Arthropods Vol. 11, No. 1, 1 March 2022 International Academy of Ecology and Environmental Sciences Arthropods ISSN 2224-4255 Volume 11, Number 1, 1 March 2022 Editor-in-Chief WenJun Zhang Sun Yat-sen University, China International Academy of Ecology and Environmental Sciences, Hong Kong E-mail: zhwj@mail.sysu.edu.cn, wjzhang@iaees.org Editorial Board Andre Bianconi (Sao Paulo State University (Unesp), Brazil) Anton Brancelj (National Institute of Biology, Slovenia) A. K. Dhawan (Punjab Agricultural University, India) John A. Fornshell (United States National Museum of Natural History, Smithsonian Institution, USA) Oscar E. Liburd (University of Florida, USA) Ivana Karanovic (Hanyang University, Korea) Lev V. Nedorezov (Russian Academy of Sciences, Russia) Enoch A Osekre (KN University of Science and Technology, Ghana) Rajinder Peshin (Sher-e-Kashmir University of Agricultural Sciences and Technology of Jammu, India) Michael Stout (Louisiana State University Agricultural Center, USA) Eugeny S. Sugonyaev (Russian Academy of Sciences, Russia) Editorial Office: arthropods@iaees.org Publisher: International Academy of Ecology and Environmental Sciences Website: http://www.iaees.org/ E-mail: office@iaees.org Arthropods, 2022, 11(1): 1-17 Article Effect of continuous rearing generations on some biological parameters of Habrobracon hebetor (Hymenoptera: Braconidae) under insectarium conditions Ghadir Momenian, Mohammad Hasan Sarayloo, Ali Afshari Department of Plant Protection, Faculty of Plant Productions, Gorgan Agricultural Sciences and Natural Resources University, Gorgan, Iran E-mail: gmomenian@gmail.com Received 30 October 2021; Accepted 5 December 2021; Published 1 March 2022 Abstract Habrabracon hebetor (Say) (Hymenoptera: Braconidae) is one of the most important biocontrol agents of insect pests in conducting IPM programs. In the present study, the effect of successive mass rearing by several generations on some biological parameters of H. hebetor was studied under laboratory conditions (28±2oC, R.H. 65±5 % & L:D (16:8)). The results of the analysis of variance showed that continuous rearing in different generations had a significant effect on all parameters (p<0.05). Eighth generation (G8) had the longest preadult stages longevity and the percentage of sex ratio with averages of 12.14 days and 66.48 (female / female+male), respectively. The longest oviposition period and highest female longevity with the averages of 18.88 and 21.88 days, respectively, was observed in the ninth generation (G9). The highest percentage of larval and pupal mortality was observed in the G1 with averages of 1.20 and 10.83%, respectively. The results also showed that the highest number of eggs laid (fecundity), pupal hatching, daily parasitism, and paralysis of larvae with averages of 14.24 eggs per female, 99.56%, 3.98 larvae, and 59.8 larvae, respectively, belonged to the fifth generation (G5). Finally, the results showed that mass rearing in continuous generations affected the biological parameters of H. hebetor and the fifth generation (G5) had the highest quality compared to other generations. Keywords Habrabracon hebetor; Continuous generations; longevity; mortality; sex ratio. Arthropods ISSN 2224­4255 URL: http://www.iaees.org/publications/journals/arthropods/online­version.asp RSS: http://www.iaees.org/publications/journals/arthropods/rss.xml E­mail: arthropods@iaees.org Editor­in­Chief: WenJun Zhang Publisher: International Academy of Ecology and Environmental Sciences 1 Introduction Effects of pesticide misuse such as increasing pressure to select pesticide-resistant populations, loss of beneficial insects, the prevalence of secondary pests, contamination of plant and livestock foods, as well as Pollution of water resources and the environment to pesticide residues has highlighted the need to use a biocontrol strategy (Kamkar and Damghani, 2013; Badran et al., 2020). Biological control or pest control IAEES www.iaees.org 2 Arthropods, 2022, 11(1): 1-17 using natural enemies is one of the most important methods proposed for sustainable agricultural development (Ahmad and Jam, 2015). Hymenopteran parasitoids serve as biological control agents for agricultural and forestry pests and as an alternative and environmentally friendly approach to overcoming the challenges of using synthetic insecticides in harvesting and post-harvesting systems (Godfray, 1994; Ueno and Ueno, 2007; Mbata and Warsi, 2019; Wang et al., 2019). Among these, after the family Ichneumonidae, the family Braconidae has been identified as the second-largest