps<br />
per parasitised T. ni (Leiby 1926, 1929).
342 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Young female wasps are better than old at searching for eggs. At 14.8¡C<br />
and 28.9¡C the period from egg to first adult emergence was 122.9 and 22.4<br />
days respectively and the duration <strong>of</strong> a generation was 162.7 and 31.2 days.<br />
The life span <strong>of</strong> a female wasp fed on a diet <strong>of</strong> 20% levulose solution was<br />
30.3 days at 14.8¡C and 28 days at 35.6¡C. Synchronisation <strong>of</strong> the parasite to<br />
T. ni was found to be nearly perfect at 25¡C. At some temperatures the<br />
parasitoid killed host eggs and at others, larvae before the 5th instar: this<br />
resulted in the death <strong>of</strong> the contained parasitoids (Stoner and Weeks 1974,<br />
1976).<br />
T. ni larvae parasitised by C. truncatellum consumed 35% more food and<br />
had a 30% higher maximum weight than unparasitised larvae, which raises a<br />
concern for at least the short-term effect <strong>of</strong> biological control (Hunter and<br />
Stoner 1975). The mortality caused by C. truncatellum appeared to be<br />
density related (Ehler and van den Bosch 1974). Average parasitisation <strong>of</strong><br />
T. ni eggs in the laboratory was 55.3% (McPherson 1993). However, the<br />
maximum recorded in cotton fields in southern California was 2.5% (Ehler<br />
1977a).<br />
C. truncatellum has been reported from larvae <strong>of</strong> Noctuidae,<br />
Geometridae, Cossidae and Coleophoridae (Peck 1963). However, Ehler<br />
(1977a) points out that it has been reported only from T. ni in Californian<br />
cotton (van den Bosch and Hagen 1966; Ehler and van den Bosch 1974) and<br />
that it appears to be specific in this environment. C. truncatellum has a<br />
Holarctic distribution, but its native home is not clear (Peck 1963).<br />
Cotesia marginiventris Hym.: Braconidae<br />
More eggs were laid by this generalist larval parasitoid in 2-day-old T. ni<br />
larvae than in younger or older larvae. The minimum development period<br />
from oviposition to adult emergence from the host was 6 days (Boling and<br />
Pitre 1970). Females were significantly more responsive to host odors after<br />
brief contact with host larval frass or host-damaged cotton leaves (Turlings<br />
et al. 1989). C. marginiventris from T. ni and 4 other species <strong>of</strong> noctuid<br />
larvae were found to contain a non-occluded, filamentous, baculo-like virus<br />
(Styer et al. 1987).<br />
Hyposoter exiguae Hym.: Ichneumonidae<br />
This solitary endoparasitoid is one <strong>of</strong> 3 main parasitoids <strong>of</strong> T. ni on cotton<br />
(Ehler 1977a) and other crops in California and has a modest ability to<br />
distinguish unparasitised from parasitised hosts (Beegle and Oatman 1975;<br />
Browning and Oatman 1984). The female prefers to oviposit in late 1st or<br />
2nd instar T. ni larvae, although all instars are acceptable. When early instars<br />
are chosen, the host larvae generally die during the 3rd or 4th instar (Ehler<br />
1977a). Parasites commencing their development in hosts 1-day-old took
4.16 Trichoplusia ni 343<br />
13.85 days for development, whereas those starting in 10-day-old larvae<br />
required only 7.4 days (Smilowitz and Iwantsch 1975; Jowyk and Smilowitz<br />
1978). The influence <strong>of</strong> temperature on development is discussed by<br />
Browning and Oatman (1981). Weight gain <strong>of</strong> T. ni larvae is severely<br />
depressed following parasitisation (Smilowitz and Iwantsch 1973; Iwantsch<br />
and Smilowitz 1975; Thompson 1982).<br />
Successful parasitisation <strong>of</strong> T. ni larvae was correlated with host age,<br />
ranging from 83% to 88% in 1st, 2nd and early 3rd instars and declining in<br />
older larvae to 27% in mid 5th instar. Females deposited an average <strong>of</strong> 2.3<br />
eggs in 1st instar and 1.3 eggs in 2nd instar larvae, superparasitisation<br />
declining in later instars (Smilowitz and Iwantsch 1975).<br />
As many parasitoid eggs were laid in virus-infected host larvae as in<br />
healthy larvae. Of those females that oviposited in infected hosts, 60%<br />
transmitted infective doses <strong>of</strong> virus to 6% <strong>of</strong> healthy hosts subsequently<br />
exposed to them. Of female parasitoids that developed in virus-infected hosts,<br />
90% transmitted infective doses to an average <strong>of</strong> 21% healthy host larvae<br />
exposed to them. T. ni larvae parasitised by H. exiguae required twice the<br />
dosage <strong>of</strong> virus for infection and the parasitoid completed development before<br />
the host larvae died (Beegle and Oatman 1974, 1975). Washed H. exiguae<br />
eggs do not develop to maturity on injection into T. ni larvae unless virus or<br />
fluid from the parasite oviduct is added (Vinson and Stoltz 1986). Because the<br />
effects <strong>of</strong> parasitisation by H. exiguae are observable within 24 hours <strong>of</strong><br />
oviposition and prior to hatching <strong>of</strong> the parasitoid, it is probable that the<br />
H. exiguae-associated virus, rather than the developing parasitoid itself, is<br />
responsible for the metabolic changes produced (Thompson 1986).<br />
Microgaster brassicae Hym.: Braconidae<br />
This solitary endoparasitoid is an important mortality factor <strong>of</strong> T. ni larvae<br />
on both cabbage and cotton. The female usually oviposits in 1st and 2nd<br />
instar host larvae and the fully-grown parasite larva leaves through the<br />
lateral abdominal wall <strong>of</strong> the medium-sized host larvae to spin a greenish or<br />
grayish cocoon. The host larva <strong>of</strong>ten survives for a few days after parasitoid<br />
emergence (Ehler 1977a). Duration <strong>of</strong> parasitoid development from egg to<br />
adult ranges from 17.7 days at 21.2¡C to 10.7 days at 32.2¡C. Adult<br />
longevity ranged from 55.5 days at 15.5¡C to 12.9 days at 32.2¡C for males<br />
and from 76.4 days to 13.7 days for females. Total progeny is largest at<br />
21.1¡C, averaging 73.2 <strong>of</strong>fspring per female. Rearing methods, mating,<br />
searching, ovipositional behaviour and interactions with other parasitoid<br />
species have been described (Browning and Oatman 1984, 1985).<br />
M. brassicae is native to North America and appears to be specific to<br />
T. ni in cotton, although it is also known from the alfalfa looper Autographa<br />
californica on lucerne (Ehler 1977a).
344 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Trichogramma minutum Hym.: Trichogrammatidae<br />
When attacking eggs <strong>of</strong> T. ni, females reared from T. ni eggs were more<br />
fecund than those from Sitotroga cerealella and searched over larger areas<br />
for hosts (Marston and Ertle 1973). At 27¡C and 50% RH T. minutum<br />
populations increased more rapidly than those <strong>of</strong> T. platneri. T. minutum<br />
does not feed from the host egg nor does it superparasitise eggs even when<br />
hosts are in short supply (Manweiler 1986).<br />
Trichogramma platneri Hym.: Trichogrammatidae<br />
After ovipositing in a host egg, females pierce it again and feed from<br />
exuding droplets <strong>of</strong> fluid. T. platneri superparasitised hosts when eggs were<br />
scarce. At about 27¡C and 50% RH, T. platneri populations increased more<br />
slowly than those <strong>of</strong> T. minutum (Manweiler 1986). The maximum number<br />
<strong>of</strong> progeny bred from a single T. ni egg was 3 and 73% <strong>of</strong> male progeny<br />
emerged from the first eggs exposed to a female. Honey was shown to<br />
increase parasitoid longevity (Hohmann et al. 1988a, b).<br />
Trichogramma pretiosum Hym.: Trichogrammatidae<br />
This species can be reared from egg to adult in vitro (H<strong>of</strong>fman et al. 1975).<br />
Females have a preference for young T. ni eggs, although eggs <strong>of</strong> all ages are<br />
accepted (Godin and Boivin 1994). A local Missouri, USA strain <strong>of</strong><br />
T. pretiosum successfully parasitised T. ni eggs in field experiments, large<br />
host eggs producing more adults than small ones and these adults were more<br />
fecund and active than those from small eggs (Boldt et al. 1973). A Texan<br />
strain was effective in the laboratory against T. ni eggs, but not in the field. It<br />
was able to develop in the same T. ni egg as Trichogramma evanescens if<br />
eggs <strong>of</strong> both parasitoids were deposited on the same day (Parker and Pinnell<br />
1972, 1974).<br />
In Texas, naturally occurring T. pretiosum assisted in controlling T. ni on<br />
cotton in field cages (Lingren et al. 1978). In southern California, average<br />
parasitisation <strong>of</strong> T. ni eggs ranged from 3 to 47% in tomatoes when releases<br />
were made at the rate <strong>of</strong> 200 000 to 318 000 adult wasps per 0.4 ha (Oatman<br />
and Platner 1978). In Florida 3 releases 3 days apart <strong>of</strong> T. pretiosum at about<br />
378 000/acre/release in a 1 acre field cage containing 7 crops resulted in<br />
substantial parasitisation <strong>of</strong> T. ni eggs and in suppression <strong>of</strong> larvae (Martin et<br />
al. 1976b). Laboratory and field cage studies with T. pretiosum were also<br />
carried out in California (Ashley et al. 1974) where female parasitoids<br />
produced from T. ni eggs were larger, more fecund and lived longer than<br />
those from artificial rearing hosts (Plodia interpunctella and Sitotroga<br />
cerealella) (Bai et al. 1992).
4.16 Trichoplusia ni 345<br />
Voria ruralis Dip.: Tachinidae<br />
Adults mate soon after eclosion and oviposition commences about 9 days<br />
later. Eggs laid on the host surface hatch within a minute and young larvae<br />
penetrate the cuticle and enter a muscle fibre. After about 3 days at 24¡C the<br />
larva pierces a hole in the dorsal wall <strong>of</strong> the host abdomen through which it<br />
inserts its posterior spiracles into the air. After rapid growth <strong>of</strong> the parasitoid,<br />
the host dies and the parasitoid larva pupates within the host integument<br />
(Thompson 1915; Brubaker 1968). When V. ruralis oviposits on 1st instars,<br />
development is slower and mortality higher than in later instars, except the<br />
late 5th instar. Development was rarely completed when eggs were laid on<br />
5th instars, unless they were laid on newly moulted individuals. Females laid<br />
an average <strong>of</strong> 310 eggs (Elsey and Rabb 1970a). Development time from egg<br />
to puparium ranged from 5.4 to 12 days, depending upon the temperature,<br />
and for the pupa 7 to 8 days at 24¡C. Time from egg to adult varied from 19.4<br />
days at 20¡C to 10.7 days at 30¡C (Brubaker 1968; Jackson et al. 1969).<br />
Parasitisation by V. ruralis causes large larvae to eat less than normal (an<br />
average <strong>of</strong> 47% reduction (Soo Hoo and Seay 1972). Up to 85%<br />
parasitisation was observed in field cages, depending upon the numbers <strong>of</strong><br />
mated V. ruralis released, with significant superparasitism at high parasitoid<br />
densities (Soo Hoo et al. 1974).<br />
V. ruralis is one <strong>of</strong> 3 major parasitoids <strong>of</strong> T. ni on crops in Florida (Martin<br />
et al. 1982) and cotton in Arizona (Werner and Butler 1979) but was present<br />
only to the extent <strong>of</strong> 0 to 0.1% in larvae on lucerne in New Mexico (Gordon<br />
et al. 1987). In northwestern USA, it was the only parasitoid recovered and<br />
occurred in 0 to 14% <strong>of</strong> T. ni larvae (Biever et al. 1992). In Virginia,<br />
V. ruralis was present in 27% <strong>of</strong> larvae in 1981 and 17% in 1982<br />
(Chamberlin and Kok 1986).<br />
V. ruralis can survive, develop in, and emerge from virus infected larvae.<br />
However, it does not act as a vector, except occasionally as a mechanical one<br />
under very restricted conditions (Vail 1981).<br />
V. ruralis is a widespread parasite and has been recorded as far north as<br />
Finland and as far south as Trinidad. It has been recorded from a range <strong>of</strong><br />
Lepidoptera. In the United States it is known mainly from larvae <strong>of</strong> various<br />
Noctuidae, especially T. ni, but less frequently from the beet armyworm,<br />
Spodoptera exigua, and other associated species (Jackson et al. 1969; Ehler<br />
1977a). Ehler and van den Bosch (1974) considered V. ruralis to be host<br />
specific to T. ni in Californian cotton.
346 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Comment<br />
It has not been feasible, except for a rather more comprehensive cover <strong>of</strong><br />
parasitoids, to include any but the most relevant <strong>of</strong> the 2000 or so references<br />
to T. ni in the literature. Further details can be accessed via the bibliographies<br />
<strong>of</strong> Sutherland and Sutherland (1972, 1984) for earlier publications and via<br />
Commonwealth Agricultural Bureau Abstracts and Lingren and Green<br />
(1984) for much <strong>of</strong> the more recent literature.<br />
Although T. ni is <strong>of</strong>ten regarded as a secondary pest <strong>of</strong> crops in its native<br />
North America, damaging numbers, nevertheless, occur from time to time,<br />
particularly when its natural enemies are killed or suppressed by broad<br />
spectrum insecticides applied for associated primary pests. Predators are<br />
<strong>of</strong>ten claimed to be more important than parasitoids in maintaining T. ni at<br />
sub-economic levels.<br />
Regrettably most, if not all, <strong>of</strong> the major predators involved lack the<br />
degree <strong>of</strong> specificity nowadays considered necessary for introduction as<br />
classical biological control agents. For this reason, further consideration is<br />
restricted to the potential <strong>of</strong> parasitoids and viruses. Far more is known from<br />
southern California than elsewhere <strong>of</strong> the parasitoid species present and their<br />
interactions. The following discussion is thus somewhat geographically<br />
biased and it should be borne in mind that additional species in other regions<br />
may well have desirable characteristics, especially for their respective<br />
climatic conditions (see later).<br />
In addition to the egg parasitoids (e.g. Trichogramma pretiosum and<br />
T. platneri), there are at least 4 other parasitoids worthy <strong>of</strong> serious<br />
consideration (Copidosoma truncatellum, Hyposoter exiguae, Microgaster<br />
brassicae (Hymenoptera) and Voria ruralis (Tachinidae)).<br />
If host specificity considerations permit clearance <strong>of</strong> these species for<br />
introduction to a new area, a decision must still be taken on which, if not all,<br />
to establish. It may be useful, therefore, to review (and extend) the<br />
information presented earlier on their attributes and interactions.<br />
Copidosoma truncatellum oviposits into the host egg, but hatches in the<br />
larva and takes about 36 days to develop to adult, so it is present throughout<br />
the entire larval period <strong>of</strong> its host. Microgaster brassicae oviposits in 1st and<br />
2nd instar T. ni larvae and requires about 14 days to develop to adult. The<br />
mature 3rd instar parasitoid larva emerges from late 3rd or early 4th instar<br />
hosts after feeding for about 9 days. Hyposoter exiguae commonly oviposits<br />
into late 1st instar T. ni and emerges from late 3rd or early 4th instar hosts. It<br />
requires about 16 days from egg to adult at 25¡C, the egg-larval period<br />
averaging about 10 days. Voria ruralis requires about 13 days from egg to<br />
adult. It will oviposit on all host instars, but development in most successful
4.16 Trichoplusia ni 347<br />
when 2nd or 3rd instars are parasitised. Larval development is then<br />
completed by the end <strong>of</strong> the 5th host larval instar (Browning and Oatman<br />
1984). As a result <strong>of</strong> overlapping life cycles, two or more <strong>of</strong> these 4 species<br />
could inhabit a host larvae simultaneously, unless a species was able to<br />
discriminate between parasitised and unparasitised hosts.<br />
As many as 10 adult Voria ruralis may emerge from a single host larva.<br />
When limited numbers <strong>of</strong> hosts are present, a female lays more than one egg<br />
on each larva. Excess eggs may result in premature mortality <strong>of</strong> the host<br />
larva and any immature parasitoids already within it. External oviposition<br />
probably prevents the ovipositing female from receiving sensory<br />
information about the presence <strong>of</strong> parasite eggs or larvae already within the<br />
host; and V. ruralis females will continue to oviposit as long as the host larva<br />
reacts with any movement.<br />
Copidosoma truncatellum parasitises host eggs <strong>of</strong> all ages following<br />
antennal drumming and ovipositor insertion. Females are apparently able to<br />
discriminate between unparasitised eggs and those parasitised by other<br />
C. truncatellum females. Microgaster brassicae females insert their<br />
ovipositors in all host larvae whether or not parasitised by Copidosoma<br />
truncatellum, although they are weakly deterred from doing so in larvae<br />
already parasitised by other M. brassicae females or by Hyposoter exiguae.<br />
Parasitoid eggs were deposited in 91% <strong>of</strong> hosts previously unparasitised,<br />
whereas those already parasitised by M. brassicae or H. exiguae showed<br />
oviposition levels <strong>of</strong> 10 and 53% respectively, indicating a response to<br />
sensory information after insertion <strong>of</strong> the ovipositor. When M. brassicae<br />
oviposited in larvae already parasitised by C. truncatellum, the latter<br />
emerged from 77.5% <strong>of</strong> the larvae, whereas M. brassicae emerged from only<br />
12.5%. However, when M. brassicae oviposited in larvae containing the<br />
slower-developing Hyposoter exiguae, the latter emerged from 16.7% <strong>of</strong><br />
larvae and M. brassicae from 76.7%. Hyposoter exiguae showed little<br />
discrimination between unparasitised larvae and those parasitised by any <strong>of</strong><br />
the other 3 species, although ovipositor insertion did not result in additional<br />
eggs in larvae already containing H. exiguae. High levels <strong>of</strong> parasitisation by<br />
H. exiguae occurred in host larvae already parasitised by C. truncatellum,<br />
possibly due to the delayed development <strong>of</strong> the latter. Nevertheless, only<br />
C. truncatellum emerged from such larvae. Further details <strong>of</strong> these and other<br />
interactions under laboratory conditions are given in the valuable paper by<br />
Browning and Oatman (1984). It is interesting that the parasitoid complex<br />
attacking T. ni on cotton in the field shows little change from season to<br />
season in species composition and relative importance. However, this does<br />
not enable a simple decision to be made on what impact there would be on<br />
abundance <strong>of</strong> T. ni (or on plant damage sustained) if one or more <strong>of</strong> the
348 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
species was omitted from an introduction program. This applies particularly<br />
to C. truncatellum, with its feature <strong>of</strong> prolonging the feeding period <strong>of</strong><br />
parasitised larvae.<br />
A further factor to be taken into consideration is the crop or range <strong>of</strong><br />
crops and the climatic conditions under which T. ni control is particularly<br />
desired, since the effectiveness <strong>of</strong> various parasitoid species is affected by<br />
both sets <strong>of</strong> factors.<br />
Relevant to this statement are studies in northwest Florida, which<br />
reported that Cotesia autographae, Cotesia marginiventris and Meteorus<br />
autographae caused high T. ni mortality during spring and early summer in a<br />
mixed cropping system (Martin et al. 1982), and work in southern Florida<br />
reporting high mortality by Diadegma insulare <strong>of</strong> T. ni larvae on brassicas.<br />
Furthermore, Stenichneumon culpator cincticornis and Vulgichneumon<br />
brevicinctor appear important in New York State (Sutherland 1966),<br />
Brachymeria ovata in North Carolina (Elsey and Rabb 1970b), and<br />
Achaetoneura archippivora and Eucelatoria armigera in western United<br />
States (Martin et al. 1984).<br />
This suggests that some parasitoid species that are not widely distributed<br />
can act as significant mortality factors under certain climatic or crop<br />
conditions. They might well have a very restricted host range, and would be<br />
available for consideration, if required.