family in the order Hymenoptera with about 17,000 species whose members attack the larval stages of Lepidoptera, Coleoptera, and Diptera (Centrella et al., 2010; Belda and Riudavets, 2013; Badran et al., 2020). The Habrabracon hebetor Say (Hymenoptera: Braconidae) is a ubiquitous insect that has been reported as a larval ecto- and cumulative parasitoid of a large number of Lepidoptera (Darwish et al., 2003; Gündüz and Gülel, 2005). This parasitoid also has significant efficiency in controlling storage pests and is one of the important ectoparasitoids of storage pests, especially the family Pyralidae (Brower et al., 1996; Ba et al., 2014; Baoua et al., 2018; Mbata and Warsi, 2019). Stored product pests cause severe economic losses due to the infestation of commodities in stored grain ecosystems, including silos, bakeries, food processing industries, etc. (Brower and Press, 1990; Cline and Press, 1990; Dubey et al., 2008; El- Aziz, 2011; Mbata and Warsi, 2019). The parasitoid H. hebetor mainly attacks the larvae of Lepidopteran stored product pests including Plodia interpunctella Hübner, Ephestia kuehniella Zeller, Ephestia cautella Walker, Anagasta kuehniella Zeller, Galleria mellonella Linnaeus and Amyelois transitella Walker (Schöller, 2014; Solá Cassi, 2017; Mbata and Warsi, 2019). Some strains of H. hebetor also attack field pests such as Helichoverpa armigera Hübner, Maruca testulalis Geyer, Spodoptera litura Fabricius and Earias vittella Fabricius (Dabhi et al., 2012; Mbata and Warsi, 2019). Mass rearing systems of biocontrol agents (even under suitable conditions) can cause the declining quality of the controlling agent and, consequently, reduce its controlling power during release. Because even if the biological agent is selected correctly, but the necessary conditions for its mass rearing are not provided, its controlling power will be affected. Therefore, today, evaluating the ability of biological agents to control pests (quality control) in the success of bio-control programs has become a very important issue (Van Lenteren, 2003; Bueno et al., 2017). However, it should be noted that the quality control of each biological control agent is unique and should be evaluated with different indices. On the other hand, it is often impossible to evaluate all of these indices on a large scale. Therefore, today, some important characteristics, including biological (sex ratio, longevity, and the number of eggs laid or fecundity) and behavioral (mobility and flight) characteristics are considered as indices (Ardeh and Ghazavi, 2010). Numerous studies have been performed on the quality control of parasitoids, especially H. hebetor. Takahashi, quoted by Benson (1974), evaluated the interaction between H. hebetor and Ephestia cautella in 20 consecutive generations in a laboratory and observed adult population fluctuations of H. hebetor in generations 6, 11, and 15 (G6, 11 and 15). Benson (1974) in the study of population dynamics of H. hebetor and E. cautella in 11 consecutive generations observed fluctuation in the number of adult wasps produced in different generations. In the above study, the lowest number of adult insects, parasitism rate, and sex ratio were seen in the 6th generation (G6) and the highest number of adult insects was seen in the 9th generation (G9). Salmanova (1991) also reported a decrease in the quality of Trichogramma wasps due to the continuous rearing in the laboratory conditions. Duton and Bigler (1995) reported that the ability to fly of Trichogramma brassicae decreases with increasing number of reared generations. Rojas et al. (1995) in their study during 8 consecutive reared generations of Bracon thurberiphage on Heliothis virescens reported reduced fertility. Thomson and Hoffman (2002) in their study of the effect of 6 consecutive generations of Trichogramma carverae on its fecundity showed that fecundity is reduced by increasing the number of reared generations but IAEES www.iaees.org Arthropods, 2022, 11(1): 1-17 3 the parasitoid efficiency in the field is not reduced. Gondez and Goglel (2005), by examining the effect of H. hebetor age on fecundity and sex ratio, showed that the fecundity of H. hebetor did not change in the first five days of life, but then decreased significantly. The sex ratio of offspring on both hosts (G. mellonella and A. kuehniella) was in favor of males and the production of females was higher on G. mellonella larvae than that on A. kuehniella larvae. Pratissoli et al. (2004) reported a decrease in the viability of Trichogramma pretiosum with increasing number of