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6 Index <strong>of</strong> scientific names <strong>of</strong> insects<br />
Abacetus optimus Peringuey Col.: Carabidae 90<br />
abdominale, Dactylosternum<br />
abdominalis, Aphelinus<br />
abjectella, Drosica<br />
abjectum, Praon<br />
abnormis, Leptomastidea<br />
abrupta, Platysoma<br />
absinthii, Aphidius<br />
absinthii, Praon<br />
acaenovinae, Aphis<br />
acalephae, Trioxys<br />
acantha, Pediobius<br />
Acanthocoris concoloratus Uhler Hem.: Pentatomidae 233<br />
Acaulona brasiliana Townsend Dip.: Tachinidae 139<br />
achaeae, Trichogramma<br />
Achaetoneura archippivora (Williston) Dip.: Tachinidae 327, 348<br />
achaetoneura, Lespesia<br />
Achrysocharis Hym.: Eulophidae 251<br />
Achrysocharis douglasi,<br />
see Chrysonotomyia douglasi 251<br />
Achrysocharoides Hym.: Eulophidae 266, 275, 277<br />
Achrysophagus Hym.: Encyrtidae 294, 297<br />
acrobates, Telenomus<br />
Acroclissoides Hym.: Pteromalidae 201, 228<br />
Acrosternum Hem.: Pentatomidae 208<br />
Acrosternum aseadum Rolston Hem.: Pentatomidae 208<br />
Acrosternum gramineum (Fabricius) Hem.: Pentatomidae 205<br />
Acrosternum hilare (Say) Hem.: Pentatomidae 205, 208, 230<br />
Acrosternum marginatum Palisot de Beauvois Hem.: Pentatomidae 208<br />
Acrosternum pennsylvanicum Palisot de Beauvois Hem.: Pentatomidae 208<br />
acuminatus, Aenasius<br />
acuta, Chrysodeixis<br />
Adialytus salicaphis (Fitch) Hym.: Aphidiidae 37, 66<br />
Adonia variegata (Goeze) Col.: Coccinellidae 43<br />
aegyptiacus, Prochiloneurus<br />
aegyptius, Ischiodon<br />
477
478 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
aegyptius, Xanthogramma<br />
Aenasius Hym.: Encyrtidae 149<br />
Aenasius acuminatus Kerrich Hym.: Encyrtidae 147<br />
Aenasius colombiensis Compere Hym.: Encyrtidae 149<br />
Aenasius theobromae Kerrich Hym.: Encyrtidae 147<br />
aequalis, Pimpla<br />
affine, Calosoma<br />
affinis, Cryptolaemus<br />
africanus, Mesochorus<br />
africanus, Stictopisthus<br />
africanus, Syrphophagus<br />
Ageniaspis Hym.: Encyrtidae 123, 125, 265<br />
Ageniaspis citricola Logvinovskaya Hym.: Encyrtidae 257, 265, 272Ð280, 282, 285<br />
Agonoscelis rutila (Fabricius) Hem.: Pentatomidae 218<br />
agraules, Pnigalio<br />
agrestoria, Echthromorpha<br />
agriae, Trichogramma<br />
Agrius convolvuli (Linnaeus) Lep.: Sphingidae 1, 2, 9Ð16<br />
Agromyza Dip.: Agromyzidae 236<br />
Agromyza obtusa,<br />
see Melanagromyza obtusa 236<br />
Agromyza phaseoli,<br />
see Ophiomyia phaseoli 236<br />
agromyzae, Neodimmockia<br />
agromyzae, Sphegigaster<br />
agromyzae, Trigonogastra<br />
alaskensis, Microplitis<br />
albiceps, Phytomyza<br />
albicollis, Hyperaspis<br />
albipes, Pleurotropitiella<br />
albizonalis, Autocharis<br />
albizonalis, Deanolis<br />
albizonalis, Noorda<br />
Alcaeorrhynchus grandis (Dallas) Col.: Pentatomidae 330<br />
Aleiodes Hym.: Braconidae 21, 23, 28<br />
Aleiodes aligharensi (Quadri) Hym.: Braconidae 21, 23, 28<br />
aliberti, Brachymeria<br />
aligarhensis, Diaphorencyrtus<br />
aligharensi, Aleiodes<br />
Allograpta nasuta (Macquart) Dip.: Syrphidae 43
Allograpta pfeifferi,<br />
see Allograpta nasuta<br />
Allotropa Hym.: Platygasteridae 150, 301<br />
Scientific Index 479<br />
Allotropa citri Muesebeck Hym.: Platygasteridae 295, 298Ð301, 307, 315<br />
Allotropa kamburovi Annecke and Prinsloo Hym.: Platygasteridae 295, 298<br />
Allotropa mecrida (Walker) Hym.: Platygasteridae 295, 298, 306<br />
Alloxysta Hym.: Charipidae 65, 67, 68, 72<br />
Alloxysta brevis (Thompson) Hym.: Charipidae 72<br />
43<br />
Alloxysta darci (Girault) Hym.: Charipidae 72<br />
aloysiisabaudiae, Trissolcus<br />
alternatus, Nabis<br />
alticeps, Leucopis<br />
amabilis, Eublemma<br />
Amatellon Hym.: Eulophidae 266, 273, 282<br />
ambiguus, Lipolexis<br />
ambiguus, Lysiphlebus<br />
Amblypelta cocophaga China Hem.: Coreidae 226<br />
Amblyteles fuscipennis Wesmael Hym.: Ichneumonidae 12<br />
americana, Asaphes<br />
americana, Carcinophora<br />
americana, Psalis<br />
americ<strong>of</strong>erus, Nabis<br />
Amobia Dip.: Sarcophagidae 190<br />
amygdali, Hyalopterus<br />
amyoti, Glaucias<br />
Anacanthocoris concoloratus (Uhler) Hem.: Coreidae 224<br />
Anagyrus Hym.: Encyrtidae 147, 152<br />
Anagyrus ananatis Gahan Hym.: Encyrtidae 145, 147, 149, 152, 154Ð156<br />
Anagyrus bohemani (Westwood) Hym.: Encyrtidae 294, 297, 305<br />
Anagyrus coccidivorus,<br />
see Anagyrus ananatis 149<br />
Anagyrus greeni Howard Hym.: Encyrtidae 294, 297<br />
Anagyrus kivuensis Compere Hym.: Encyrtidae 149, 307<br />
Anagyrus pseudococci (Girault) Hym.: Encyrtidae 287, 297, 299Ð301, 303, 305Ð307,<br />
311, 315, 316<br />
Anagyrus sawadai Ishii Hym.: Encyrtidae 294, 297, 305, 307<br />
analis, Lebia<br />
ananatis, Anagyrus<br />
Anasa tristis (De Geer) Hem.: Pentatomidae 230<br />
Anastatus Hym: Eupelmidae 203, 211, 214, 215, 223, 227
480 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Anastatus bifasciatus Boyer de Fonscolombe Hym.: Eupelmidae 203, 223<br />
Anastatus dasyni Ferri re Hym: Eupelmidae 202<br />
Anastatus japonicus Ashmead Hym: Eupelmidae 203, 225<br />
anchorago, Stiretrus<br />
angelicae, Trioxys<br />
angelicus, Pseudaphycus<br />
Angita insularis Hym.: Ichneumonidae 324<br />
Angitia plutellae,<br />
see Diadegma insulare<br />
angolensis, Chilocorus<br />
angustifrons, Pseudaphycus<br />
Anisochrysa basalis,<br />
see Mallada basalis 291<br />
Anisolabis annulipes, see Euborellia annulipes 90<br />
Ankylopteryx octopunctata Fabricius Neu.: Chrysopidae 271<br />
annulipes, Anisolabis<br />
annulipes, Euborellia<br />
Anochaetus Hym.: Formicidae 92<br />
anomidis, Apanteles<br />
Anomis erosa (HŸbner) Lep.: Noctuidae 18<br />
Anomis flava (Fabricius) Lep.: Noctuidae 1, 2, 17Ð31<br />
Anoplolepis custodiens (F. Smith) Hym.: Formicidae 305<br />
Anoplolepis longipes (Jerdon) Hym.: Formicidae 155<br />
antennata, Nezara<br />
antennata, Pauesia<br />
antestiae, Gryon<br />
antestiae, Hadronotus<br />
Antestia orbana Kirk Hem.: Pentatomidae 225<br />
Anthocoris Hem.: Anthocoridae 43<br />
Antilochus coquebertii (Fabricius) Hem.: Pyrrhocoridae 137, 138<br />
antinorii, Bogosia<br />
Antonina graminis (Maskell) Hem.: Pseudococcidae 155<br />
apachus, Polistes<br />
Apanteles Hym.: Braconidae 12, 14, 23, 29, 90, 192<br />
Apanteles anomidis Watanabe Hym.: Braconidae 21, 23, 29, 31<br />
Apanteles ruficrus,<br />
see Cotesia ruficrus 23<br />
Apanteles syleptae Ferri re Hym.: Braconidae 23, 28<br />
apetzi, Scymnus<br />
Aphanogmus dictynna (Waterston) Hym.: Ceraphronidae 166, 168, 182<br />
Aphelinus Hym.: Aphelinidae 37, 51, 63, 64, 68, 72, 82, 83
Scientific Index 481<br />
Aphelinus abdominalis (Dalmer) Hym.: Aphelinidae 37, 50, 61, 65, 67, 68, 71<br />
Aphelinus asychis Walker Hym.: Aphelinidae 50<br />
Aphelinus basalis, see Aphelinus abdominalis<br />
Aphelinus brevis, see Alloxysta brevis<br />
37, 67<br />
Aphelinus chaoniae Walker Hym.: Aphelinidae 50<br />
Aphelinus flavipes, see Aphelinus abdominalis 37, 50, 65, 71<br />
Aphelinus gossypii Timberlake Hym.: Aphelinidae 37, 50, 68, 71, 72, 83<br />
Aphelinus humilis Mercet Hym.: Aphelinidae 50<br />
Aphelinus kashmiriensis, see Aphelinus gossypii 50<br />
Aphelinus mali (Haldeman) Hym.: Braconidae 50, 63, 68, 72<br />
Aphelinus mariscusae (Risbec) Hym.: Aphelinidae 37<br />
Aphelinus nigritus, see Aphelinus varipes 50<br />
Aphelinus paramali Zehavi and Rosen Hym.: Aphelinidae 50<br />
Aphelinus semiflavus Howard Hym.: Aphelinidae 56, 69, 72<br />
Aphelinus varipes (Foerster) Hym.: Aphelinidae 50, 61<br />
Aphidencyrtus Hym.: Encyrtidae 57, 63, 65<br />
Aphidencyrtus aligarhensis, see Diaphorencyrtus aligarhensis 121, 131<br />
Aphidencyrtus aphidiphagus, see Syrphophagus aphidivora 72<br />
Aphidencyrtus aphidivora, see Syrphophagus aphidivora 42<br />
Aphidencyrtus diaphorinae, see Diaphorencyrtus aligarhensis<br />
aphidimyza, Aphidoletes<br />
aphidiphagus, Aphidencyrtus<br />
aphidis, Pachyneuron<br />
121<br />
Aphidius Hym.: Aphidiidae 38, 53<br />
Aphidius absinthii Marshall Hym.: Aphidiidae 37, 67<br />
Aphidius avenae, see Aphidius picipes 63<br />
Aphidius cardui, see Lysiphlebus fabarum 39<br />
Aphidius colemani Viereck Hym.: Aphidiidae 37, 51, 60, 61, 64, 66, 68, 73, 81Ð83<br />
Aphidius ervi Haliday Hym.: Aphidiidae 37, 51<br />
Aphidius flavipes, see Aphelinus abdominalis 37<br />
Aphidius floridaensis Smith Hym.: Aphidiidae 51<br />
Aphidius funebris Mackauer Hym.: Aphidiidae 37<br />
Aphidius gifuensis Ashmead Hym.: Aphidiidae 51, 68, 74, 83<br />
Aphidius lonicerae, see Aphidius urticae 51<br />
Aphidius matricariae (Haliday) Hym.: Aphidiidae 5, 37, 51, 67, 74, 82<br />
Aphidius phorodontis, see Aphidius matricariae 51, 74<br />
Aphidius picipes (Nees) Hym.: Aphelinidae 51, 63<br />
Aphidius platensis, see Aphidius colemani 37, 73
482 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Aphidius ribis Haliday Hym.: Aphidiidae 37<br />
Aphidius rosae Haliday Hym.: Aphidiidae 38<br />
Aphidius salicis Haliday Hym.: Aphidiidae 38<br />
Aphidius similis Starù and Carver Hym.: Aphidiidae 38, 51<br />
Aphidius sonchi Marshall Hym.: Aphidiidae 51<br />
Aphidius transcaspicus, see Aphidius colemani 73<br />
Aphidius urticae Haliday Hym.: Aphidiidae 51<br />
Aphidius uzbekistanicus Luzhetzki Hym.: Aphidiidae<br />
aphidivora, Aphidencyrtus<br />
aphidivora, Syrphophagus<br />
38, 51<br />
Aphidoletes aphidimyza (Rondani) Dip.: Cecidomyiidae 49, 57, 69, 79, 80<br />
Aphis Hem.: Aphididae 75, 76, 78, 79, 82<br />
Aphis acaenovinae Eastop Hem.: Aphididae 76<br />
Aphis citricola, see Aphis spiraecola 70, 78<br />
Aphis craccivora Koch Hem.: Aphididae 1, 2, 33Ð83<br />
Aphis fabae Scopoli Hem.: Aphididae 73, 75<br />
Aphis gossypii Glover Hem.: Aphididae 1, 2, 33Ð83<br />
Aphis nerii Boyer de Fonscolombe Hem.: Aphididae 60, 70, 73, 74, 76Ð78<br />
Aphis punicae Passerini Hem.: Aphididae 73<br />
Aphis spiraecola Patch Hem.: Aphididae 66, 70, 76, 78<br />
Aphis zizyphi Theobald Hem.: Aphididae<br />
apicalia, Exorista<br />
apiciflavus, Scymnus<br />
73<br />
Apleurotropis Hym.: Eulophidae 266, 273<br />
Aplomya theclarum (Scudder) Dip.: Tachinidae 327<br />
Aprostocetus Hym.: Eulophidae<br />
archippivora, Achaetoneura<br />
archippivora, Lespesia<br />
241, 266<br />
Archytas californiae (Walker) Dip.: Tachinidae<br />
arcuata, Coccinella<br />
arcuata, Ectophasiopsis<br />
argenteopilosus, Eriborus<br />
327<br />
Argyrophylax atropivora, see Zygobothria atropivora<br />
armigera, Eucelatoria<br />
armigera, Helicoverpa<br />
12<br />
Asaphes americana, see Asaphes lucens 72<br />
Asaphes lucens (Provancher) Hym.: Pteromalidae 72<br />
Asaphoideus niger Girault Hym.: Pteromalidae 271, 272
Ascotolinx funeralis Girault Hym.: Eulophidae 266, 272, 276<br />
aseadum, Acrosternum<br />
asiaticus, Trioxys<br />
Asolcus mitsukurii, see Trissolcus mitsukurii 224<br />
asychis, Aphelinus<br />
atomella, Melanagromyza<br />
atricornis, Phytomyza<br />
atropivora, Argyrophylax<br />
atropivora, Sturmia<br />
atropivora, Zygobothria<br />
attrisium, Rhychium<br />
auberti, Diaphorina<br />
auctus, Trioxys<br />
aurantii, Toxoptera<br />
auratocauda, Cadurcia<br />
auratocauda, Sturmia<br />
auropunctata, Wasmannia<br />
australicum, Trichogramma<br />
australiensis, Eupelmus<br />
Autocharis albizonalis, see Deanolis albizonalis 106<br />
Autographa brassicae, see Trichoplusia ni 318<br />
Autographa californica (Speyer) Lep.: Noctuidae 338, 342<br />
autographae, Cotesia<br />
autographae, Meteorus<br />
avenae, Aphidius<br />
Axiagastus campbelli Distant Hem.: Pentatomidae 226<br />
axyridis, Harmonia<br />
ayyari, Tetrastichus<br />
Azteca Hym.: Formicidae 176<br />
Bactrocera ferrugineus (Fabricius) Dip.: Tephritidae 109<br />
Bactrocera frauenfeldi (Schiner) Dip.: Tephritidae 109<br />
balteatus, Episyrphus<br />
barberi, Sympherobius<br />
Baryscapus galactopus (Ratzeberg) Hym.: Eulophidae 324<br />
basalis, Anisochrysa<br />
basalis, Aphelinus<br />
basalis, Chrysopa<br />
Scientific Index 483
484 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
basalis, Mallada<br />
basalis, Trissolcus<br />
basicurvus, Trioxys<br />
bella, Leucopis<br />
Belonuchus ferrugatus (Erichson) Col.: Staphylinidae 92, 93, 102<br />
Belonuchus quadratus Kraatz. Col.: Staphylinidae 92, 102<br />
beneficia, Polycystomyia<br />
Bessa remota (Aldrich) Dip.: Tachinidae 327<br />
bicarinativentris, Plutarchia<br />
bicarinatum, Tetramorium<br />
bicolor, Charops<br />
bicolor, Galerita<br />
bicolor, Microdon<br />
bicolor, Paragus<br />
bicolor, Propagalerita<br />
bifasciatus, Anastatus<br />
biguttatus, Scymnus<br />
bilobus, Zelus<br />
bilucenarius, Nephus<br />
bilucenarius, Scymnus<br />
bimaculata, Microcharops<br />
bimaculata, Sturmia<br />
binaevatus, Scymnus<br />
Biosteres Hym.: Braconidae 251<br />
biplaga, Earias<br />
bipunctatus, Nephus<br />
bipunctatus, Scymnus<br />
bipustulatus, Chilocorus<br />
blackburni, Chelonus<br />
blanda blanda, Zenilla<br />
Blepyrus insularis (Cameron) Hym.: Encyrtidae 294, 297<br />
Blepyrus saccharicola Gahan Hym.: Encyrtidae 294, 297, 300, 307<br />
Bogosia antinorii Rondani Dip.: Tachinidae 201, 202, 211, 217, 232Ð234<br />
Bogosia helva (Wiedermann) Dip.: Tachinidae 139<br />
bohemani, Anagyrus<br />
bolivari, Neoprochiloneurus<br />
boninensis, Chrysopa<br />
boninensis, Mallada
Scientific Index 485<br />
borbonicus, Paragus<br />
borellii, Labia<br />
Brachycantha Col.: Coccinellidae 146, 152<br />
Brachycaudus Hem.: Aphididae 75<br />
Brachycaudus cardui (Linnaeus) Hem.: Aphididae 75<br />
Brachymeria Hym.: Chalcididae 21, 24, 27, 191, 192, 337<br />
Brachymeria aliberti (Schmitz) Hym.: Chalcididae 24, 28<br />
Brachymeria intermedia (Nees) Hym.: Chalcididae 323<br />
Brachymeria lasus (Walker) Hym.: Chalcididae 24, 30, 191, 192, 323<br />
Brachymeria madecassa Steffan Hym.: Chalcididae 24, 29<br />
Brachymeria multicolor (Kieffer) Hym.: Chalcididae 21, 24, 29<br />
Brachymeria obscurata, see Brachymeria lasus 24, 192<br />
Brachymeria ovata (Say) Hym.: Chalcididae 323, 337, 348<br />
Brachymeria paolii Masi Hym.: Chalcididae 24, 29<br />
Brachymeria tibialis Steffan Hym.: Chalcididae 21, 24<br />
Bracon Hym.: Braconidae 190, 192, 265, 276, 282<br />
Bracon greeni Ashmead Hym.: Braconidae 190, 194<br />
Bracon phyllocnistidis (Muesebeck) Hym.: Braconidae 265, 274, 282<br />
brasiliana, Acaulona<br />
brassicae, Autographa<br />
brassicae, Brevicoryne<br />
brassicae, Liriomyza<br />
brassicae, Microgaster<br />
brassicae, Microplitis<br />
brevicapillum, Trichogramma<br />
brevicinctor, Vulgichneumon<br />
Brevicoryne brassicae (Linnaeus) Hem.: Aphididae 48<br />
brevipes, Dysmicoccus<br />
brevipes, Pseudococcus<br />
brevipetiolatus, Zaommomentedon<br />
brevipetioletus, Visnuella<br />
brevis, Alloxysta<br />
brevis, Aphelinus<br />
Brinckochrysa scelestes (Banks) Neu.: Chrysopidae 291<br />
brochymenae, Trissolcus<br />
bromeliae, Diaspis<br />
Brumoides lineatus (Weise) Col.: Coccinellidae 291<br />
Brumus suturalis (Fabricius) Col.: Coccinellidae 43, 291, 309
486 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
brunneicornis, Sphegigaster<br />
brunnipennis, Plautia<br />
c-nigrum, Hyperaspis<br />
cadaverica, Scleroderma<br />
Cadurcia auratocauda (Curran) Dip.: Tachinidae<br />
caenosus, Dictyotis<br />
californiae, Archytas<br />
californica, Autographa<br />
californica, Coccinella<br />
californicus, Ooencyrtus<br />
22<br />
Calliceras dictynna, see Aphanogmus dictynna 168<br />
Calliephaltes Hym.: Ichneumonidae 195<br />
Callitula filicornis Del. Hym.: Pteromalidae 243<br />
Callitula viridicoxa (Girault) Hym.: Eurytomidae 243, 251<br />
Callitula yasudi Yasuda Hym.: Pteromalidae 243<br />
Calosoma affine Chaudoir Col.: Carabidae 332<br />
Calosoma peregrinator GuŽrin Col.: Carabidae 332, 340<br />
Calosoma schayeri Erichson Col.: Carabidae<br />
campaniformis, Eumenes<br />
campbelli, Axiagastus<br />
26<br />
Camplyocheta Dip.: Tachinidae 22<br />
Campoletis Hym.: Ichneumonidae 324<br />
Campoletis flavicincta Ashmead Hym.: Ichneumonidae 324<br />
Campoletis sonorensis (Cameron) Hym.: Ichneumonidae 324<br />
Campoletis websteri, see Campoletis sonorensis 324<br />
Camponotus Hym.: Formicidae 153<br />
Camponotus friedae Forel Hym.: Formicidae 155<br />
Campyloneura Hym.: Braconidae 190, 192<br />
Cantheconidia furcellata, see Eucanthecona furcellata<br />
capensis, Condica<br />
capensis, Prospalta<br />
capicola, Platynaspis<br />
22<br />
Carcelia Dip.: Tachinidae 111, 327<br />
Carcelia cosmophilae (Curran) Dip.: Tachinidae 22, 27<br />
Carcelia illota (Curran) Dip.: Tachinidae 22, 27<br />
Carcelia kockiana Tours. Dip.: Tachinidae 22<br />
Carcinophora americana (Palisot de Beauvois) Derm.: Labiidae 90, 102
Scientific Index 487<br />
cardiae, Diaphorina<br />
Cardiochiles nigriceps Viereck Hym.: Braconidae<br />
cardui, Aphidius<br />
cardui, Brachycaudus<br />
cardui, Lysiphlebus<br />
carnea, Chrysopa<br />
carnea, Chrysoperla<br />
322<br />
Carpocoris mediterraneus Tamanini Hem.: Pentatomidae 224<br />
Carpophilus Col.: Nitidulidae 144<br />
Casinaria infesta (Cresson) Hym.: Ichneumonidae 324<br />
Cathartus Col.: Silvanidae<br />
centaureae, Trioxys<br />
centrosematis, Ophiomyia<br />
92, 93, 102<br />
Cephalonomia Hym.: Bethylidae 175<br />
Cephalonomia stephanoderis Betrem Hym.: Bethylidae 157, 166, 168, 172Ð177, 182<br />
Ceraphron dictynna, see Aphanogmus dictynna<br />
cerasicola, Ephedrus<br />
182<br />
Ceratomegilla maculata fuscilabris De Geer Col.: Coccinellidae<br />
cerealella, Sitotroga<br />
332<br />
Cermatulus nasalis (Westwood) Hem.: Pentatomidae<br />
ceroplastae, Coccophagus<br />
chaoniae, Aphelinus<br />
22, 27, 218, 225<br />
Charichirus Col.: Staphylinidae 92<br />
Charips, see Alloxysta 65<br />
Charops Hym.: Ichneumonidae 25<br />
Charops bicolor (SzŽpligeti) Hym.: Ichneumonidae 12, 14, 21, 25, 29<br />
Chartocerus walkeri Hayat Hym.: Signiphoridae 120, 123, 125<br />
Cheilomenes lunata (Fabricius) Col.: Coccinellidae 43<br />
Cheilomenes sexmaculata Fabricius Col.: Coccinellidae 43, 65, 67, 122, 128<br />
Cheilomenes sulphurea (Oliver) Col.: Coccinellidae 43<br />
Cheilomenes vicina (Mulsant) Col.: Coccinellidae 43<br />
Cheiloneurus Hym.: Encyrtidae 123, 125, 129<br />
Chelonus Hym.: Braconidae 190, 192, 195, 322<br />
Chelonus blackburni Cameron Hym.: Braconidae 321, 322<br />
Chelonus curvimaculatus Cameron Hym.: Braconidae 322<br />
Chelonus formosanus Sonan Hym.: Braconidae 322<br />
Chelonus insularis (Cresson) Hym.: Braconidae 321, 322, 341<br />
Chelonus, texanus Cresson Hym.: Braconidae 340
488 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Chetogena Dip.: Tachinidae 327<br />
Chilo simplex, see Chilo suppressalis 15<br />
Chilo suppressalis Walker Lep.: Pyralidae 15<br />
Chilocorus angolensis Crotch Col.: Coccinellidae 309<br />
Chilocorus bipustulatus (Linnaeus) Col.: Coccinellidae<br />
chilonis, Trichogramma<br />
chilotraeae, Trichogramma<br />
292, 303<br />
Chlorocytus Hym.: Pteromalidae 242, 246<br />
Chloropulvinaria psidii (Maskell) Hem.: Coccidae<br />
chloropus, Telenomus<br />
314<br />
Chromatomyia horticola (Goureau) Dip.: Agromyzidae 239, 252<br />
Chrysocharis Hym.: Eulophidae 266, 273, 274, 276<br />
Chrysocharis pentheus (Walker) Hym.: Eulophidae 266, 274, 275<br />
Chrysodeixis acuta (Walker) Lep.: Noctuidae 28<br />
Chrysodeixis includens Walker Lep.: Noctuidae 340<br />
Chrysonotomyia Hym.: Eulophidae 241, 266, 273, 277<br />
Chrysonotomyia douglasi (Girault) Hym.: Eulophidae 241, 251<br />
Chrysonotomyia erythraea (Silvestri) Hym.: Eulophidae 241, 251<br />
Chrysonotomyia formosa (Westwood) Hym.: Eulophidae 241, 251<br />
Chrysopa Neu.: Chrysopidae 22, 146, 291, 331<br />
Chrysopa basalis, see Mallada basalis 271<br />
Chrysopa bipunctatus, see Nephus bipunctatus<br />
Chrysopa boninensis Okamoto Neu.: Chrysopidae<br />
Chrysopa carnea, see Chrysoperla carnea<br />
122, 271, 273, 282, 283<br />
Chrysopa formosa Brauer Neu.: Chrysopidae 62<br />
Chrysopa intima MacLachlan Neu.: Chrysopidae 62<br />
Chrysopa irregularis Banks Neu.: Chrysopidae 146, 152<br />
Chrysopa kulingensis Navas Neu.: Chrysopidae 190, 192<br />
Chrysopa lacciperda, see Odontochrysa lacciperda 291, 302, 314<br />
Chrysopa lanata Banks Neu.: Chrysopidae 331<br />
Chrysopa nigricornis Burmeister Neu.: Chrysopidae 331<br />
Chrysopa orestes Banks Neu.: Chrysopidae 65<br />
Chrysopa pallens Tieder Neu.: Chrysopidae 62<br />
Chrysopa perla (Linnaeus) Neu.: Chrysopidae 69<br />
Chrysopa ramburi Schneider Neu.: Chrysopidae 146, 152<br />
Chrysopa rufilabris Burmeister Neu.: Chrysopidae 331<br />
Chrysopa scelestes, see Brinckochrysa scelestes 291<br />
Chrysopa septempunctata, see Chrysopa pallens 62
Scientific Index 489<br />
Chrysopa sinica (Tieder) Neu.: Chrysopidae 62, 69, 271<br />
Chrysoperla carnea (Stephens) Neu.: Chrysopidae 43, 69, 291, 302, 303, 331, 340<br />
Chrysoperla sinica, see Chrysopa sinica 62<br />
Chrysopilus Dip.: Rhagionidae 92<br />
Chrysopilus ferruginosus (Wiedermann) Dip.: Rhagionidae 92, 93, 95, 96<br />
Chrysoplatycerus splendens (Howard) Hym.: Encyrtidae 294, 297, 305, 316<br />
ciliata, Zygobothria<br />
cinctipennis, Closterocerus<br />
cingulatus, Dysdercus<br />
circulus, Halticoptera<br />
Cirrospiloideus phyllocnistoides, see Citrostichus phyllocnistoides 268<br />
Cirrospilus Hym.: Eulophidae 102, 241, 266, 272, 273, 275, 278<br />
Cirrospilus ingenuus Gahan Hym.: Eulophidae 264, 266, 274, 275, 277, 283<br />
Cirrospilus longefasciatus Ferri re Hym.: Eulophidae 266<br />
Cirrospilus lyncus Walker Hym.: Eulophidae 266, 274<br />
Cirrospilus phyllocnistis Ishii Hym.: Eulophidae 266, 277<br />
Cirrospilus phyllocnistoides, see Citrostichus phyllocnistoides 268<br />
Cirrospilus pictus (Nees) Hym.: Eulophidae 266, 275, 276<br />
Cirrospilus quadristriatus (Subba Rao & Ramamani) Hym.: Eulophidae 257, 266, 272,<br />
273, 275, 277Ð280, 283, 285<br />
Cirrospilus variegatus (Masi) Hym.: Eulophidae 266, 277<br />
Cirrospilus vittatus (Walker) Hym.: Eulophidae 266, 275<br />
cirsii, Trioxys<br />
citrella, Phyllocnistis<br />
citri, Allotropa<br />
citri, Diaphorina<br />
citri, Kratoysma<br />
citri, Planococcus<br />
citricola, Ageniaspis<br />
citricola, Aphis<br />
citricola, Psylla<br />
citriculus, Pseudococcus<br />
citrisuga, Psylla<br />
citroimpura, Trioza<br />
Citrostichus phyllocnistoides (Narayanan) Hym.: Eulophidae 268, 272, 273, 275, 277,<br />
279, 280, 283, 285<br />
Clausenia josefi Rosen Hym.: Encyrtidae 294, 297<br />
Clausenia purpurea Ishii Hym.: Encyrtidae 303
490 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
clavata, Gymnosoma<br />
Cleodiplosis koebelei (Felt) Dip.: Cecidomyiidae 151, 154<br />
Cleothera Col.: Coccinellidae 150<br />
Closterocerus Hym.: Eulophidae 177, 273<br />
Closterocerus cinctipennis Ashmead Hym.: Eulophidae 268, 278<br />
Closterocerus trifasciatus Westwood Hym.: Eulophidae 268, 275, 277<br />
Coccodiplosis smithi (Felt) Dip.: Cecidomyiidae<br />
coccidivora, Triommata<br />
coccidivorus, Anagyrus<br />
293<br />
Coccidoxenoides Hym.: Encyrtidae 299, 301<br />
Coccidoxenoides peregrinus Timberlake Hym.: Encyrtidae<br />
304, 305, 307, 311, 312, 315, 316<br />
287, 294, 296, 297, 300Ð302,<br />
Coccinella arcuata, see Harmonia octomaculata 60, 300<br />
Coccinella californica Mann. Col.: Coccinellidae 292<br />
Coccinella novemnotata francisciana Casey Col.: Coccinellidae 332<br />
Coccinella repanda, see Coccinella transversalis 60, 292<br />
Coccinella septempunctata (Linnaeus) Col.: Coccinellidae 26, 43, 62, 65Ð67, 69, 306<br />
Coccinella semipunctata Col.: Coccinellidae 292<br />
Coccinella transversalis (Thunberg) Col.: Coccinellidae 60, 292<br />
Coccinella transversoguttata Faldeman Col.: Coccinellidae 332<br />
Coccinella undecimpunctata Linnaeus Col.: Coccinellidae<br />
coccophaga, Amblypelta<br />
69<br />
Coccophagus Hym.: Aphelinidae 123, 126<br />
Coccophagus ceroplastae (Howard) Hym.: Aphelinidae 123, 126<br />
Coccophagus heteropneusticus Compere Hym.: Aphelinidae 311<br />
Coccus pseudomagnoliarum (Kuwana) Hem.: Coccidae<br />
c<strong>of</strong>fea, Phymastichus<br />
c<strong>of</strong>feicola, Heterospilus<br />
cognatoria, Hadrojoppa<br />
colemani, Aphidius<br />
299<br />
Collops marginellus Le Conte Col.: Melyridae 332<br />
Collops vittatus (Say) Col.: Melyridae<br />
colombiensis, Aenasius<br />
communis, Trioxys<br />
comperei, Telenomus<br />
complanatus, Trioxys<br />
complexa, Sinea<br />
332<br />
Compsilura concinnata (Meigen) Dip.: Tachinidae 327
Scientific Index 491<br />
comstocki, Pseudococcus<br />
comstockii, Euplectrus<br />
concinnata, Compsilura<br />
concolor, Pachyneuron<br />
concoloratus, Anacanthocoris<br />
Condica capensis GuenŽe Lep.: Noctuidae 194<br />
confrater, Eupeodes<br />
confrater, Syrphus<br />
confusa, Sinea<br />
confusum, Trichogramma<br />
confusus, Lysiphlebus<br />
congregata, Cotesia<br />
Conocephalus saltator (Saussure) Ort.: Tettigoniidae 146<br />
conquisator, Itoplectis<br />
constrictus, Scymnus<br />
convergens, Hippodamia<br />
convolvuli, Agrius<br />
convolvuli, Herse<br />
Copidosoma Hym.: Encyrtidae 195, 324, 337<br />
Copidosoma floridanum (Ashmead) Hym.: Encyrtidae 324, 326, 337<br />
Copidosoma truncatellum Dalman Hym.: Encyrtidae 317, 321, 324, 335, 337Ð342,<br />
346Ð348<br />
coquebertii, Antilochus<br />
corbetti, Elasmus<br />
Coruna Hym.: Pteromalidae 218<br />
Cosmophila, see Anomis<br />
Cosmophila erosa, see Anomis erosa 18<br />
Cosmophila flava, see Anomis flava 18<br />
Cosmophila indica, see Anomis flava 18<br />
cosmophilae, Carcelia<br />
cosmophilae, Zenillia<br />
Cosmopolites Col.: Curculionidae 86<br />
Cosmopolites pruinosus Heller Col.: Curculionidae 86<br />
Cosmopolites sordidus (Germar) Col.: Curculionidae 1, 2, 85Ð104<br />
Cotesia Hym.: Braconidae 322, 336<br />
Cotesia autographae Muesebeck Hym.: Braconidae 322, 348<br />
Cotesia congregata (Say) Hym.: Braconidae 322<br />
Cotesia glomerata (Linnaeus) Hym.: Braconidae 322, 337
492 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Cotesia laeviceps (Ashmead) Hym.: Braconidae 322<br />
Cotesia marginiventris (Cresson) Hym.: Braconidae 322, 336, 340Ð342, 348<br />
Cotesia plutellae Kurdjimov Hym.: Braconidae 322, 336, 337<br />
Cotesia ruficrus (Haliday) Hym.: Braconidae 23, 322, 336<br />
Cotesia yakutatensis Ashmead Hym.: Braconidae<br />
craccivora, Aphis<br />
crassipennis, Ectophasia<br />
crawfordi, Ophelosia<br />
322<br />
Cremastus flavoorbitalis, see Trathala flavoorbitalis 193<br />
Cremastus hapaliae Cushman Hym.: Ichneumonidae 191<br />
Crematogaster Hym.: Formicidae 153<br />
Crematogaster curvispinosa Mayr Hym.: Formicidae<br />
cristatus, Telenomus<br />
169, 175<br />
Cristicaudus nepalensis (Takada) Hym.: Aphidiidae<br />
crossota, Plautia<br />
crypticus, Trissolcus<br />
53<br />
Cryptochetum sp. Dip.: Cryptochetidae 293<br />
Cryptogonus orbiculus, see Nephus bipunctatus 296<br />
Cryptolaemus Col.: Coccinellidae 146, 150, 152, 155<br />
Cryptolaemus affinis Crotch Col.: Coccinellidae 153, 292, 304<br />
Cryptolaemus montrouzieri Mulsant Col.: Coccinellidae<br />
296, 299, 300Ð306, 312, 315, 316<br />
146, 150, 152, 153, 290, 292,<br />
Cryptolaemus wallacii Crotch Col.: Coccinellidae 153<br />
Cryptoprymna Hym.: Pteromalidae 243, 247<br />
Cryptus Hym.: Ichneumonidae 193<br />
Cryptus rutovinctus Pratt Hym.: Ichenumonidae<br />
culpator cincticornis, Stenichneumon<br />
curculionis, Physoderes<br />
curvicauda, Labia<br />
curvimaculatus, Chelonus<br />
curvispinosa, Crematogaster<br />
324<br />
Cuspicona simplex Walker Hem.: Pentatomidae<br />
custodiens, Anoplolepis<br />
225<br />
Cycloneda sanguinea Linnaeus Col.: Coccinellidae 332, 341<br />
Cydia ptychora, see Leguminivora ptychora 194<br />
Cylas formicarius (Fabricius) Col.: Curculionidae<br />
cylindrica, Sphaerophoria<br />
11<br />
Cylindromyia Dip.: Tachinidae 22
Scientific Index 493<br />
Cylindromyia rufifemur Paramonov Dip.: Tachinidae 202, 217, 233<br />
Cynipoide Hym.: Cynipidae 241, 247, 251<br />
Cyrtogaster Hym.: Pteromalidae<br />
cyrus, Telenomus<br />
245<br />
dactylopii, Leptomastix<br />
dactylopii, Prochiloneurus<br />
Dactylosternum abdominale (Fabricius) Col.: Hydrophilidae 85, 91, 93, 97, 102, 103<br />
Dactylosternum hydrophiloides Macleay Col.: Hydrophilidae 85, 91, 96Ð98, 100, 102<br />
Dactylosternum intermedium Reg. Col.: Hydrophilidae 91, 102<br />
Dactylosternum pr<strong>of</strong>undus Auct. Col.: Hydrophilidae 91, 102<br />
Dactylosternum subdepressum Lap. Col.: Hydrophilidae 91, 98<br />
Dactylosternum subquadratum Fairmaire Col.: Hydrophilidae 91<br />
darci, Alloxysta<br />
darvicola, Eurytoma<br />
dasyni, Anastatus<br />
dasyops, Winthemia<br />
Deanolis albizonalis (Hampson) Lep.: Pyralidae 110<br />
Deanolis sublimbalis Snellen Lep.: Pyralidae 2, 105Ð112<br />
deion, Trichogramma<br />
delhiensis, Lysiphlebus<br />
Delta pyriforme Fabricius Hym.: Eumenidae 25, 30<br />
deltiger, Toxares<br />
delucchii, Asecodes<br />
delucchii, Teleopterus<br />
demolitor, Microplitis<br />
Dendrocerus Hym.: Megaspilidae 65, 68<br />
dendrolimi, Trichogramma<br />
Dermatopelte Hym.: Eulophidae 191, 192<br />
Dermatopolle, see Dermatopelte 191<br />
derogata, Syllepte<br />
detorquens, Technomyrmex<br />
Diadegma Hym.: Ichneumonidae 325<br />
Diadegma insulare (Cresson) Hym.: Ichneumonidae 325, 348<br />
Diadegma plutellae Hym.: Ichneumonidae 325<br />
diadema, Sinea<br />
Diadiplosis hirticornis Felt Dip.: Cecidomyiidae 293<br />
Diaeretiella rapae (M'Intosh) Hym.: Braconidae 38, 48, 53
494 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Diaphorencyrtus aligarhensis (Shafee, Alam and Agarwal) Hym.: Encyrtidae 113, 120,<br />
121, 123Ð133<br />
Diaphorencyrtus diaphorinae, see Diaphorencyrtus aligarhensis 121<br />
Diaphorina auberti Hollis Hem.: Psyllidae 116, 131<br />
Diaphorina cardiae Crawford Hem.: Psyllidae 131<br />
Diaphorina citri Kuwayama Hem.: Psyllidae 1, 113Ð134<br />
Diaretus rapae, see Diaeretiella rapae<br />
Diaspis bromeliae (Keiren) Hem.: Diaspididae 146, 153<br />
Dichomeris eridantis Meyrick Lep.: Gelechiidae 194<br />
Dichrodiplosis sp. Dip.: Cecidomyiidae 293<br />
Dicrodiplosis guatemalensis Felt Dip.: Cecidomyiidae 151<br />
dictynna, Aphanogmus<br />
dictynna, Calliceras<br />
dictynna, Ceraphron<br />
Dictyotis caenosus (Westwood) Hem.: Pentatomidae 225<br />
dilabida, Sturmia<br />
dimidiata, Harmonia<br />
dimidiata, Leis<br />
Dindymus rubiginosus (Fabricius) Hem.: Pyrrhocoridae 166, 168<br />
Diomus Col.: Coccinellidae 150, 300<br />
Diomus flavifrons see Diomus pumilio<br />
Diomus margipallens, see Scymnus margipallens 150<br />
Diomus pumilio (Weise) Col.: Coccinellidae 292, 306, 310, 313<br />
discolor, Micraspis<br />
Disophrys lutea (BrullŽ) Hym.: Braconidae 24, 30<br />
dispar, Lymantria<br />
Dissolcus fulviventris, see Gryon fulviventris 227<br />
diversus, Pheidologeton<br />
dolichostigma, Melanagromyza<br />
Dolicoderus Hym.: Formicidae 223<br />
dolosus, Enicospilus<br />
donacis, Melanaphis<br />
douglasi, Achrysocharis<br />
douglasi, Chrysonotomyia<br />
Drosica abjectella Walker Lep.: Tineidae 147, 154<br />
dryi, Tamarixia<br />
duplifascialis, Hendecasis<br />
Dysdercus Hem.: Pyrrhocoridae 135, 138
Scientific Index 495<br />
Dysdercus cingulatus (Fabricius) Hem.: Pyrrhocoridae 1, 2, 135Ð139<br />
dysmicocci, Pseudaphycus<br />
Dysmicoccus brevipes (Cockerell) Hem.: Pseudococcidae 1, 2, 141Ð156, 313<br />
Dysmicoccus neobrevipes Beardsley Hem.: Pseudococcidae 141Ð144, 152Ð154<br />
Earias Lep.: Noctuidae 28<br />
Earias biplaga Walker Lep.: Noctuidae 30<br />
Earias insulana Boisduval Lep.: Noctuidae<br />
eastopi, Trioza<br />
194<br />
Echthromorpha agrestoria (Swederus) Hym.: Ichneumonidae 25, 27<br />
Echthromorpha punctum Brutte Hym.: Ichneumonidae 325<br />
Ectophasia crassipennis (Fabricius) Dip.: Tachinidae 202, 224<br />
Ectophasiopsis arcuata (Bigot) Dip.: Tachinidae 202, 213, 221, 233, 324<br />
edentata, Voria<br />
Elachertus Hym.: Eulophidae 268, 273, 283<br />
Elasmus Hym.: Elasmidae 265, 273, 275<br />
Elasmus corbetti Ferri re Hym.: Elasmidae 265<br />
Elasmus tischeriae (Howard) Hym.: Elasmidae 265<br />
Elasmus zehntneri Ferri re Hym.: Elasmidae<br />
elegantilis, Neoleucinodes<br />
Elpe, see Camplyocheta<br />
265, 276, 284<br />
Encarsia Hym.: Aphelinidae 124, 125<br />
Encarsia shafeei, see Encarsia transvena 125<br />
Encarsia transvena (Timberlake) Hym.: Aphelinidae 125<br />
Endaphis maculans (Barnes) Dip.: Cecidomyiidae 57, 79<br />
Enicospilus Hym.: Ichneumonidae 25, 30, 325<br />
Enicospilus dolosus (Tosquinet) Hym.: Ichneumonidae 25, 28<br />
Enicospilus samoana (Kohl) Hym.: Ichneumonidae 25, 27<br />
Ephedrus Hym.: Aphidiidae 38, 71, 83<br />
Ephedrus cerasicola Starù Hym.: Aphidiidae 38, 67<br />
Ephedrus nacheri Quilis Hym.: Aphidiidae 38, 53, 64<br />
Ephedrus persicae Froggatt Hym.: Aphidiidae 38, 53, 64, 74, 83<br />
Ephedrus plagiator (Nees) Hym.: Aphidiidae 38, 53, 64, 71, 74, 75<br />
Epiclerus nomocerus Hym.: Tetracampidae 244<br />
Episyrphus balteatus (De Geer) Dip.: Syrphidae<br />
epius, Spalgis<br />
equatus, Trioxys<br />
43<br />
Eriborus argenteopilosus (Cameron) Hym.: Ichneumonidae 191, 194
496 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
eridantis, Dichomeris<br />
Eriococcus Hem.: Eriococcidae 312<br />
Eriosoma lanigerum (Hausmann) Hem.: Aphididae<br />
erosa, Anomis<br />
erosa, Cosmophila<br />
ervi, Aphidius<br />
erythraea, Chrysonotomyia<br />
erytreae, Trioza<br />
72<br />
Eublemma amabilis Moore Lep.: Noctuidae 194<br />
Euborellia annulipes (Lucas) Derm.: Labiidae 90, 102<br />
Euborellia pallipes (Shiraki) Derm.: Labiidae 22, 29<br />
Eucanthecona furcellata (Wolff) Hem.: Pentatomidae 22, 30<br />
Eucelatoria armigera (Coquillett) Dip.: Tachinidae 327, 328<br />
Eucelatoria rubentis (Coquillett) Dip.: Tachinidae 328<br />
Euclytia flava (Townsend) Dip.: Tachinidae 202<br />
Eucoilidea Hym.: Cynipidae 241, 247, 255<br />
Euderus Hym.: Eulophidae 241, 251, 268<br />
Eulissus Col.: Staphylinidae 92<br />
Eumenes campaniformis (Fabricius) Hym.: Eumenidae 25, 30<br />
Eumenes pyriformis, see Delta pyriforme 25, 30<br />
Eupelmus Hym.: Eupelmidae 242<br />
Eupelmus australiensis (Girault) Hym.: Eupelmidae 239, 242<br />
Eupelmus grayi Girault Hym.: Eupelmidae 242<br />
Eupelmus popa, see Eupelmus australiensis 239<br />
Eupelmus urozonus Dalman Hym.: Eupelmidae 242, 271<br />
Eupeodes confrater (Wiedemann) Dip.: Syrphidae 43<br />
Euphorocera Dip.: Tachinidae 328<br />
Euphorocera tachinomoides Townsend Dip.: Tachinidae 328<br />
Euplectrus Hym.: Eulophidae 324<br />
Euplectrus comstockii Howard Hym.: Eulophidae 324<br />
Euplectrus manilae Ashmead Hym.: Eulophidae 24<br />
Euplectrus platyhypenae Howard Hym.: Eulophidae 324, 337<br />
Eurydinotellus viridicoxa, see Callitula viridicoxa 251<br />
Euryrhopalus propinquus Kerrich Hym.: Encyrtidae 145, 147, 149<br />
Euryrhopalus schwarzi (Howard) Hym.: Encyrtidae 145, 147, 149<br />
Euryrophalus pretiosa, see Euryrophalus schwarzi 145, 147<br />
Eurytenes nanus, see Opius phaseoli 252<br />
Eurytoma Hym.: Eurytomidae 239, 242, 246, 247, 271, 277
Scientific Index 497<br />
Eurytoma larvicola Girault Hym.: Eurytomidae 242, 245<br />
Eurytoma poloni Girault Hym.: Eurytomidae 242, 247, 251<br />
Eurytoma syleptae Ferri re Hym.: Eurytomidae 28<br />
Eusandalum incompleta (Bou‹ek) Hym.: Eupelmidae<br />
euschisti, Trissolcus<br />
271, 274, 277<br />
Eutectona macheralis (Walker) Lep.: Pyralidae 194<br />
Euthyrhynchus floridanus (Linnaeus) Col.: Pentatomidae 330<br />
Eutochia pulla (Erichson) Col.: Tenebrionidae 92<br />
Eutrichopodopsis nitens, see Trichopoda giacomellii 202, 221, 232<br />
Euzophera ferticella Ragonot Lep.: Pyralidae<br />
evanescens, Trichogramma<br />
194<br />
Evania appendigaster (Linnaeus) Hym.: Evaniidae<br />
exaltatorius, Trogus<br />
exigua, Spodoptera<br />
exiguae, Hyposoter<br />
exiguum, Trichogramma<br />
111<br />
Exochomus flavipes (Thunberg) Col.: Coccinellidae 43, 292, 305, 310<br />
Exochomus flaviventris Mader Col.: Coccinellidae 292<br />
Exochomus metallicus Korsch Col.: Coccinellidae<br />
exoletum, Praon<br />
299, 310<br />
Exorista apicalia Baranov Dip.: Tachinidae 23<br />
Exorista sorbillans (Wiedemann) Dip.: Tachinidae 23, 27<br />
Exoristobia philippensis Ashmead Hym.: Encyrtidae<br />
exsaguis, Zelus<br />
201, 221<br />
fabae, Aphis<br />
fabarum, Lysiphlebus<br />
facialis, Pediobius<br />
fecundus, Ooencyrtus<br />
Ferrisia virgata Cockerell Hem.: Pseudococcidae 314<br />
ferrugatus, Belonuchus<br />
ferrugineus , Bactrocera<br />
ferruginosus, Chrysopilus<br />
ferticella, Euzophera<br />
ficus, Planococcus<br />
filicornis, Callitula<br />
flava, Anomis<br />
flava, Cosmophila
498 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
flava, Euclytia<br />
flavicincta, Campoletis<br />
flavipes, Aphelinus<br />
flavipes, Aphidius<br />
flavipes, Exochomus<br />
flavitarsis, Mischocyttarus<br />
flaviventris, Exochomus<br />
flavoorbitalis, Cremastus<br />
flavoorbitalis, Trathala<br />
flavopicta, Spilochalcis<br />
floridaensis, Aphidius<br />
floridanum, Copidosoma<br />
floridanus, Euthyrhynchus<br />
Fopius Hym.: Braconidae 240, 248, 251<br />
formicarius, Cylas<br />
formosa, Chrysonotomyia<br />
formosa, Chrysopa<br />
formosana, Schizobremia<br />
formosanus, Chelonus<br />
fragilis, Meteorus<br />
fragilis, Pseudococcus<br />
frauenfeldi, Bactrocera<br />
friedae, Camponotus<br />
fullawayi, Zaplatycerus<br />
Fulvius nigricornis Poppius Hem.: Miridae 90, 102<br />
fulviventris, Dissolcus<br />
fulviventris, Gryon<br />
fulviventris, Hadronotus<br />
funebris, Aphidius<br />
funeralis, Ascotolinx<br />
furcellata, Cantheconidia<br />
furcellata, Eucanthecona<br />
furnacalis, Ostrinia<br />
fuscifemora, Nepiera<br />
fuscipennis, Amblyteles<br />
galactopus, Baryscapus<br />
Galepsomyia Hym.: Eulophidae 268, 273, 275
Scientific Index 499<br />
Galerita bicolor (Drury) Col.: Carabidae 90, 102<br />
Gambrus ultimus (Cresson) Hym.: Ichneumonidae 325<br />
Gelis tenellus (Say) Hym.: Ichneumonidae 325<br />
geminata, Solenopsis<br />
Geocoris Hem.: Lygaeidae 22, 69<br />
Geocoris pallens StŒl Hem.: Lygaeidae 330, 340<br />
Geocoris punctipes Say Hem.: Lygaeidae 330<br />
Geotomus pygmaeus Dallas Hem.: Cydnidae 90, 102, 103<br />
gestuosus, Pterocormus<br />
giacomellii, Trichopoda<br />
gifuensis, Aphidius<br />
gifuensis, Telenomus<br />
gilva, Timberlakia<br />
Gitonides perspicax Knab Dip.: Drosophilidae 147, 153<br />
Glaucias amyoti (Dallas) Hem.: Pentatomidae 225<br />
globula, Hyperaspis<br />
glomerata, Cotesia<br />
Glyptapanteles vitripennis Curtis Hym.: Braconidae 336<br />
Goniozus Hym.: Bethylidae 166, 168, 182<br />
gossypii, Aphelinus<br />
gossypii, Aphis<br />
gracilis, Lipolexis<br />
graelsii, Xanthodes<br />
gramineum, Acrosternum<br />
graminis, Antonina<br />
graminum, Schizaphis<br />
grandis, Alcaeorrhynchus<br />
granulatus, Rogas<br />
grayi, Eupelmus<br />
greeni, Anagyrus<br />
greeni, Bracon<br />
gregori, Sympiesis<br />
Gryon Hym.: Scelionidae 201, 205, 228, 233<br />
Gryon antestiae, see Gryon fulviventris 227<br />
Gryon fulviventris (Crawford) Hym.: Scelionidae 205, 227<br />
Gryon japonicum (Ashmead) Hym.: Scelionidae 205, 212, 221<br />
Gryon obesum Masner Hym.: Scelionidae 205, 212, 221<br />
guatemalensis, Dicrodiplosis
500 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
guineense, Tetramorium<br />
gustavoi, Trichopoda<br />
Gymnosoma clavata (Rohdendorf) Dip.: Tachinidae 202, 233<br />
Gymnosoma kuramanum Matsumura Dip.: Tachinidae 202<br />
Gymnosoma rotundata (Fabricius) Dip.: Tachinidae 202, 233, 234<br />
Hadrojoppa cognatoria Smith Hym.: Ichneumonidae 12<br />
Hadronotus antestiae, see Gryon fulviventris 227<br />
Hadronotus fulviventris, see Gryon fulviventris 277<br />
Halticoptera Hym.: Pteromalidae 243, 247<br />
Halticoptera circulus (Walker) Hym.: Pteromalidae 243, 252<br />
Halticoptera patellana (Dalman) Hym.: Pteromalidae 243, 246, 249<br />
Hambletonia pseudococcina Compere Hym.: Encyrtidae<br />
hampei, Hypothenemus<br />
hampei, Stephanoderes<br />
hamygurivara, Sphegigaster<br />
hapaliae, Cremastus<br />
145, 148, 149, 152, 154Ð156<br />
Harmonia axyridis (Pallas) Col.: Coccinellidae 62, 63, 66, 128<br />
Harmonia dimidiata (Fabricius) Col.: Coccinellidae 43, 68<br />
Harmonia octomaculata (Fabricius) Col.: Coccinellidae 60, 292, 300<br />
Hebertia Hym.: Pteromalidae 243<br />
Helicoverpa Lep.: Noctuidae 27<br />
Helicoverpa armigera (HŸbner) Lep.: Noctuidae<br />
helva, Bogosia<br />
helymus, Metachaeta<br />
helymus, Periscepsia<br />
28, 30, 194<br />
Hemerobius Neu.: Hemerobiidae 331<br />
Hemiptarsenus Hym.: Eulophidae 241<br />
Hemiptarsenus semialbicornis, see Hemiptarsenus varicornis 252<br />
Hemiptarsenus varicornis (Girault) Hym.: Eulophidae<br />
hemipterus, Xenoencyrtus<br />
241, 252<br />
Hendecasis duplifascialis Hampson Lep.: Pyralidae 194<br />
Herbertia Hym.: Pteromalidae 239, 243<br />
Herse convolvuli, see Agrius convolvuli 10<br />
Hesperus sparsior (Bernhauer) Col.: Staphylinidae 92<br />
Heteropsylla cubana Crawford Hem.: Psyllidae 134<br />
Heteropsylla spinulosa Muddiman, Hodkinson & Hollis Hem.: Psyllidae 134
Scientific Index 501<br />
Heterospilus c<strong>of</strong>feicola Schmiedeknecht Hym.: Braconidae<br />
177, 178, 181, 182<br />
166, 169Ð171, 173, 175,<br />
Hexacladia hilaris Burks Hym.: Encyrtidae<br />
hilare, Acrosternum<br />
hilaris, Hexacladia<br />
hindecasisella, Phanerotoma<br />
203<br />
Hippodamia convergens (GuŽrin-MŽneville) Col.: Coccinellidae 69, 332, 341<br />
Hippodamia parenthesis (Say) Col.: Coccinellidae 332<br />
Hippodamia quinqesignata punctulata Le Conte Col.: Coccinellidae 332<br />
Hippodamia variegata (Goeze) Col.: Coccinellidae<br />
hirticornis, Diadiplosis<br />
62, 69<br />
Hister niloticus Marseul Col.: Histeridae<br />
hockiana, Carcelia<br />
h<strong>of</strong>fmanni, Scymnus<br />
hokkaidensis, Trioxys<br />
91<br />
Holcopelte Hym.: Eulophidae 268<br />
Hololepta Col.: Histeridae 91, 97, 98, 102<br />
Hololepta minuta Erichson Col.: Histeridae 97<br />
Hololepta quadridenta (Fabricius) Col.: Histeridae 91, 96Ð 98, 100, 101<br />
Hololepta striaditera Marseul Col.: Histeridae 91<br />
Homalotylus Hym.: Encyrtidae 306<br />
Horismenus Hym.: Eulophidae 268, 273<br />
Horismenus sardus Hym.: Eulophidae<br />
horticola, Chromatomyia<br />
howardi, Tetrastichus<br />
hullensis, Trissolcus<br />
humilis, Aphelinus<br />
humilis, Iridomyrmex<br />
268<br />
Hyalopterus pruni (Ge<strong>of</strong>froy) Hem.: Aphididae<br />
hybneri, Piezodorus<br />
hydrophiloides, Dactylosternum<br />
73<br />
Hypantropha Dip.: Tachinidae 328<br />
Hyperaspis Col.: Coccinellidae 150, 292, 300, 310<br />
Hyperaspis albicollis Gorham Col.: Coccinellidae 150<br />
Hyperaspis c-nigrum Mulsant Col.: Coccinellidae 150, 300<br />
Hyperaspis globula Casey Col.: Coccinellidae 310<br />
Hyperaspis jucunda Mulsant Col.: Coccinellidae 310<br />
Hyperaspis lateralis Mulsant Col.: Coccinellidae 292
502 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Hyperaspis polita Weise Col.: Coccinellidae 292, 306<br />
Hyperaspis silvestrii Weise Col.: Coccinellidae 150, 154<br />
Hyperomyzus lactucae (Linnaeus) Hem.: Aphididae 62<br />
Hyphantrophaga Dip.: Tachinidae 327<br />
Hyposolenus laevigatus, see Plaesius laevigatus 91<br />
Hyposoter exiguae (Viereck) Hym.: Ichneumonidae 317, 325, 338, 340Ð343, 346, 347<br />
Hypothenemus Col.: Scolytidae 163, 164, 178<br />
Hypothenemus hampei (Ferrari) Col.: Scolytidae 1, 2, 157Ð183<br />
illota, Carcelia<br />
importatus, Opius<br />
includens, Chrysodeixis<br />
includens, Nephus<br />
includens, Pseudoplusia<br />
includens, Scymnus<br />
incompleta, Eusandalum<br />
incompleta, Ratzeburgiola<br />
inconspicua, Palexorista<br />
indagator, Sycanus<br />
indefensa, Plutarchia<br />
indica, Cosmophila<br />
indicum, Monomorium<br />
indicus, Paragus<br />
indicus, Trioxys<br />
infesta, Casinaria<br />
ingenuus, Cirrospilus<br />
innota, Sarcodexia<br />
insidiosus, Orius<br />
insulana, Earias<br />
insulare, Diadegma<br />
insularis, Angitia<br />
insularis, Blepyrus<br />
insularis, Chelonus<br />
insularis, Omicrogiton<br />
intacta, Thopeutis<br />
intermedia, Brachymeria<br />
intermedium, Dactylosternum<br />
interocularis, Thyreocephalus
Scientific Index 503<br />
interpunctella, Plodia<br />
intima, Chrysopa<br />
Iridomyrmex humilis Mayr Hym.: Formicidae<br />
irregularis, Chrysopa<br />
144<br />
Ischiodon aegyptius (Wiedemann) Dip.: Syrphidae 43<br />
Ischiodon scutellaris Fabricius Dip.: Syrphidae 43, 68<br />
Iseropus stercorator orgyiae (Ashmead) Hym.: Ichneumonidae 325<br />
Isyropa Dip.: Tachinidae 22<br />
Itamoplex Hym.: Ichneumonidae 191Ð193<br />
Itoplectis conquisator (Say) Hym.: Ichneumonidae 325<br />
japonica, Lysiphlebia<br />
japonica, Propylea<br />
japonicum, Gryon<br />
japonicum, Trichogramma<br />
japonicus, Anastatus<br />
japonicus, Meteorus<br />
japonicus, Pleurotropposis<br />
japonicus, Stenomesius<br />
javanus, Plaesius<br />
johnsoni, Ooencyrtus<br />
jokahamae, Polistes<br />
josefi, Clausenia<br />
jucunda, Hyperaspis<br />
kamburovi, Allotropa<br />
kashmiriensis, Aphelinus<br />
kenyae, Planococcus<br />
kivuensis, Anagyrus<br />
koebelei, Cleodiplosis<br />
Kratoysma Hym.: Eulophidae 269, 272, 277<br />
Kratoysma citri Bou‹ek Hym.: Eulophidae 269, 276, 284<br />
kraunhiae, Planococcus<br />
kulingensis, Chrysopa<br />
kuramanum, Gymnosoma<br />
Labia borellii Burr Derm.: Labiidae 90<br />
Labia curvicauda (Motschulsky) Derm.: Labiidae 90
504 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Labidura riparia Pallas Derm.: Labiduridae 330<br />
lacciperda, Chrysopa<br />
lacciperda, Odontochrysa<br />
lacciperda, Plesiochrysa<br />
lactucae, Hyperomyzus<br />
lacunatus, Psix<br />
laeviceps, Cotesia<br />
laevigatus, Hyposolenus<br />
laevigatus, Plaesius<br />
lanata, Chrysopa<br />
lanigerum, Eriosoma<br />
lanipes, Trichopoda<br />
laphygmae, Meteorus<br />
larvicola, Eurytoma<br />
lasus, Brachymeria<br />
lateralis, Hyperaspis<br />
Lebia analis Dejean Col.: Carabidae 332<br />
Leguminivora ptychora (Meyrick) Lep.: Tortricidae 194<br />
leibyi, Schizocerophaga<br />
Leis dimidiata, see Harmonia dimidiata<br />
leopardina, Marietta<br />
lepelleyi, Trissolcus<br />
Leptochirus unicolor, see Priochirus unicolor 92, 93, 102<br />
Leptomastidea abnormis (Girault) Hym.: Encyrtidae 287, 295Ð297, 299Ð301, 303Ð307,<br />
313, 315, 316<br />
Leptomastix Hym.: Encyrtidae 301<br />
Leptomastix dactylopii Howard Hym.: Encyrtidae 148, 149, 287, 295Ð297, 299Ð307,<br />
312Ð316<br />
Leptomastix nigrocoxalis Compere Hym.: Encyrtidae 295, 297<br />
Leptomastix trilongifasciatus Girault Hym.: Encyrtidae 295, 297<br />
Lespesia sp. Dip.: Tachinidae 328<br />
Lespesia achaetoneura Dip.: Tachinidae 328<br />
Lespesia archippivora (Riley) Dip.: Tachinidae 328<br />
Leucinodes orbonalis GuenŽe Lep.: Pyralidae 1, 2, 185Ð195<br />
Leucopis Dip.: Chamaemyiidae 43, 65<br />
Leucopis alticeps Czerny Dip.: Chamaemyiidae 293, 306<br />
Leucopis bella Loew Dip.: Chamaemyiidae 293<br />
Leucopis silesiaca Eggar Dip.: Chamaemyiidae 293
Scientific Index 505<br />
lilacinus, Planococcus<br />
lineatus, Brumoides<br />
Lioderma Col.: Histeridae 91, 102<br />
Lioderma quadridentata, see Hololepta quadridentata 91<br />
liogaster, Opius<br />
Lipolexis ambiguus, see Lysiphlebus ambiguus 39<br />
Lipolexis gracilis Fšrster Hym.: Aphidiidae 38, 53, 68, 75<br />
Lipolexis pseudoscutellaris, see Lipolexis scutellaris 54<br />
Lipolexis scutellaris Mackauer Hym.: Aphidiidae 39, 54, 64, 66Ð68, 75, 82, 83<br />
Liriomyza brassicae (Riley) Dip.: Agromyzidae 252<br />
Liriomyza sativae Blanchard Dip.: Agromyzidae 252<br />
Liriomyza trifolii (Burgess) Dip.: Agromyzidae 251<br />
liriomyzae, Meruana<br />
Lissauchenius venator LafertŽ Col.: Carabidae 26<br />
littoralis, Spodoptera<br />
lituratis, Piezodorus<br />
Lixophaga Dip.: Tachinidae 195<br />
Lobodiplosis pseudococci, see Vincentodiplosis pseudococci 151<br />
lodosi, Trissolcus<br />
longefasciatus, Cirrospilus<br />
longicornis, Paratrechina<br />
longipes, Anoplolepis<br />
longispinus, Pseudococcus<br />
longiventris, Paragus<br />
lonicerae, Aphidius<br />
louisianae, Scymnus<br />
lucens, Asaphes<br />
lunata, Cheilomenes<br />
lurida, Scotinophara<br />
lutea, Disophrys<br />
lutea, Oligochrysa<br />
lutea, Ophelosia<br />
Lygocerus, see Dendrocerus 65<br />
Lymantria dispar (Linnaeus) Lep.: Lymantriidae 225<br />
lyncus, Cirrospilus 39<br />
Lysaphidus Hym.: Aphidiidae 37<br />
Lysaphidus platensis, see Aphidius colemani 37<br />
Lysaphidus schimitscheki (Fahriner) Hym.: Aphidiidae 54
506 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Lysiphlebia Hym.: Aphidiidae 39<br />
Lysiphlebia japonica (Ashmead) Hym.: Aphidiidae 39, 54, 63, 64, 66, 75, 82, 83<br />
Lysiphlebia mirzai Shuja Uddin Hym.: Aphidiidae 54, 70, 75<br />
Lysiphlebia rugosa Starù & Schlinger Hym.: Aphidiidae 39<br />
Lysiphlebus Hym.: Aphidiidae 40, 55, 63<br />
Lysiphlebus ambiguus, see Lysiphlebus fabarum 39, 54, 71<br />
Lysiphlebus cardui, see Lysiphlebus fabarum 54<br />
Lysiphlebus confusus, see Lysiphlebus fabarum 39, 54, 75<br />
Lysiphlebus delhiensis (Subba Rao & Sharma) Hym.: Aphidiidae 39<br />
Lysiphlebus fabarum (Marshall) Hym.: Aphidiidae 39, 54, 60, 61, 65Ð67, 71, 75, 83<br />
Lysiphlebus salicaphis, see Adialytus salicaphis 37<br />
Lysiphlebus shaanxiensis Chou and Xian Hym.: Aphidiidae 54<br />
Lysiphlebus testaceipes (Cresson) Hym.: Aphidiidae<br />
76, 77, 82, 83<br />
5, 40, 55, 60, 61, 63, 64, 66Ð69, 74,<br />
macheralis, Eutectona<br />
macheralis, Pyrausta<br />
macrophallus, Sturmia<br />
macrosiphophagum, Toxares<br />
maculans, Endaphis<br />
maculata fuscilabris, Ceratomegilla<br />
maculipennis, Pseudaphycus<br />
maculiventris, Podisus<br />
madecassa, Brachymeria<br />
Madremyia saundersii (Williston) Dip.: Tachinidae 328<br />
maidis, Rhopalosiphum<br />
malayensis, Ooencyrtus<br />
mali, Aphelinus<br />
Mallada basalis (Walker) Neu.: Chrysopidae 291, 302<br />
Mallada boninensis Okamoto Neu.: Chrysopidae 271, 291, 302<br />
manilae, Euplectrus<br />
manilae, Snellenius<br />
marginata, Mesograpta<br />
marginatum, Acrosternum<br />
marginellus, Collops<br />
marginipallens, Diomus<br />
marginiventris, Cotesia<br />
margipallens, Scymnus
Scientific Index 507<br />
mariae, Spilochalcis<br />
Marietta Hym.: Aphelinidae 128<br />
Marietta javensis, see Marietta leopardina 124, 129<br />
Marietta leopardina Motschulsky Hym.: Aphelinidae 124, 126, 149<br />
mariscusae, Aphelinus<br />
maro, Trissolcus<br />
marshalli, Sericophoromyia<br />
Maruca testulalis, see Maruca vitrata 194<br />
Maruca vitrata Fabricius Lep.: Pyralidae 194<br />
matricariae, Aphidius<br />
mauritiusi, Scymnus<br />
mecrida, Allotropa<br />
mediterraneus, Carpocoris<br />
mediterraneus, Pnigalio<br />
megacephala, Pheidole<br />
megacephala, Pheidole<br />
Megaselia rufipes (Meigen) Dip.: Phoridae 12, 15<br />
Melanagromyza Dip.: Agromyzidae 236<br />
Melanagromyza atomella (Malloch) Dip.: Agromyzidae 253<br />
Melanagromyza dolichostigma de Meijere Dip.: Agromyzidae 236, 247, 251, 254<br />
Melanagromyza obtusa (Malloch) Dip.: Agromyzidae 236<br />
Melanagromyza phaseoli, see Ophiomyia phaseoli<br />
Melanagromyza sojae (Zehnter) Dip.: Agromyzidae 236, 247, 248, 251, 253, 254<br />
melanagromyzae, Biosteres<br />
melanagromyzae, Opius<br />
Melanaphis donacis (Passerini) Hem.: Aphididae 73<br />
melvillei, Tropidophryne<br />
Menochilus sexmaculatus, see Cheilomenes sexmaculata 65<br />
menthastri, Sphaerophoria<br />
Merismorella shakespearei, see Syntomopus shakespearei 254<br />
Meruana liriomyzae Bou‹ek Hym.: Eulophidae 241, 252<br />
Mesochorus Hym.: Ichneumonidae 25, 29<br />
Mesochorus africanus (Ferri re) Hym.: Ichneumonidae 28<br />
Mesograpta marginata (Say) Dip.: Syrphidae 332<br />
Metachaeta helymus Walker Dip.: Tachinidae 328<br />
metallicus, Exochomus<br />
Meteorus Hym.: Braconidae 21<br />
Meteorus autographae Muesebeck. Hym.: Braconidae 322, 348
508 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Meteorus fragilis Wesmael Hym.: Braconidae 24, 29<br />
Meteorus japonicus, see Meteorus pulchricornis 24<br />
Meteorus laphygmae Viereck Hym.: Braconidae 323<br />
Meteorus pulchricornis (Wesmael) Hym.: Braconidae 21, 24, 29<br />
Metopius Hym.: Ichneumonidae<br />
metricus, Polistes<br />
25<br />
Micraspis discolor (Fabricius) Col.: Coccinellidae 65<br />
Microbracon phyllocnistidis, see Bracon phyllocnistidis 265, 274<br />
Microcharops bimaculata (Ashmead) Hym.: Ichneumonidae 325, 335<br />
Microcharops tibialis (Cresson) Hym.: Ichneumonidae 325<br />
Microdon bicolor Sack Dip.: Syrphidae 43<br />
Microgaster brassicae (Muesebeck) Hym.: Braconidae<br />
347<br />
317, 323, 338Ð341, 343, 346,<br />
Microgaster demolitor, see Microplitis demolitor 336<br />
Microgaster plutellae (Muesebeck) Hym.: Braconidae 323<br />
Microgaster rufiventris, see Microplitis rufiventris 356<br />
Micromus posticus (Walker) Neu.: Hemerobiidae 69<br />
Micromus timidus (Fabricius) Neu.: Hemerobiidae 68<br />
Microplitis alaskensis Ashmead Hym.: Braconidae 323<br />
Microplitis brassicae, see Microgaster brassicae 323<br />
Microplitis demolitor Wilkinson Hym.: Braconidae 356<br />
Microplitis rufiventris Kokujev Hym.: Braconidae<br />
mikan, Sympiesomorpha<br />
minio, Pnigalio<br />
minuta, Hololepta<br />
minutum, Trichogramma<br />
minutus, Orius<br />
mirzai, Lysiphlebia<br />
356<br />
Mischocyttarus flavitarsis (Saussure) Hym.: Vespidae<br />
mitsukurii, Asolcus<br />
mitsukurii, Trissolcus<br />
molestus, Rogas<br />
333<br />
Monomorium Hym.: Formicidae 153<br />
Monomorium indicum Forel Hym.: Myrmecidae<br />
montana, Winthemia<br />
montanus, Patrocloides<br />
montrouzieri, Cryptolaemus<br />
mormideae, Telenomus<br />
67
Scientific Index 509<br />
multicolor, Brachymeria<br />
multilineatum, Zagrammosoma<br />
murrayii, Psylla<br />
myzophagum, Praon<br />
Myzus Hem.: Aphididae 75<br />
Myzus persicae Sulzer Hem.: Aphididae 5, 59, 63, 71Ð74<br />
Nabis alternatus Parshley Col.: Nabidae 330<br />
Nabis americ<strong>of</strong>erus Carayon Hem.: Nabidae 330, 340<br />
Nabis roseipennis (Reuter) Hem.: Nabidae 330<br />
Nabis sin<strong>of</strong>erus Hsiao Hem.: Nabidae 22<br />
nacheri, Ephedrus<br />
nakagawai, Telenomus<br />
nanus, Eurytenes<br />
narangae, Zacharops<br />
nasalis, Cermatulus<br />
nasuta, Allograpta<br />
nasuta, Prorops<br />
nearctaphidis, Trioxys<br />
nebulosa, Pseudiastata<br />
neobrevipes, Dysmicoccus<br />
Neochrysocharis, see Chrysonotomyia 266, 277<br />
Neodimmockia agromyzae, see Hemiptarsenus varicornis 252<br />
Neoleucinodes Lep.: Pyralidae 186<br />
Neoleucinodes elegantilis (GuenŽe) Lep.: Pyralidae 186, 195<br />
Neoprochiloneurus bolivari (Mercet) Hym.: Encyrtidae 306<br />
Neorileya Hym.: Eurytomidae 203<br />
nepalensis, Cristicaudus<br />
Nephopterix rhodobasalis Hampson Lep.: Pyralidae 194<br />
Nephus Col.: Coccinellidae 300, 310<br />
Nephus bilucenarius, see Scymnus bilucenarius 145, 150<br />
Nephus bipunctatus (Kugelann) Col.: Coccinellidae 292, 298, 306, 310<br />
Nephus includens, see Scymnus includens 292<br />
Nephus pictus, see Scymnus bilucenarius 145<br />
Nephus reunioni (Fursch) Col.: Coccinellidae 292, 302, 304, 306, 310, 316<br />
Nepiera fuscifemora Graf Hym.: Ichneumonidae 326<br />
nerii, Aphis<br />
Nesolynx phaeosoma (Waterston) Hym.: Eulophidae 28<br />
Netelia Hym.: Ichneumonidae 325
510 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Nezara Hem.: Pentatomidae 198, 202<br />
Nezara antennata Scott Hem.: Pentatomidae 208, 224, 229, 233, 234<br />
Nezara viridula (Linnaeus) Hem.: Pentatomidae 1, 137, 197Ð233<br />
nezarae, Ooencyrtus<br />
ni, Trichoplusia<br />
niger, Asaphoideus<br />
niger, Thysanus<br />
niger, Xenoencyrtus<br />
nigriceps, Cardiochiles<br />
nigricornis, Chrysopa<br />
nigricornis, Fulvius<br />
nigrifrontalis, Trichopoda<br />
nigripes, Sigalphus<br />
nigritus, Aphelinus<br />
nigrocoxalis, Leptomastix<br />
nigr<strong>of</strong>asciatus, Phonoctonus<br />
nigronervosa, Pentalonia<br />
niloticus, Hister<br />
nitens, Eutrichopodopsis<br />
noctuae, Zenillia<br />
nomocerus, Epiclerus<br />
Noorda albizonalis, see Deanolis albizonalis 106<br />
Norbanus Hym.: Pteromalidae 239<br />
notata, Sympiesis<br />
novemnotata francisciana, Coccinella<br />
Nyereria Hym.: Braconidae 24, 28<br />
obesum, Gryon<br />
obscurata, Brachymeria<br />
obscurus, Pseudococcus<br />
obtusa, Agromyza<br />
obtusa, Melanagromyza<br />
octomaculata, Harmonia<br />
octopunctata, Ankylopteryx<br />
Ocyptera, see Cylindromyia 42<br />
Odontochrysa lacciperda Kimmins Neu.: Chrysopidae 291, 302, 314<br />
Oechalia schellembergii (GuŽrin-MŽneville) Hem.: Pentatomidae 22, 27, 218<br />
oenone, Trissolcus
Scientific Index 511<br />
ogyges, Trissolcus<br />
oleracei, Opius<br />
Oligochrysa lutea (Walker) Neu.: Chrysopidae 291, 300<br />
Olla v-nigrum (Mulsant) Col.: Coccinellidae 332<br />
Omicrogiton insularis Orchym. Col.: Hydrophilidae 91, 102<br />
Ooencyrtus Hym.: Encyrtidae 201, 204, 223, 225, 230<br />
Ooencyrtus californicus Hym.: Encyrtidae 203<br />
Ooencyrtus fecundus Ferri re & VoegelŽ Hym.: Encyrtidae 204<br />
Ooencyrtus johnsoni (Howard) Hym.: Encyrtidae 204<br />
Ooencyrtus malayensis Ferri re Hym.: Encyrtidae 204<br />
Ooencyrtus nezarae Ishii Hym.: Encyrtidae 204, 212, 221, 224, 227<br />
Ooencyrtus pityocampae (Mercet) Hym.: Encyrtidae 204<br />
Ooencyrtus submetallicus (Howard) Hym.: Encyrtidae 204, 212Ð214, 217, 221, 227, 230<br />
Ooencyrtus trinidadensis Crawford Hym.: Encyrtidae<br />
Ophelosia crawfordi Riley Hym.: Encyrtidae 295, 298<br />
204, 213, 221<br />
Ophiomyia centrosematis (de Meijere) Dip.: Agromyzidae 239, 248, 252<br />
Ophiomyia phaseoli (Tryon) Dip.: Agromyzidae 1, 2, 235Ð255<br />
Ophiomyia spencerella Greathead Dip.: Agromyzidae 236, 239, 253, 255<br />
Opius Hym.: Braconidae 240, 247<br />
Opius importatus Fischer Hym.: Braconidae 235, 239, 240, 249, 250, 255<br />
Opius liogaster SzŽpligeti Hym.: Braconidae 240, 248, 252<br />
Opius melanagromyzae, see Opius phaseoli 252<br />
Opius oleracei Fischer Hym.: Braconidae 239, 240<br />
Opius phaseoli Fischer Hym.: Braconidae<br />
optimus, Abacetus<br />
orbana, Antestis<br />
orbonalis, Leucinodes<br />
orestes, Chrysopa<br />
235, 239, 240, 246, 248Ð250, 252, 253, 255<br />
Orius Hem.: Anthocoridae 43<br />
Orius insidiosus Say Col.: Anthocoridae 330<br />
Orius minutus (Linnaeus) Hem.: Anthocoridae 22, 63, 271, 291<br />
Orius tristicolor (White) Hem.: Anthocoridae 330, 340, 341<br />
Ostrinia furnacalis (GuenŽe) Lep.: Pyralidae<br />
ovata, Brachymeria<br />
14<br />
Oxyharma subaenea (Dodd) Hym.: Pteromalidae 243, 254<br />
Pachyneuron Hym.: Pteromalidae 67, 77<br />
Pachyneuron aphidis BouchŽ Hym.: Pteromalidae 57, 65, 68, 78
512 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Pachyneuron concolor (Fšrster) Hym.: Pteromalidae 123, 125<br />
Pachyneuron solitarius (Ratz) Hym.: Pteromalidae 306<br />
Pachyneuron vitodurense Delucchi Hym.: Pteromalidae 68<br />
Pachyophthalmus Dip.: Tachinidae<br />
pacificus, Planococcus<br />
pacificus, Telenomus<br />
190<br />
Paederus Col.: Staphylinidae 223<br />
Palexorista inconspicua (Meigen) Dip.: Tachinidae 23<br />
Palexorista quadrizonula (Thomson) Dip.: Tachinidae<br />
pallens, Chrysopa<br />
pallens, Geocoris<br />
pallidicollis, Pullus<br />
pallidus, Trioxys<br />
pallipes, Euborellia<br />
paolii, Brachymeria<br />
parachrysops, Sturmia<br />
23, 30, 31<br />
Paragus bicolor, see Microdon bicolor 43<br />
Paragus borbonicus Macquart Dip.: Syrphidae 43<br />
Paragus indicus, see Paragus tibialis 43<br />
Paragus longiventris Loewe Dip.: Syrphidae 43<br />
Paragus serratus (Fabricius) Dip.: Syrphidae 43<br />
Paragus tibialis (FallŽn) Dip.: Syrphidae<br />
paramali, Aphelinus<br />
43<br />
Paranaemia vittegera (Mulsant) Col.: Coccinellidae 332<br />
Parapanteles Hym.: Braconidae 24, 28<br />
Paratrechina longicornis (Latreille) Hym.: Formicidae 145<br />
Paratrigonogastra stella, see Sphegigaster stella<br />
parenthesis, Hippodamia<br />
patellana, Halticoptera<br />
247, 254<br />
Patrocloides montanus (Cresson) Hym.: Ichneumonidae 325, 340<br />
Pauesia antennata Makerjl Hym.: Aphidiidae 40<br />
Pauridia peregrina, see Coccidoxenoides peregrinus 294, 297, 311<br />
Pediobius Hym.: Eulophidae 239, 241<br />
Pediobius acantha (Walker) Hym.: Eulophidae 241<br />
Pediobius facialis (Giraud) Hym.: Eulophidae 324, 336<br />
Pediobius sexdentatus Hym.: Pteromalidae<br />
pennipes, Trichopoda<br />
pennsylvanicum, Acrosternum<br />
326, 338
Scientific Index 513<br />
pensylvanica, Vespula<br />
Pentalonia nigronervosa Coquerel Hem.: Aphididae 67, 68, 74, 81<br />
pentheus, Chrysocharis<br />
perdignus, Pseudaphycus<br />
peregrina, Pauridia<br />
peregrinator, Calosoma<br />
peregrinus, Coccidoxenoides<br />
Periscepsia helymus, see Metachaeta helymus 327<br />
perla, Chrysopa<br />
perplexus, Rogas<br />
persicae, Ephedrus<br />
persicae, Myzus<br />
perspicax, Gitonides<br />
perticella, Euzophera<br />
petiolatus, Semielacher<br />
pfeifferi, Allograpta<br />
phaeosoma, Nesolynx<br />
Phanerotoma Hym.: Braconidae 190, 192, 195<br />
Phanerotoma hindecasisella Cameron Hym.: Braconidae 190, 194<br />
phaseoli, Agromyza<br />
phaseoli, Melanagromyza<br />
phaseoli, Ophiomyia<br />
phaseoli, Opius<br />
Pheidole Hym.: Formicidae 67, 153, 156<br />
Pheidole megacephala (Fabricius) Hym.: Formicidae 92, 93, 144, 152Ð154, 223, 300<br />
Pheidologeton diversus (Jerdon) Hym.: Formicidae 155<br />
Phenacoccus solani Ferris Hem.: Pseudococcidae 313<br />
philippensis, Exoristobia<br />
Phonoctonus Hem.: Reduviidae 139<br />
Phonoctonus nigr<strong>of</strong>asciatus StŒl Hem.: Reduviidae 139<br />
Phonoctonus subimpictus StŒl Hem.: Reduviidae 139<br />
Phorocera Dip.: Tachinidae 328<br />
phorodontis, Aphidius<br />
Phorticus pygmaeus Poppius Hem.: Nabidae 90, 102<br />
Phryxe vulgaris (FallŽn) Dip.: Tachinidae 328<br />
phyllocnistidis, Bracon<br />
phyllocnistidis, Microbracon<br />
Phyllocnistis Lep.: Gracillariidae 258
514 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Phyllocnistis citrella Stainton Lep.: Gracillariidae 1, 2, 257Ð286<br />
phyllocnistis, Cirrospilus<br />
phyllocnistis, Scotolinx<br />
phyllocnistoides, Cirrospiloideus<br />
phyllocnistoides, Cirrospilus<br />
phyllocnistoides, Cirrostichus<br />
phyllocnistoides, Tetrastichus<br />
Phymastichus c<strong>of</strong>fea La Salle Hym.: Eulophidae 157, 166, 168, 174, 178, 182<br />
Physoderes curculionis China Hem.: Reduviidae 90, 102<br />
Phytomyza albiceps Meigen Dip.: Agromyzidae 254<br />
Phytomyza atricornis, see Chromatomyia horticola 239, 252<br />
picipes, Aphidius<br />
pictus, Cirrospilus<br />
pictus, Nephus<br />
pictus, Scymnus<br />
Pieris rapae Linnaeus Lep.: Pieridae 337<br />
Piezodorus hybneri (Gmelin) Hem.: Pentatomidae 218, 227<br />
Piezodorus lituratus (Fabricius) Hem.: Pentatomidae 224<br />
pilipes, Trichopoda<br />
Pimpla aequalis (Provancher) Hym.: Ichneumonidae 325<br />
pinguis, Winthemia<br />
ÔPireniniÕ Hym.: Pteromalidae 276<br />
pisum, Acyrthosiphon<br />
pityocampae, Ooencyrtus<br />
Plaesius javanus Erichson Col.: Histeridae 85, 91, 93, 95Ð100, 103, 104<br />
Plaesius laevigatus Marseul Col.: Histeridae 85, 91, 93, 95, 96, 100, 104<br />
plagiator, Ephedrus<br />
Planococcus Hem.: Pseudococcidae 288<br />
Planococcus citri (Risso) Hem.: Pseudococcidae 1, 2, 155, 287Ð316<br />
Planococcus ficus (Signoret) Hem.: Pseudococcidae 288<br />
Planococcus kenyae (Le Pelley) Hem.: Pseudococcidae 288, 311<br />
Planococcus kraunhiae (Kuwana) Hem.: Pseudococcidae 296, 305<br />
Planococcus lilacinus Cockerell Hem.: Pseudococcidae 313, 314<br />
Planococcus pacificus Cox Hem.: Pseudococcidae 288, 313<br />
platensis, Aphidius<br />
platensis, Lysaphidius<br />
platneri, Trichogramma<br />
platyhypenae, Euplectrus
Scientific Index 515<br />
Platynaspis Col.: Coccinellidae 310<br />
Platynaspis capicola Crotch Col.: Coccinellidae 68<br />
Platysoma Col.: Histeridae 91<br />
Platysoma abrupta Erichson Col.: Histeridae 91, 102<br />
Plautia brunnipennis (Montrouzier and Signoret) Hem.: Pentatomidae 225<br />
Plautia crossota (Dallas) Hem.: Pentatomidae 206<br />
Plesiochrysa lacciperda, see Chrysopa lacciperda<br />
pleuralis, Alloxysta<br />
291<br />
Pleurotropitiella albipes Blanchard Hym.: Eulophidae 203<br />
Pleurotropposis japonicus (Kamijo) Hym.: Eulophidae 269<br />
Plodia interpunctella (HŸbner) Lep.: Pyralidae<br />
plusiae, Siphona<br />
344<br />
Plutarchia Hym.: Eurytomidae 242, 247<br />
Plutarchia bicarinativentris Girault Hym.: Eurytomidae 239, 242<br />
Plutarchia indefensa (Walker) Hym.: Eurytomidae 242, 248, 254<br />
Plutella xylostella Linnaeus Lep.: Yponomeutidae<br />
plutellae, Angitia<br />
plutellae, Cotesia<br />
plutellae, Diadegma<br />
plutellae, Microgaster<br />
337<br />
Pnigalio Hym.: Eulophidae 269, 273, 274, 277, 278<br />
Pnigalio agraules Walker Hym.: Eulophidae 269, 276<br />
Pnigalio mediterraneus, see Pnigalio agraules 276<br />
Pnigalio minio (Walker) Hym.: Eulophidae<br />
podisi, Telenomus<br />
269, 277, 284<br />
Podisus Hem.: Pentatomidae 220<br />
Podisus maculiventris (Say) Hem.: Pentatomidae 27, 331<br />
Polistes Hym.: Vespidae 26, 29<br />
Polistes apachus Saussure Hym.: Vespidae 333<br />
Polistes jokahamae Radoszkowski Hym.: Vespidae 26, 29<br />
Polistes metricus Say Hym.: Vespidae<br />
polita, Hyperaspis<br />
poloni, Eurytoma<br />
333<br />
Polycystomyia beneficia, see Callitula viridicoxa 251<br />
Polycystus Hym.: Pteromalidae 243, 246<br />
Polycystus propinquus Waterston Hym.: Pteromalidae<br />
popa, Eupelmus<br />
posticus, Micromus<br />
243
516 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Praon Hym.: Braconidae 40, 56, 77<br />
Praon abjectum (Haliday) Hym.: Braconidae 40, 55<br />
Praon absinthii Bagnall Hym.: Braconidae 55, 64<br />
Praon exsoletum (Nees) Hym.: Braconidae 40, 55<br />
Praon myzophagum Mackauer Hym.: Braconidae 56<br />
Praon volucre (Haliday) Hym.: Braconidae<br />
pretiosa, Euryrophalus<br />
pretiosum, Trichogramma<br />
41, 56, 62, 77<br />
Priochirus unicolor (Laporte) Col.: Staphylinidae 92, 93, 102<br />
Pristomerus Hym.: Ichneumonidae 325<br />
Pristomerus spinator (Fabricius) Hym.: Ichneumonidae 325<br />
Pristomerus testaceus Morley Hym.: Ichneumonidae 191, 194<br />
Prochiloneurus dactylopii (Howard) Hym.: Encyrtidae<br />
pr<strong>of</strong>undum, Dactylosternum<br />
305<br />
Propagalerita bicolor, see Galerita bicolor<br />
propinquus, Euryrhopalus<br />
propinquus, Polycystus<br />
90<br />
Propylea japonica (Thunberg) Col.: Coccinellidae 62, 63<br />
Prorops nasuta Waterston Hym.: Bethylidae 157, 166, 168, 170Ð182<br />
Prospalta capensis, see Condica capensis 194<br />
Protomicroplitis Hym.: Braconidae<br />
pruinosus, Cosmopolites<br />
pruni, Hyalopterus<br />
24, 28<br />
Psalis americana, see Carcinophora americana 90, 102<br />
Pseudaphycus Hym.: Encyrtidae 148, 149, 152, 300<br />
Pseudaphycus angelicus (Howard) Hym.: Encyrtidae 152, 295, 298<br />
Pseudaphycus angustifrons Gahan Hym.: Encyrtidae 148<br />
Pseudaphycus dysmicocci Bennett Hym.: Encyrtidae 148, 149, 155<br />
Pseudaphycus maculipennis Mercet Hym.: Encyrtidae 295, 298, 314<br />
Pseudaphycus perdignus Compere and Zinna Hym.: Encyrtidae 295, 300, 301, 309<br />
Pseudiastata nebulosa Coquerell Dip.: Drosophilidae 147, 151, 152, 155, 156<br />
Pseudiastata pseudococcivora Sabrosky Dip.: Drosophilidae<br />
pseudococci, Anagyrus<br />
pseudococci, Lobodiplosis<br />
pseudococci, Vincentodiplosis<br />
pseudococcina, Hambletonia<br />
pseudococcivora, Pseudiastata<br />
151<br />
Pseudococcus brevipes, see Dysmicoccus brevipes 142
Scientific Index 517<br />
Pseudococcus citriculus Green Hem.: Pseudococcidae 303, 311<br />
Pseudococcus comstocki Kuwana Hem.: Pseudococcidae 313<br />
Pseudococcus fragilis Brain Hem.: Pseudococcidae 311<br />
Pseudococcus longispinus Targioni-Tozzetti Hem.: Pseudococcidae 311<br />
Pseudococcus obscurus Essig Hem.: Pseudococcidae 311<br />
Pseudococcus vitis Niediel Hem.: Pseudococcidae<br />
pseudomagnoliarum, Coccus<br />
313<br />
Pseudoperichaeta Dip.: Tachinidae 190<br />
Pseudoplusia includens, see Chrysodeixis includens<br />
pseudoscutellaris, Lipolexis<br />
psidii, Chloropulvinaria<br />
340<br />
Psix lacunatus Johnson & Masner Hym.: Scelionidae 205<br />
Psix striaticeps (Dodd) Hym.: Scelionidae 205, 220<br />
Psylla Hem.: Psyllidae 131<br />
Psylla citricola Yang and Li Hem.: Psyllidae 116<br />
Psylla citrisuga Yang and Li Hem.: Psyllidae 116<br />
Psylla murrayii Mathur Hem.: Psyllidae 116<br />
Psyllaephagus Hym.: Encyrtidae 123, 125, 129<br />
Psyllaephagus diaphorinae, see Diaphorencyrtus aligarhensis 121<br />
Pterocormus gestuosus (Cresson) Hym.: Ichneumonidae 326<br />
Pteromalus Hym.: Pteromalidae 205<br />
Pterosema subaenea, see Oxyharma subaenea<br />
ptychora, Cydia<br />
ptychora, Leguminivora<br />
pulchricornis, Meteorus<br />
pulla, Eutochia<br />
254<br />
Pullus Col.: Coccinellidae 43<br />
Pullus pallidicollis Mulsant Col.: Coccinellidae 292, 302<br />
Pulvinaria Hem.: Coccidae<br />
pumilio, Diomus<br />
punctata, Xanthopimpla<br />
punctipes, Geocoris<br />
punctum, Echthromorpha<br />
punicae, Aphis<br />
purpurea, Clausenia<br />
pygmaeus, Geotomus<br />
pygmaeus, Phorticus<br />
312<br />
Pyrausta macheralis, see Eutectona macheralis 194
518 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
pyreformis, Eumenes<br />
pyriforme, Delta<br />
pyrrhopya, Winthemia<br />
Quadrastichus Hym.: Eulophidae 269, 272, 275, 277, 281<br />
quadratus, Belonuchus<br />
quadridenta, Hololepta<br />
quadridentata, Lioderma<br />
quadripustulata, Winthemia<br />
quadristriata, Scotolinx<br />
quadristriatus, Cirrospilus<br />
quadrivittatus, Scymnus<br />
quadrizonula, Palexorista<br />
quatrosignata, Sticholotis<br />
quinqesignata punctulata, Hippodamia<br />
radiata, Tamarixia<br />
radiatus, Tetrastichus<br />
ramburi, Chrysopa<br />
rapae, Diaeretiella<br />
rapae, Diaretus<br />
rapae, Pieris<br />
Ratzeburgiola incompleta, see Eusandalum incompleta 271, 277<br />
remota, Bessa<br />
renardii, Zelus<br />
repanda, Coccinella<br />
reunioni, Nephus<br />
reunioni, Scymnus<br />
rhanis, Synopeas<br />
Rhizobius ventralis Erichson Col.: Coccinellidae 146<br />
rhodobasalis, Nephopterix<br />
Rhopalosiphum maidis (Fitch) Hem.: Aphididae 77<br />
Rhychium attrisium Van der Vecht Hym.: Vespidae 111<br />
ribis, Aphidius<br />
rietscheli, Trioxys<br />
riparia, Labidura<br />
Riptortus serripes (Fabricius) Hem.: Alydidae 218<br />
robusta, Sphaerophoria
oepkei, Scymnus<br />
Rogas Hym.: Braconidae<br />
Rogas granulatus De Gant Hym.: Braconidae 323<br />
Rogas molestus Cresson Hym.: Braconidae 323<br />
Rogas rufocoxalis Gahan Hym.: Braconidae 323<br />
rosae, Aphidius<br />
roseipennis, Nabis<br />
rotundata, Gymnosoma<br />
rubentis, Eucelatoria<br />
rubicola, Trioxys<br />
rubiginosus, Dindymus<br />
rubricatus, Xenoencyrtus<br />
rudus, Trissolcus<br />
ruficrus, Apanteles<br />
ruficrus, Cotesia<br />
rufifemur, Cylindromyia<br />
rufilabris, Chrysopa<br />
rufipes, Megaselia<br />
rufocoxalis, Rogas<br />
rufopicta, Winthemia<br />
rugosa, Lysiphlebia<br />
rugosa, Sphegigaster<br />
rugosa, Trigonogastra<br />
ruralis, Voria<br />
rutila, Agonoscelis<br />
rutovinctus, Cryptus<br />
Scientific Index 519<br />
saccharicolus, Blepyrus<br />
salicaphis, Adialytus<br />
salicaphis, Lysiphlebus<br />
salicis, Aphidius<br />
saltator, Conocephalus<br />
samoana, Enicospilus<br />
sanctus, Sympherobius<br />
sandanis, Sympiesis<br />
sanguinea, Cycloneda<br />
Sarcodexia innota (Walker) Dip.: Sarcophagidae 202<br />
Sarcodexia sternodontis Townsend Dip.: Sarcophagidae 202, 327
520 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Sarcophaga sp. Dip.: Sarcophagidae<br />
sardus, Horismenus<br />
sativae, Liriomyza<br />
saundersii, Madremyia<br />
sawadai, Anagyrus<br />
scapuliferus, Scymnus<br />
328<br />
Scarites Col.: Carabidae<br />
scelestes, Brinckochrysa<br />
scelestes, Chrysopa<br />
schayeri, Calosoma<br />
schellenbergii, Oechalia<br />
schimitscheki, Lysaphidus<br />
90, 102<br />
Schizaphis graminum (Rondani) Hem.: Aphididae 5<br />
Schizobremia formosana Felt Dip.: Cecidomyiidae 147, 154<br />
Schizocerophaga leibyi Townsend Dip.: Tachinidae<br />
schwarzi, Euryrhopalus<br />
328<br />
Scleroderma Hym.: Bethylidae 180<br />
Scleroderma cadaverica Benoit Hym.: Bethylidae 168, 180<br />
Scotinophara lurida (Burmeister) Hem.: Pentatomidae 224<br />
Scotolinx phyllocnistis, see Cirrospilus phyllocnistis 267<br />
Scotolinx quadristriata, see Cirrospilus quadristriata<br />
scutellaris, Ischiodon<br />
scutellaris, Lipolexis<br />
scuticarinatus, Trissolcus<br />
267<br />
Scymnus Col.: Coccinellidae 43, 66, 146, 150, 152, 293, 305<br />
Scymnus apetzi Mulsant Col.: Coccinellidae 292, 306<br />
Scymnus apiciflavus (Motschulsky) Col.: Coccinellidae 146, 292<br />
Scymnus bilucenarius (Mulsant) Col.: Coccinellidae 145, 146, 150, 152<br />
Scymnus biguttatus Mulsant Col.: Coccinellidae 292, 306<br />
Scymnus binaevatus Mulsant Col.: Coccinellidae 293, 296, 310<br />
Scymnus bipunctatus, see Nephus bipunctatus 292, 296, 310<br />
Scymnus constrictus Mulsant Col.: Coccinellidae 68<br />
Scymnus h<strong>of</strong>fmanni Weise Col.: Coccinellidae 62, 63, 66<br />
Scymnus includens (Kirsch) Col.: Coccinellidae 292, 314<br />
Scymnus louisianae Chapin Col.: Coccinellidae 69<br />
Scymnus margipallens Mulsant Col.: Coccinellidae 150, 154<br />
Scymnus mauritiusi Korsch. Col.: Coccinellidae 146, 153<br />
Scymnus pictus Gorham Col.: Coccinellidae 150
Scientific Index 521<br />
Scymnus quadrivittatus Mulsant Col.: Coccinellidae 150<br />
Scymnus reunioni, see Nephus reunioni 292, 302, 304, 310<br />
Scymnus roepkei De Fluiter Col.: Coccinellidae 292<br />
Scymnus scapuliferus Mulsant Col.: Coccinellidae 43<br />
Scymnus sordidus Horn Col.: Coccinellidae 152, 292, 300, 310<br />
Scymnus subvillosus (Goeze) Col.: Coccinellidae 292, 306<br />
Scymnus trepidulus Weise Col.: Coccinellidae 43<br />
Scymnus uncinatus Sicard Col.: Coccinellidae 145, 150<br />
semialbicornis, Hemiptarsenus<br />
Semielacher Hym.: Eulophidae 269, 276<br />
Semielacher petiolatus (Girault) Hym.: Eulophidae 257, 264, 269, 281, 284, 285<br />
semiflavus, Aphelinus<br />
semifumatum, Trichogramma<br />
Senometopia, see Carcelia 111<br />
Senotainia Dip.: Sarcophagidae 327<br />
septempunctata, Chrysopa<br />
septempunctata, Coccinella<br />
Sericophoromyia marshalli Villeneuve Dip.: Tachinidae 23<br />
serratus, Paragus<br />
serripes, Riptortus<br />
sexdentatus, Pediobius<br />
sexmaculata, Cheilomenes<br />
sexmaculatus, Menochilus<br />
seychellensis, Telenomus<br />
shaanxiensis, Lysiphlebus<br />
shakespearei, Merismorella<br />
shakespearei, Syntomopus<br />
side, Spilochalcis<br />
Sigalphus nigripes He & Chen Hym.: Braconidae 24<br />
signata, Chrysopa<br />
Signiphora Hym.: Signiphoridae 123, 129<br />
silesiaca, Leucopis<br />
silvestri, Hyperaspis<br />
similis, Aphidius<br />
simplex, Chilo<br />
simplex, Cuspicona<br />
Sinea complexa Caudell Hem.: Reduviidae 331<br />
Sinea confusa Caudell Hem.: Reduviidae 331
522 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Sinea diadema (Fabricius) Hem.: Reduviidae 331<br />
sinensis, Trioxys<br />
sinica, Chrysopa<br />
sinica, Chrysoperla<br />
sin<strong>of</strong>erus, Nabis<br />
Siphona Dip.: Tachinidae 328<br />
Siphona plusiae Coquillett Dip.: Tachinidae 328<br />
sipius, Trissolcus<br />
Sitotroga cerealella (Olivier) Lep.: Gelechiidae 344<br />
smithi, Coccodiplosis<br />
Snellenius manilae (Ashmead) Hym.: Braconidae 323<br />
sojae, Melanagromyza<br />
solani, Phenacoccus<br />
Solenopsis Hym.: Formicidae 152, 153, 275<br />
Solenopsis geminata (Fabricius) Hym.: Formicidae 144, 153, 154, 223<br />
solitarius, Pachyneuron<br />
solitus, Telenomus<br />
solocis, Trissolcus<br />
sonchi, Aphidius<br />
sonorensis, Campoletis<br />
sorbillans, Exorista<br />
sordidus, Cosmopolites<br />
sordidus, Scymnus<br />
Spalgis epius (Westwood) Lep.: Lycaenidae 294, 302, 314<br />
sparsior, Hesperus<br />
spencerella, Ophiomyia<br />
Sphaerophoria cylindrica (Say) Dip.: Syrphidae 332<br />
Sphaerophoria menthastri (Linnaeus) Dip.: Syrphidae 332<br />
Sphaerophoria robusta (Curran) Dip.: Syrphidae 332<br />
Sphegigaster Hym.: Pteromalidae 244, 247<br />
Sphegigaster agromyzae, see Sphegigaster voltairei 254<br />
Sphegigaster brunneicornis (Ferri re) Hym.: Pteromalidae 243, 245, 246, 254<br />
Sphegigaster hamygurivara Hym.: Pteromalidae 243<br />
Sphegigaster rugosa (Waterston) Hym.: Pteromalidae 243, 246, 254<br />
Sphegigaster stella (Girault) Hym.: Pteromalidae 244, 247, 254<br />
Sphegigaster stepicola Bou‹ek Hym.: Pteromalidae 244, 245, 254<br />
Sphegigaster voltairei (Girault) Hym.: Pteromalidae 244, 245, 247, 254<br />
Spilochalcis flavopicta Cresson Hym.: Chalcididae 323
Scientific Index 523<br />
Spilochalcis nr. mariae (Riley) Hym.: Chalcididae 323<br />
Spilochalcis side Walker Hym.: Chalcididae 323<br />
spinator, Pristomerus<br />
spiraecola, Aphis<br />
splendens, Chrysoplatycerus<br />
Spodoptera exigua HŸbner Lep.: Noctuidae 16, 30, 194, 345<br />
Spodoptera littoralis (Boisduval) Lep.: Noctuidae 30<br />
stella, Paratrigonogastra<br />
stella, Sphegigaster<br />
Stenichneumon culpator cincticornis (Cresson) Hym.: Ichneumonidae 326, 348<br />
Stenomesius japonicus (Ashmead) Hym.: Eulophidae 269<br />
Stephanoderes hampei, see Hypothenemus hampei 158<br />
stephanoderis, Cephalonomia<br />
stepicola, Sphegigaster<br />
stercorator orgyiae, Iseropus<br />
sternodontis, Sarcodexia<br />
Sticholotis quatrosignata Weise Col.: Coccinellidae 146<br />
Stictopisthus africanus, see Mesochorus africanus 28<br />
Stiretrus anchorago (Fabricius) Col.: Pentatomidae 331<br />
striaditera, Hololepta<br />
striata, Trathala<br />
striaticeps, Psix<br />
striatipes, Sympiesis<br />
Sturmia atropivora, see Zygobothria atropivora 12<br />
Sturmia auratocauda, see Cadurcia auratocauda 22<br />
Sturmia bimaculata, see Palexorista inconspicua 23<br />
Sturmia dilabida Villeneuve Dip.: Tachinidae 12, 16<br />
Sturmia macrophallus, see Zygobothria ciliata 12<br />
Sturmia parachrysops Bezzi Dip.: Tachinidae 190<br />
subaenea, Oxyharma<br />
subaenea, Pterosema<br />
subaeneus, Thysanus<br />
subdepressum, Dactylosternum<br />
subimpictus, Phonoctonus<br />
sublimbalis, Deanolis<br />
submetallicus, Ooencyrtus<br />
subquadratum, Dactylosternum<br />
subvillosus, Scymnus
524 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
sulphurea, Cheilomenes<br />
suppressalis, Chilo<br />
suturalis, Brumus<br />
Sycanus indagator StŒl Col.: Reduviidae<br />
syleptae, Apanteles<br />
syleptae, Eurytoma<br />
331<br />
Syllepte derogata Fabricius Lep.: Pyralidae 28, 194<br />
Sympherobius barberi (Banks) Neu.: Chrysopidae 291, 295, 298, 305<br />
Sympherobius sanctus Tjeder Neu.: Chrysopidae 291, 303<br />
Sympiesis Hym.: Eulophidae 264, 269, 276, 279<br />
Sympiesis gregori Hym.: Eulophidae 269, 276<br />
Sympiesis striatipes (Ashmead) Hym.: Eulophidae 270, 274, 275, 277<br />
Sympiesomorpha mikan, see Stenomesius japonicus 269<br />
Synopeas rhanis (Walker) Hym.: Platygasteridae 80<br />
Syntomopus Hym.: Pteromalidae 244<br />
Syntomopus shakespearei (Girault) Hym.: Pteromalidae 244, 254<br />
Syrphophagus africanus Gahan Hym.: Encyrtidae 68<br />
Syrphophagus aphidivora (Mayr) Hym.: Encyrtidae 42, 68, 72<br />
Syrphophagus taiwanus Hayat and Lin Hym.: Encyrtidae 123, 125<br />
Syrphus Dip.: Syrphidae 43, 67, 69, 293, 300<br />
Syrphus balteatus, see Episyrphus balteatus 43<br />
Syrphus confrater, see Eupeodes confrater 43<br />
tachinomoides, Euphorocera<br />
taiwanus, Syrphophagus<br />
Tamarixia Hym.: Eulophidae 128, 133<br />
Tamarixia dryi (Waterston) Hym.: Eulophidae 129, 130<br />
Tamarixia leucaenae Bou‹ek Hym.: Eulophidae 134<br />
Tamarixia radiata (Waterston) Hym.: Eulophidae 113, 120, 121, 123Ð134<br />
Tapinoma Hym.: Formicidae 275<br />
Technomyrmex detorquens Walker Hym.: Formicidae 153<br />
Telenomus Hym.: Scelionidae 12, 14, 15, 201, 205, 213, 218, 220, 221, 227, 233, 326<br />
Telenomus acrobates Giard. Hym.: Scelionidae 306<br />
Telenomus chloropus (Thomson) Hym.: Scelionidae<br />
227, 229, 234<br />
200, 205, 211Ð213, 217, 221, 224,<br />
Telenomus comperei Crawford Hym.: Scelionidae 205, 226<br />
Telenomus cristatus Johnson Hym.: Scelionidae 205<br />
Telenomus cyrus Nixon Hym.: Scelionidae 206, 230
Scientific Index 525<br />
Telenomus gifuensis Ashmead Hym.: Scelionidae 206, 212, 221, 224<br />
Telenomus mormideae Costa Lima Hym.: Scelionidae 206, 220, 221<br />
Telenomus nakagawai, see Telenomus chloropus 211, 224, 229<br />
Telenomus pacificus (Gahan) Hym.: Scelionidae 205, 226<br />
Telenomus podisi (Ashmead) Hym.: Scelionidae 205, 221<br />
Telenomus seychellensis Kieffer Hym.: Scelionidae 205, 220<br />
Telenomus solitus Johnson Hym.: Scelionidae 326<br />
Teleopterus Hym.: Eulophidae 270, 272, 288<br />
Teleopterus delucchii Bou‹ek Hym.: Eulophidae 270, 278<br />
tenellus, Gelis<br />
testaceipes, Lysiphlebus<br />
testaceus, Pristomerus<br />
testulalis, Maruca<br />
tetracanthus, Zelus<br />
Tetramorium bicarinatum (Nylander) Hym.: Formicidae 85, 92, 100, 101<br />
Tetramorium guineense, see Tetramorium bicarinatum 92, 100<br />
Tetrastichus Hym.: Eulophidae 120, 123, 125, 129, 131, 133, 241, 246, 270, 273, 274,<br />
276, 277, 285<br />
Tetrastichus ayyari, see Tetrastichus howardi 24<br />
Tetrastichus howardi (Olliff) Hym.: Eulophidae 24, 29<br />
Tetrastichus phyllocnistoides, see Citrostichus phyllocnistoides 268<br />
Tetrastichus radiatus, see Tamarixia radiata 121<br />
texanus, Chelonus<br />
thalense, Trichogramma<br />
theclarum, Aplomya<br />
theobromae, Aenasius<br />
theristis, Pammene<br />
Thopeutis intacta Snellen Lep.: Pyralidae 284<br />
thyantae, Trissolcus<br />
Thyreocephalus interocularis Eppelsheim Col.: Staphylinidae 85, 92, 93, 103<br />
Thysanus niger Ashmead Hym.: Encyrtidae 148<br />
Thysanus subaeneus Forst. Hym.: Encyrtidae 306<br />
tibialis, Brachymeria<br />
tibialis, Microcharops<br />
tibialis, Paragus<br />
Timberlakia gilva Prinsloo Hym.: Encyrtidae 295, 298<br />
timidus, Micromus<br />
tischeriae, Elasmus
526 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Toxares macrosiphophagum Shuja Uddin Hym.: Aphidiidae 56<br />
Toxares zakai Shuja Uddin Hym.: Aphidiidae 41<br />
Toxoptera aurantii (Boyer de Fonscolombe) Hem.: Aphididae<br />
transcaspicus, Aphidius<br />
transvena, Encarsia<br />
transversalis, Coccinella<br />
transversoguttata, Coccinella<br />
64, 75, 79<br />
Trathala flavoorbitalis (Cameron) Hym.: Ichneumonidae 185, 191Ð193, 195<br />
Trathala striata Cameron Hym.: Ichneumonidae<br />
trepidulus, Scymnus<br />
191<br />
Trichogramma Hym.: Trichogrammatidae 13, 14, 16, 21, 25, 30, 195, 317, 321, 327, 341<br />
Trichogramma achaeae Nagaraja & Nagarkatti Hym.: Trichogrammatidae 13Ð16<br />
Trichogramma agriae Nagaraja Hym.: Trichogrammatidae 13Ð16<br />
Trichogramma australicum Girault Hym.: Trichogrammatidae 13Ð16, 326<br />
Trichogramma brevicapillum Pinto & Platner Hym.: Trichogrammatidae 326<br />
Trichogramma chilonis Ishii Hym.: Trichogrammatidae 13, 14, 16, 25, 29, 30, 111<br />
Trichogramma chilotraeae Nagaraja and Nagarkatti Hym.: Trichogrammatidae 11, 326<br />
Trichogramma confusum Viggiani Hym.: Trichogrammatidae 13, 15<br />
Trichogramma deion Pinto & Oatman Hym.: Trichogrammatidae 326<br />
Trichogramma dendrolimi Matsumura Hym.: Trichogrammatidae 21, 25, 29<br />
Trichogramma evanescens Westwood Hym.: Trichogrammatidae 326, 344<br />
Trichogramma exiguum Pinto & Platner Hym.: Trichogrammatidae 326<br />
Trichogramma japonicum Ashmead Hym.: Trichogrammatidae 25, 30, 326<br />
Trichogramma minutum Riley Hym.: Trichogrammatidae<br />
344<br />
13, 15, 25, 27, 29, 326, 339,<br />
Trichogramma platneri Nagarkatti Hym.: Trichogrammatidae 326, 344, 346<br />
Trichogramma pretiosum Riley Hym.: Trichogrammatidae 27, 327, 337, 338, 344, 346<br />
Trichogramma semifumatum (Perkins) Hym.: Trichogrammatidae 327, 340<br />
Trichogramma thalense Pinto & Oatman Hym.: Trichogrammatidae 327<br />
Trichoplusia ni HŸbner Lep.: Noctuidae 1, 2, 317Ð347<br />
Trichopoda Dip.: Tachinidae 203, 208, 232<br />
Trichopoda giacomellii (Blanchard) Dip.: Tachinidae<br />
231, 232, 234<br />
202, 211, 218, 220, 221, 225, 226,<br />
Trichopoda gustavoi, see Trichopoda giacomellii 202<br />
Trichopoda lanipes (Fabricius) Dip.: Tachinidae 202, 232<br />
Trichopoda nigrifrontalis, see Trichopoda giacomelli 202<br />
Trichopoda pennipes (Fabricius) Dip.: Tachinidae<br />
221, 224Ð226, 230, 231, 233<br />
197, 201, 208, 209, 212Ð215, 217,
Scientific Index 527<br />
Trichopoda pilipes (Fabricius) Dip.: Tachinidae<br />
233<br />
trifasciatus, Closterocerus<br />
trifolii, Liriomyza<br />
197, 203, 211Ð214, 217, 221, 225, 231,<br />
Trigonogastra agromyzae, see Sphegigaster voltairei 245, 254<br />
Trigonogastra rugosa, see Sphegigaster rugosa<br />
trilongifasciatus, Leptomastix<br />
trinidadensis, Ooencyrtus<br />
254<br />
Triommata coccidivora (Felt) Dip.: Cecidomyidae 293, 302<br />
Trioxys Hym.: Aphidiidae 42, 57<br />
Trioxys acalephae (Marshall) Hym.: Aphidiidae 41, 56<br />
Trioxys angelicae (Haliday) Hym.: Aphidiidae 41, 56, 65, 71, 78<br />
Trioxys asiaticus Telenga Hym.: Aphidiidae 41, 56<br />
Trioxys auctus (Haliday) Hym.: Aphidiidae 41, 56<br />
Trioxys basicurvus Shuja Uddin Hym.: Aphidiidae 56, 64<br />
Trioxys centaureae (Haliday) Hym.: Aphidiidae 41<br />
Trioxys cirsii (Curtis) Hym.: Aphidiidae 41<br />
Trioxys communis Gahan Hym.: Aphidiidae 56, 63, 66Ð68, 78, 82, 83<br />
Trioxys complanatus Quilis Hym.: Aphidiidae 42, 56<br />
Trioxys equatus Samanta, Tamili and Raychaudhuri Hym.: Aphidiidae 56<br />
Trioxys hokkaidensis Takada Hym.: Aphidiidae 42<br />
Trioxys indicus Subba Rao & Sharma Hym.: Aphidiidae 42, 57, 61Ð65, 78, 79, 82, 83<br />
Trioxys nearctaphidis (Mackauer) Hym.: Aphidiidae 42<br />
Trioxys pallidus Haliday Hym.: Aphidiidae 57<br />
Trioxys rietscheli Mackauer Hym.: Aphidiidae 57, 68<br />
Trioxys rubicola Shuja Uddin Hym.: Aphidiidae 64<br />
Trioxys sinensis Mackauer Hym.: Aphidiidae 42, 57, 67<br />
Trioza citroimpura Yang and Li Hem.: Psyllidae 116<br />
Trioza eastopi, see Trioza litseae 129<br />
Trioza erytreae (Del Guerico) Hem.: Psyllidae 119, 120, 129, 130<br />
Trioza litseae Bordaga Hem.: Psyllidae 129<br />
Trissolcus Hym.: Scelionidae 201, 206, 211, 221, 226, 227, 233<br />
Trissolcus aloysiisabaudiae (Fouts) Hym.: Scelionidae 206, 220<br />
Trissolcus basalis (Wollaston) Hym.: Scelionidae 197, 201, 206, 209, 211Ð228, 230<br />
Trissolcus brochymenae (Ashmead) Hym.: Scelionidae 206<br />
Trissolcus crypticus Clarke Hym.: Scelionidae 207, 210, 211<br />
Trissolcus euschisti Ashmead Hym.: Scelionidae 207<br />
Trissolcus hullensis (Harrington) Hym.: Scelionidae 207
528 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Trissolcus lepelleyi (Nixon) Hym.: Scelionidae 207, 220<br />
Trissolcus lodosi (Szab—) Hym.: Scelionidae 207<br />
Trissolcus maro Nixon Hym.: Scelionidae 207, 220<br />
Trissolcus mitsukurii (Ashmead) Hym.: Scelionidae 206, 211Ð215, 217, 221, 224, 229<br />
Trissolcus oenone (Dodd) Hym.: Scelionidae 206<br />
Trissolcus ogyges (Dodd) Hym.: Scelionidae 206<br />
Trissolcus rudus Le Hym.: Scelionidae 206<br />
Trissolcus scuticarinatus (Costa Lima) Hym.: Scelionidae 206<br />
Trissolcus sipius (Nixon) Hym.: Scelionidae 206, 220<br />
Trissolcus solocis Johnson Hym.: Scelionidae 206<br />
Trissolcus thyantae Ashmead Hym.: Scelionidae 206<br />
Trissolcus urichi Hym.: Scelionidae 206<br />
Trissolcus utahensis Hym.: Scelionidae<br />
tristicolor, Orius<br />
tristis, Anasa<br />
206<br />
Trogus exaltatorius Panzer Hym.: Ichneumonidae 12<br />
Tropidophryne melvillei Compere Hym.: Encyrtidae<br />
truncatellum, Copidosoma<br />
309<br />
ultimus, Gambrus<br />
uncinatus, Scymnus<br />
undecimpunctata, Coccinella<br />
unicolor, Leptochirus<br />
unicolor, Priochirus<br />
urichi, Trissolcus<br />
urozonus, Eupelmus<br />
urticae, Aphidius<br />
utahensis, Trissolcus<br />
uzbekistanicus, Aphidius<br />
v-nigrum, Olla<br />
varicornis, Hemiptarsenus<br />
variegata, Adonia<br />
variegata, Hippodamia<br />
variegatus, Cirrospilus<br />
varipes, Aphelinus<br />
venator, Lissauchenius<br />
ventralis, Rhizobius
Scientific Index 529<br />
Vespula pensylvanica (Saussure) Hym.: Vespidae 333<br />
vicina, Cheilomenes<br />
Vincentodiplosis pseudococci (Felt) Dip.: Cecidomyiidae 145, 151, 156<br />
viridicoxa, Callitula<br />
viridicoxa, Eurydinotellus<br />
viridula, Nezara<br />
Visnuella brevipetiolatu, see Zaommomentedon brevipetiolatus 270, 275<br />
vitis, Pseudococcus<br />
vitodurense, Pachyneuron<br />
vitrata, Maruca<br />
vitripennis, Glyptapanteles<br />
vittatus, Cirrospilus<br />
vittatus, Collops<br />
vittegera, Paranaemia<br />
voltairei, Sphegigaster<br />
volucre, Praon<br />
Voria edentata (Baran<strong>of</strong>) Dip.: Tachinidae 328<br />
Voria ruralis (FallŽn) Dip.: Tachinidae 317, 328, 336Ð341, 345Ð347<br />
vulgaris, Phryxe<br />
Vulgichneumon brevicinctor (Say) Hym.: Ichneumonidae 326, 348<br />
walkeri, Chartocerus<br />
wallacii, Cryptolaemus<br />
Wasmannia auropunctata (Roger) Hym.: Formicidae 174<br />
Winthemia Dip.: Tachinidae 329, 337<br />
Winthemia dasyops (Wiedemann) Dip.: Tachinidae 21, 23, 28<br />
Winthemia montana Rein. Dip.: Tachinidae 328<br />
Winthemia pinguis Dip.: Tachinidae 337<br />
Winthemia pyrrhopyga (Wiedemann) Dip.: Tachinidae 337<br />
Winthemia quadripustulata (Fabricius) Dip.: Tachinidae 328<br />
Winthemia rufopicta (Bigot) Dip.: Tachinidae 329<br />
Xanthodes graellsii (Feisthamel) Lep.: Noctuidae 30<br />
Xanthogramma aegyptium, see Ischiodon aegyptius 43<br />
Xanthopimpla punctata (Fabricius) Hym.: Ichneumonidae 25, 191, 192<br />
Xenoencyrtus Hym.: Encyrtidae 204<br />
Xenoencyrtus hemipterus (Girault) Hym.: Encyrtidae 204, 211, 213, 214, 218, 221<br />
Xenoencyrtus niger, see Xenoencyrtus hemipterus 204, 211, 213, 221
530 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Xenoencyrtus rubricatus Riek Hym.: Encyrtidae 204<br />
xylostella, Plutella<br />
yakutatensis, Cotesia<br />
yasudi, Callitula<br />
Zacharops narangae Cushman Hym.: Braconidae 25<br />
Zagrammosoma Hym.: Braconidae 278<br />
Zagrammosoma multilineatum (Ashmead) Hym.: Eulophidae<br />
zakai, Toxares<br />
270, 273, 278<br />
Zaommomentedon Hym.: Eulophidae 270, 276<br />
Zaommomentedon brevipetiolatus Kamijo Hym.: Eulophidae 270, 275, 277, 281, 285<br />
Zaplatycerus fullawayi Timberlake Hym.: Encyrtidae<br />
zehntneri, Elasmus<br />
149<br />
Zelus bilobus Say Hem.: Reduviidae 331<br />
Zelus exsaguis StŒl Hem.: Reduviidae 331<br />
Zelus renardii Kalenati Hem.: Reduviidae 331<br />
Zelus tetracanthus StŒl Hem.: Reduviidae 331<br />
Zenilla blanda blanda (Osten Sacken) Dip.: Tachinidae 329<br />
Zenillia cosmophilae, see Carcelia cosmophilae 22<br />
Zenillia noctuae, see Carcelia illota<br />
zizyphi, Aphis<br />
20<br />
Zygobothria atropivora (Robineau-Desvoidy) Dip.: Tachinidae 12, 16<br />
Zygobothria ciliata (Wulp) Dip.: Tachinidae 12, 15, 23
7 General Index<br />
abelmoschus, Hibiscus<br />
aconitifolia, Vigna<br />
Acridotheres tristis 26<br />
Aegle marmelos 262<br />
alatae 34, 35, 46, 70<br />
alfalfa looper, see Autographa californica 342<br />
Allothrombium pulvinum 58, 63<br />
Alseodaphne semicarpifolia 262<br />
Althaea rosea 19<br />
Amaranthus 19<br />
Ananas comosus 142<br />
anise, see Clausena anisumolens<br />
anisopliae, Metarhizium<br />
anisumolens, Clausena<br />
annuus, Helianthus<br />
antiquorum, Colocasia<br />
117<br />
Apocynum venotum 29<br />
apterae 34, 46, 47<br />
Arabian c<strong>of</strong>fee, see C<strong>of</strong>fea arabica 159, 165, 181<br />
arabica, C<strong>of</strong>fea 181<br />
Arbutilon 19<br />
areca palm 144<br />
Argentine ant, see Iridomyrmex humilis 144<br />
Arthrobotrys 59<br />
artificial diet 11, 35, 170, 177, 187, 320<br />
arvensis, Convolvulus<br />
<strong>Asian</strong> citrus psyllid, see Diaphorina citri<br />
asiatica, Toddalia<br />
114<br />
asparagus 320<br />
Aspergillus flavus<br />
astrigata, Pardosa<br />
138, 298, 301, 334<br />
Atalantia 118<br />
atropurpureum, Macroptilium<br />
atropurpureus, Phaseolus<br />
aurantiifolia, Citrus<br />
aurea, Vigna<br />
531
532 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
aureus, Phaseolus<br />
australisiaca, Microcitrus<br />
avocado 108<br />
Bacillus thuringiensis 21, 27, 320, 334<br />
Bacillus thuringiensis wuhanensis<br />
bacteriophora, Heterorhabditis<br />
26<br />
baculovirus 191, 342<br />
banana, see Musa sapientum 68, 85--104, 144, 145, 154, 163, 289<br />
banana aphid, see Pentalonia nigronervosa 67, 68, 74<br />
banana weevil borer, see Cosmopolites sordidus 85, 86<br />
barley 5<br />
bassiana, Beauveria<br />
batatas, Ipomoea<br />
bats 321<br />
bean, see Phaseolus vulgaris<br />
238, 251, 320<br />
35, 47, 63, 161, 199, 210, 219, 236,<br />
bean fly, see Ophiomyia phaseoli 235, 236<br />
bean stem borer, see Melanagromyza sojae<br />
Beauveria 128, 167, 199<br />
Beauveria bassiana 59, 69, 93, 122, 128, 157, 166, 169, 170, 174--<br />
176, 182, 183, 334<br />
beet armyworm, see Spodoptera exigua<br />
bele, see Hibiscus manihot<br />
Bergera koenigii 116, 117<br />
19<br />
bigheaded ant, see Pheidole megacephalaI 144<br />
bindweed, see Convulvulus arvensis 11<br />
black berry nightshade, see Solanum nigrum 187<br />
black citrus aphid, see Toxoptera aurantii 75<br />
black gram, see Vigna mungo 237<br />
black legume aphid, see Aphis craccivora 34<br />
bluish dogbane, see Apocynum venotum 29<br />
Botrytis stephanoderis see Beauveria bassiana 169<br />
bottle gourd 78<br />
Bouea burmanica 105, 108<br />
brinjal, see Solanum melongena 185<br />
brinjal fruit borer, see Leucinodes orbonalis 186<br />
brinjal stem borer, see Euzophera ferticella 194<br />
broccoli 321
unneum, Metarrhizium<br />
burmanica, Bouea<br />
cabbage 317, 320, 321, 337<br />
cabbage aphid, see Brevicoryne brassicae 48<br />
cabbage looper, see Trichoplusia ni 317, 318<br />
cabbage white butterfly, see Pieris rapae<br />
caerulea, Geoplana<br />
caerulea, Kontikia<br />
337<br />
Caesalpinia 163<br />
cairica, Ipomoea<br />
cajan, Cajanus<br />
Cajanus cajan<br />
calcarata, Vigna<br />
237<br />
caloxylon, Merrillia<br />
Canavalia ensiformis<br />
canephora, C<strong>of</strong>fea<br />
cannabinus, Hibiscus<br />
238<br />
canola 320<br />
cantaloupe 320<br />
cape gooseberry 187<br />
Capsella 35<br />
capsicum 187, 200, 320<br />
carcocapsae, Steinernema<br />
carrot 320<br />
cassava 289<br />
castor, see Ricinus communis 232<br />
Catharanthus roseus 116<br />
catimor c<strong>of</strong>fee 176<br />
cauliflower 320<br />
celery 320, 341<br />
Centrosema 163<br />
Cephalosporium lecanii 59, 69, 122, 128<br />
Chenopodium 338<br />
chick pea 236<br />
chico 108<br />
chinense, Clerodendrum<br />
Chinese cabbage 63<br />
Chiracanthium inclusum 278<br />
General Index 533
534 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
chlordimeform 27<br />
chrysanthemum 5, 65<br />
Ciconia nigra 16<br />
Cinnamomum zeylanica<br />
cinnamomum, Jasminum<br />
citri, Xanthomonas<br />
262<br />
citricola scale, see Coccus pseudomagnoliarum 299<br />
citron, see Citrus medica 117<br />
Citrullus lanatus 19<br />
Citrus 35, 47, 64--66, 77, 113--134, 257--316<br />
Citrus aurantifolia 117, 262<br />
Citrus hystrix 117<br />
Citrus limon 117, 262<br />
Citrus madurensis 117<br />
Citrus maxima 117<br />
Citrus maxima var. racemosa 117<br />
Citrus medica 117, 262<br />
Citrus reticulata 115, 117<br />
Citrus sinensis 117<br />
citrus canker fungus, see Xanthomonas citri 263<br />
citrus greening 113, 114, 116, 119, 120, 128--130<br />
citrus leaf mottle 128, 130<br />
citrus leafminer, see Phyllocnistis citrella 257, 258<br />
citrus mealybug, see Planococcus citri 287, 288<br />
citrus psyllid, see Diaphorina citri 114<br />
citrus tristeza virus 36, 47<br />
citrus vein phloem degeneration 128<br />
Cladosporium oxysporum 298, 305<br />
Clausena anisumolens 116, 117<br />
Clausena excavata 117<br />
Clausena indica 118<br />
Clausena lansium 117<br />
Clerodendrum chinense 14, 15<br />
Clubiona 278<br />
cocoa 35, 47, 289<br />
coconut bug, see Amblypelta cocophaga 226<br />
coconut spathe bug, see Axiagastus campbelli<br />
coerulia, Tweedia<br />
226
General Index 535<br />
C<strong>of</strong>fea 163, 288<br />
C<strong>of</strong>fea arabica 158, 159, 165, 176, 181<br />
C<strong>of</strong>fea canephora 159, 165, 181<br />
c<strong>of</strong>feanum, Colleotrichium<br />
c<strong>of</strong>fee 35, 47, 144, 157--183, 289, 290, 302--304<br />
c<strong>of</strong>fee berry borer, see Hypothenemus hampei 158<br />
c<strong>of</strong>fee berry disease, see Colletotrichum c<strong>of</strong>feanum 170<br />
c<strong>of</strong>fee leaf rust, see Hemileia vastatrix 170, 176<br />
Colleotrichium c<strong>of</strong>feanum 170<br />
Colocasia antiquorum 14, 15<br />
Colocasia esculenta<br />
communis, Ricinus<br />
comosus, Ananas<br />
71<br />
Convolvulus arvensis 11<br />
cotton 17, 19, 21, 27--30, 46, 47, 60, 62, 63, 65, 68, 69, 78, 137, 161,<br />
200, 220, 320, 335, 338, 340--345, 347<br />
cotton aphid, see Aphis gossypii 46<br />
cotton semi looper, see Anomis flava 18<br />
cotton stainer, see Dysdercus cingulatus 135, 136<br />
cowpea, see Vigna unguiculata or Vigna sinensis<br />
219, 222, 227, 236, 238, 239<br />
19, 35, 36, 200,<br />
cowpea aphid, see Aphis craccivora 34<br />
crazy ant, see Paratrechina longicornis 145<br />
creeping sensitive plant, see Mimosa invisa<br />
croceus, Xysticus<br />
134<br />
Crotalaria 163, 251--254<br />
Crotalaria juncea 238<br />
Crotalaria laburnifolia 238, 245<br />
Crotalaria mucronata 238<br />
cucumber 47, 64, 66, 67, 73, 83, 187, 320<br />
curry bush, see Bergera koenigii 117<br />
custard apple 299, 300<br />
Cyclosa insulana 65<br />
cyfluthrin 112
536 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
date 289<br />
DD136 Steinernema carcocapsae 138<br />
Deccan hemp, see Hibiscus cannabinus 19<br />
deltamethrin 112<br />
Dendroica palmarum 333<br />
desi cotton 19<br />
Dialium lacourtianum 163<br />
diamondback moth, see Plutella xylostella<br />
diazoma, Thelohania<br />
337<br />
Dioscorea 164<br />
dissecta, Merremia<br />
Dolichos lablab, see Lablab niger 238<br />
egg plant, see Solanum melongena 47, 64, 65, 78, 83, 185--195<br />
egg plant fruit and shoot borer, see Leucinodes orbonalis 186<br />
elephant lemon, see Citrus medica 262<br />
endosulfan 180, 182<br />
ensiformis, Canavalia<br />
Entomophthora 13, 59, 335<br />
Entomophthora exitialis 59<br />
Entomophthora fresenii 298<br />
Entomophthora fumosa 298, 300<br />
Entomophthora gammae 334<br />
Entomophthora sp. ÔgrylliÕ type 13<br />
Entomophthora sphaerosperma 334<br />
Eremocitrus glauca 262<br />
Erigonidium graminicolum<br />
esculenta, Colocasia<br />
esculentum, Lycopersicon<br />
esculentus, Hibiscus<br />
excavata, Clausena<br />
exitialis, Entomophthora<br />
exotica, Murraya<br />
26, 62
General Index 537<br />
faba, Vicia<br />
feltiae, Steinernema<br />
fig 288, 289, 306<br />
fire ant, see Solenopsis geminata 144<br />
flavus, Aspergillus<br />
flax 231<br />
Fortunella 116, 117<br />
French bean, see Phaseolus vulgaris 237, 245, 249, 253<br />
fresenii, Entomophthora<br />
fresenii, Neozygites<br />
fruit sucking moth 19<br />
fumosoroseus, Paecilomyces<br />
Fusarium 298<br />
Fusarium lateritium 122, 128<br />
Garcinia mangostana 262<br />
gardenia 290<br />
Geoplana caerulea see Kontikia caerulea 100<br />
glasshouse 35, 59, 66, 69, 71, 73, 77, 288, 290, 305, 315, 316<br />
glauca, Eremocitrus<br />
glauca, Nicotiana<br />
glaucum, Pennisetum<br />
Gliricidia 63<br />
Gliricidia maculata 68<br />
Gliricidia sepium<br />
glutinesa, Swinglea<br />
35, 164<br />
Glycine max 67, 219, 238, 245, 246<br />
Glycine soja 248<br />
golden gram, see Vigna aurea<br />
graminicolum, Erigonidium<br />
237<br />
granulosis virus 21, 26<br />
grape 19, 119, 199, 288--290, 306, 315<br />
grapefruit 303, 305<br />
green gram, see Vigna aurea or Vigna radiata 19, 237, 247<br />
green lacewing, see Chrysoperla carnea 43<br />
green peach aphid, see Myzus persicae 74<br />
green semi looper, see Anomis flava 18<br />
green vegetable bug, see Nezara viridula 198<br />
groundnut 35, 144, 199, 226<br />
groundnut aphid, see Aphis craccivora 34
538 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
guava 47, 108, 200, 302<br />
gypsy moth 225<br />
Habana velox 278<br />
haricot bean, see Phaseolus vulgaris<br />
hederifolia, Ipomoea<br />
237<br />
Helianthus annuus 219<br />
Hemileia vastatrix 170<br />
Hemipterotarseius 137, 138<br />
Hentzia palmarum 279<br />
Heterorhabditis 93, 94, 167, 169<br />
Heterorhabditis bacteriophora 94<br />
Heterorhabditis zealandica 94<br />
Hibiscus 60, 64, 76, 83, 137, 163<br />
Hibiscus abelmoschus 19<br />
Hibiscus cannabinus 19<br />
Hibiscus esculentus 19, 67<br />
Hibiscus manihot 19<br />
Hibiscus rosa-sinensis 19<br />
Hibiscus sabadariffa 19<br />
hirsutum cotton 19<br />
hollyhock, see Althaea rosea 19<br />
honey 226, 284, 313, 344<br />
honeydew 47, 74, 82, 132, 144, 156, 290, 303, 315<br />
horehound bug, see Agonoscelis rutila 218<br />
huanglunbin 116<br />
hyacinth bean, see Lablab niger<br />
hystrix, Citrus<br />
238<br />
Indian mynah 21, 29<br />
indica, Clausena<br />
indica, Ipomoea<br />
indicum, Sesamum<br />
indicum, Solanum<br />
insulana, Cyclosa<br />
invisa, Mimosa<br />
Ipomoea 15<br />
Ipomoea batatas 11, 19<br />
Ipomoea cairica 11
Ipomoea hederifolia 11<br />
Ipomoea indica 11<br />
Ipomoea pescapreae 11<br />
jackfruit 108<br />
jasmin orange, see Murraya paniculata 116, 117<br />
Jasminum cinnamomum 262<br />
Jasminum humile 262<br />
Jasminum sambac 262<br />
javanicus, Paecilomyces<br />
javanicus, Spicaria<br />
jute, see Hibiscus sabadariffa 19, 230<br />
kapok, see Hibiscus cannabinus 137<br />
kenaf 17, 19, 27, 30<br />
kidney bean, see Phaseolus vulgaris<br />
koenigii, Bergera<br />
koenigii, Murraya<br />
76<br />
Kontikia caerulea 100<br />
kumquat, see Fortunella 117<br />
General Index 539<br />
lablab, see Lablab niger 200, 246<br />
Lablab niger 238<br />
laburnifolia, Crotalaria<br />
lacourtianum, Dialium<br />
lanatus, Citrullus<br />
lanceolata, Vepris<br />
lansium, Clausena<br />
lateritium, Fusarium<br />
lathryoides, Macroptilium<br />
lathyroides, Phaseolus<br />
leaf mottle 128<br />
lecanii, Cephalosporium<br />
lecanii, Verticillium<br />
Leea 19<br />
lemon, see Citrus limon 116, 117, 130, 262, 289, 302, 312<br />
Leonurus sibericus 232<br />
lettuce 317, 320, 338, 340<br />
lettuce necrotic yellows 62
540 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Leucaena 163<br />
Leucaena leucocephala 164<br />
Ligustrum 163<br />
likubin 130<br />
lime, see Citrus aurantifolia<br />
limon, Citrus<br />
117, 130, 262, 274, 302, 312<br />
little red fire ant, see Wasmannia auropunctata 174<br />
looplure 319<br />
loquat 164<br />
Loranthus 262<br />
lucerne 35, 199, 338, 339, 344<br />
lunatus, Phaseolus<br />
lupin, see Lupinus 35<br />
Lupinus 35<br />
Lycopersicon esculentum 19, 187<br />
macadamia 19, 220, 222--224, 289<br />
Macroptilium atropurpureum 238<br />
Macroptilium lathryoides<br />
maculata, Gliricidia<br />
238<br />
Madagascar periwinkle, see Catharanthus roseus<br />
madurensis, Citrus<br />
116<br />
maize 14, 137, 161, 199, 320<br />
Malva 338<br />
mandarin, see Citrus reticulata 115, 117, 302<br />
Mangifera 108<br />
Mangifera indica 105, 106, 108, 109<br />
Mangifera minor 105, 108<br />
Mangifera odorata 105, 108<br />
mango, see Mangifera indica<br />
mangostana, Garcinia<br />
47, 105--112, 187, 289, 290<br />
mangosteen, see Garcinia mangostana<br />
manihot, Hibiscus<br />
262<br />
Manila hemp, see Musa textilis<br />
marcescens, Serratia<br />
marmelos, Aegle<br />
89, 137<br />
Marpissa tigrina 122, 133<br />
Mauritius papeda, see Citrus hystrix 117
General Index 541<br />
max, Glycine 35<br />
maxima var. racemosa, Citrus<br />
maxima, Citrus<br />
medic, see Medicago<br />
medica, Citrus<br />
Medicago 35<br />
Melanospora 128<br />
melon, see Citrullus lanatus 19, 65<br />
melon aphid, see Aphis gossypii<br />
melongena, Solanum<br />
46<br />
Merremia dissecta 11<br />
Merrillia caloxylon 117<br />
Metarhizium 167<br />
Metarhizium anisopliae 93, 169, 175, 334<br />
Metarrhizium brunneum 334<br />
methidathion 302<br />
methomyl 130<br />
Microcitrus australisiaca 117<br />
Mimosa invisa 134<br />
minor, Mangifera<br />
Misumenops tricuspidatus 26, 62<br />
Misumena ratia 333<br />
Mohinga 101<br />
morning glory, see Ipomoea hederifolia 11<br />
moth bean, see Vigna aconitifolia<br />
mucronata, Crotalaria<br />
11, 237<br />
mung bean, see Vigna radiata or Vigna aurea<br />
mungo, Vigna<br />
11, 219, 237, 248<br />
Murraya exotica 118, 262<br />
Murraya koenigii, see Bergera koenigii 117, 262<br />
Murraya paniculata 115--117, 127--131, 133<br />
Musa 86<br />
Musa sapientum 89<br />
Musa textilis 89<br />
muskmallow, see Hibiscus abelmoschus 19<br />
mustard 338<br />
myriacanthum, Solanum
542 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Neoaplectana carpocapsae, see Steinernema feltiae 194<br />
Neozygites fresenii 59, 69<br />
Nerium 60<br />
Nicotiana glauca<br />
niger, Lablab<br />
nigra, Ciconia<br />
nigrum, Solanum<br />
338, 339<br />
Nomuraea rileyi 169, 334, 335<br />
Nosema trichoplusiae 334<br />
octomaculatum, Theridion<br />
odorata, Mangifera<br />
oil palm<br />
okra, see Hibiscus esculentus 17, 19, 35, 47, 65, 137, 227<br />
okra semi looper, see Anomis flava 18<br />
oleander aphid, see Aphis nerii 73, 76<br />
oleander, see Nerium 60, 77<br />
orange, see Citrus sinensis 117, 199, 302, 304<br />
oriental stink bug, see Nezara antennata 233<br />
oviparae 34<br />
Oxyanthus 163<br />
Oxyopes salticus 323<br />
oxysporum, Cladosporium<br />
Paecilomyces 122, 128<br />
Paecilomyces fumosoroseus 59, 69<br />
Paecilomyces javanicus 167, 169<br />
Paecilomyces tenuipes<br />
palmarum, Dendroica<br />
169<br />
Panagrolaimus 169<br />
Pandanus 144<br />
panduratus, Phaseolus<br />
paniculata, Murraya<br />
papaya 108<br />
paprika 35, 47<br />
paraffin oil 94<br />
Pardosa 322<br />
Pardosa astrigata 63
General Index 543<br />
parsley 320<br />
Passerculus sandwichensis 333<br />
passionfruit 199, 289, 299, 300<br />
pea, see Pisum sativum 187, 199, 238, 246, 320<br />
peanut 161<br />
pear 19<br />
pearl millet, see Pennisetum glaucum 137<br />
pecan 199, 234<br />
Pennisetum americanum, see Pennisetum glaucum 137<br />
Pennisetum glaucum 137<br />
petroleum oil 119, 133, 263, 264, 286<br />
pescapreae, Ipomoea<br />
Phaseolus 11, 163, 237<br />
Phaseolus aureus 67, 247<br />
Phaseolus lunatus 164<br />
Phaseolus panduratus 238<br />
Phaseolus semierectus 238<br />
Phaseolus vulgaris 76, 219, 236--238, 245<br />
pheromone 87, 112, 137, 161, 187, 208, 217, 218, 259, 286, 289,<br />
Phidippus<br />
319<br />
333<br />
Phidippus regius 333<br />
picorna virus 201<br />
pigeon pea, see Cajanus cajan 6, 35, 64, 77--79, 236, 237<br />
pineapple, see Ananas comosus 141--156<br />
pineapple mealybug wilt 144, 145, 152, 156<br />
pineapple mealybug, see Dysmicocus brevipes 142<br />
pineapple scale, see Diaspis bromeliae 153<br />
Pisum sativum 238<br />
planarian 100<br />
plantain 89<br />
plant resistance 5, 6, 11, 88, 187, 238, 262, 320<br />
polyacrylamide gel 95<br />
polyhedrosis virus 21, 26, 317, 321, 334, 337, 338, 340<br />
pomegranate 288, 306<br />
Poncirus 116<br />
Poncirus trifoliata 117<br />
Pongamia semicarpifolia 262
544 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
potato, see Solanum tuberosum<br />
pseudocacia, Robinia<br />
47, 65, 187, 199, 289, 320<br />
prothiophos 94<br />
Psychotria 164<br />
pulvinum, Allothrombium<br />
pummelo, see Citrus maxima var. racemosa 116, 117, 260, 277<br />
pumpkin, butternut 289<br />
radiata, Vigna<br />
rape, see canola<br />
ratia, Misumena<br />
320<br />
red banded borer, see Deanolis sublimbalis 106<br />
red banded mango caterpillar, see Deanolis albizonalis 106<br />
red cotton bug, see Dysdercus cingulatus 136<br />
red seed bug, see Dysdercus cingulatus<br />
regius, Phidippus<br />
reticulata, Citrus<br />
136<br />
Rhamnus 64, 83<br />
rhodes grass scale, see Antonina graminis 155<br />
rice 144, 200, 230, 238<br />
rice bean, see Vigna calcarata 237<br />
Ricinus 19<br />
Ricinus communis 232<br />
rikettsia 334<br />
rileyi, Nomuraea<br />
Robinia pseudacacia 76<br />
robusta c<strong>of</strong>fee, see C<strong>of</strong>fea canephora<br />
rosa-sinensis, Hibiscus<br />
rosea, Althaea<br />
159, 165, 181<br />
roselle, see Hibiscus sabadariffa 19<br />
roselle cotton 18<br />
roseus, Catharanthus<br />
Rubus 163<br />
Rumex 35<br />
runner bean, see Phaseolus vulgaris 237
General Index 545<br />
sabadariffa, Hibiscus<br />
safflower 194<br />
salticus, Oxyopes<br />
sambac, Jasminum<br />
sandwichensis, Passerculus<br />
santol 108<br />
sapientum, Musa<br />
semicarpifolia, Alseodaphne<br />
semicarpifolia, Pongamia<br />
semierectus, Phaseolus<br />
sepium, Gliricidia<br />
Serratia marcescens 334<br />
sesame, see Sesamum indicum 200, 219<br />
Sesamum indicum 219<br />
Seville orange 286<br />
sho<strong>of</strong>lower, see Hibiscus rosa-sinensis<br />
sibericus, Leonurus<br />
19<br />
Sida 19<br />
sineguelas 108<br />
sinensis, Citrus<br />
sinensis, Vigna<br />
sisal 144<br />
snap bean, see Phaseolus vulgaris<br />
soja, Glycine<br />
237<br />
Solanum 187<br />
Solanum indicum 187<br />
Solanum melongena 186, 187<br />
Solanum myriacanthum 187<br />
Solanum nigrum 187<br />
Solanum tuberosum 187<br />
Solanum xanthocarpum 187<br />
sooty mould 44, 144, 300<br />
sordidin 87<br />
sorghum 5, 137, 199, 218, 230<br />
southern green stink bug, see Nezara viridula 198<br />
soybean, see Glycine max or G. soja 67, 144, 177, 199, 200, 201,<br />
218, 219, 221--224, 226, 227, 229, 231, 232, 234, 236--<br />
238, 245--248, 320
546 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
soybean stem borer, see Melanagromyza sojae 247<br />
soybean top borer, see Melanagromyza dolichostigma 247<br />
soybean, Davis 234<br />
soybean, PI 717444 200, 234<br />
sphaerosperma, Entomophthora<br />
Spicaria javanicus, see Paecilomyces javanicus<br />
Spicaria see Paecilomyces<br />
169<br />
spinach 199, 320<br />
squash 46, 47, 76<br />
squash bug, see Anasa tristis 230<br />
star apple 108<br />
Steinernema 93, 94<br />
Steinernema carpocapsae 94, 138, 194, 263<br />
Steinernema feltiae, see Steinernema carpocapsae<br />
stephanoderis, Botrytis<br />
94<br />
strawberry 69<br />
subterranean clover 35<br />
sugarbeet 337<br />
sugarcane 77, 144, 145, 152<br />
sunflower, see Helianthus anuus 65, 194, 199, 219<br />
swallow 166<br />
sweet potato, see Ipomoea batatas 9, 11, 14, 16, 19, 187<br />
sweet potato hawk moth, see Agrius convolvuli 10, 14<br />
sweet potato hornworm, see Agrius convolvuli 10, 14<br />
sweet potato weevil, see Cosmopolities sordidus 11<br />
Swinglea glutinesa 118<br />
tangerine 277<br />
taro, see Colocasia esculenta 11, 47, 64, 71, 83, 153<br />
tenuipes, Paecilomyces<br />
Tephrosia 163<br />
textilis, Musa<br />
Thelohania diazoma 334<br />
Theridion 65<br />
Theridion octomaculatum 62<br />
thuringiensis wuhanensis, Bacillus<br />
thuringiensis, Bacillus<br />
tigrina, Marpissa
General Index 547<br />
tobacco 199, 289, 320<br />
Toddalia asiatica 117<br />
tomato, see Lysopersicum esculentum 19, 187, 195, 199, 200, 210,<br />
toti virus<br />
218, 219, 223, 227, 317, 320, 335, 338, 344<br />
201<br />
Trachelas volutus 278<br />
transgenic line 320<br />
tree tobacco, see Nicotiana glauca<br />
trichoplusiae, Nosema<br />
tricuspidatus, Misumenops<br />
trifoliata, Poncirus<br />
trifoliata, Triphasia<br />
338, 339<br />
Trifolium 35<br />
Triphasia trifoliata 117<br />
tristeza virus 34, 47<br />
tristis, Acridotheres<br />
tuberosum, Solanum<br />
Tweedia coerulea 74<br />
Uloborus 65<br />
ungiculata, Vigna<br />
urd bean, see Vigna mungo 11, 237<br />
Urena 19, 137<br />
vastatrix, Hemileia<br />
venotum, Apocynum<br />
Vepris lanceolata 118<br />
Verticillium lecanii, see Cephalosporium lecanii<br />
vexillata, Vigna<br />
59, 69, 122, 128<br />
Vicia 35<br />
Vicia faba 67<br />
Vigna 237<br />
Vigna aconitifolia 237<br />
Vigna aurea 237<br />
Vigna calcarata 237<br />
Vigna mungo 11, 14, 237<br />
Vigna radiata 11, 14, 19, 219, 238<br />
Vigna sinensis 248
548 <strong>Biological</strong> <strong>Control</strong> <strong>of</strong> <strong>Insect</strong> <strong>Pests</strong>: <strong>Southeast</strong> <strong>Asian</strong> <strong>Prospects</strong><br />
Vigna unguiculata 19, 68, 219, 238, 239, 245<br />
Vigna vexillata 11<br />
Vitis 163<br />
vivapary 34<br />
vulgaris, Phaseolus<br />
watermelon 47, 320<br />
wheat 231<br />
white-bellied stork, see Ciconia nigra 16<br />
wild radish 227<br />
wild mung 11<br />
woolly apple aphid, see Eriosoma lanigerum 72<br />
xanthocarpum, Solanum<br />
Xanthomonas citri 263<br />
Xysticus croceus 62<br />
zealandica, Heterorhabditis<br />
zeylanica, Cinnamomum