rearing generations. Badran et al. (2020) reported that the second generation of H. hebetor had the longest pre-adult development time (12.4 days) and the highest fecundity (1304 eggs/female) among 20 generations reared on E. kuehniella. Ghaemmaghami et al. (2021a, b) stated that the female adult longevity of T. brassicae varied significantly between successive generations (45 generations). They also reported that the highest values of gross reproduction rate (GRR), net reproduction rate (R0), intrinsic rate of increase (r), finite rate of increase (λ) and mean generation time (T) were found in generations 5 and 10 (G5 and G10). Considering the importance of H. hebetor as one of the important agents in biocontrol strategy, this study was conducted to evaluate the effect of long-term mass rearing on the quality of H. hebetor. 2 Materials and Methods 2.1 Rearing of larvae of the Mediterranean flour moth (E. kuehniella) To achieve a sufficient and homogeneous larval colony in terms of rearing and nutritional conditions, the larvae were reared in the same laboratory conditions (25±2°C, 70±5% RH, and a photoperiod of 16:8 (L:D) h). Plastic basins with 14 cm in high and 48 cm in diameter were used for rearing, which was filled with food mixture and moth eggs to a height of 3 cm. After disinfecting the nutrient composition with Aluminium phosphide (phostoxin) tablets, the eggs were poured evenly on the nutrient composition into the basins, which included wheat flour (75%) and wheat bran (25%) (Yazdanian, 2001). The number of eggs consumed per kg of nutrient composition was 0.18 to 0.2 g of eggs. To provide the moisture needed to hatch the eggs and prevent them from drying out, the openings of the basins were covered with a thin black cotton cloth, and water was sprayed with a spray. 2.2 Collecting of adult H. hebetor In the spring, adult H. hebetor collected from rapeseed fields, which were in the flowering stage, by using funnel-shaped traps consisting of a slotted funnel and two layers of ruching. 40-50 4th and 5th larvae were placed between two layers of ruching and the set was fixed in the opening of the funnel. The traps were placed on a wooden base or rapeseed plants after preparation. 50 female wasps were collected. To eliminate environmental effects, reduce test errors, and standardize test materials, adult wasps were reared for one generation in laboratory conditions and used as the primary colony in the main experiment. 2.3 Mass rearing of H. hebetor last age larvae of A. kueniella were used to rear H. hebetor wasps. To separate the larvae from the intertwined nutrient composition, basins were placed on a gentle heat source. Then a black cloth was placed on the basins so that the cloth was in contact with the nutrient composition inside the basins. As the temperature inside the plastic basins gradually increased, the flour moth larvae began to rise out of the basins and came out of the diet. On a sheet of A4 paper, 5 of the 5th instar larvae of flour moth were placed. Each wasp was released into a clear glass with 10 cm in height and 5 cm in diameter and covered with cloth and netting. The glasses were placed upside down on the papers containing the larvae. After 24 hours, the glasses were removed and the parasitized larvae were kept in the experimental cabin under the desired conditions until adult parasitoids emerged. 2.4 Experiment IAEES www.iaees.org 4 Arthropods, 2022, 11(1): 1-17 After observing the wasps of each generation in the rearing cabin, 25 pairs (male and female) of them were randomly selected from the population inside the cabin and placed in a test tube with a length of 10 cm and aperture diameter of 1 cm for 24 hours to mate. The test tubes were blocked with cotton soaked in 5% honey water. After 24 hours, the wasps are removed from the test tubes and a pair of them are released into a disposable glass with 10 cm in height and 5 cm in diameter and covered their opening with a cloth and net. 10 larvae of Mediterranean flour moths were supplied to wasps indirectly on a paper daily, and cotton pieces soaked in 5% honey water were hung from the top of the glass to feed the wasps. Every day after counting the wasp eggs, the sheets containing the eggs were cut and transferred to plastic Petri dishes and other measurements were recorded inside them. 2.5 Parameters studied 2.5.1 Pre-adult stages longevity The number of days from egg stage to emergence of adult insects was counted in each replication and each generation and the average longevity of pre-adult stages in each generation was obtained. 2.5.2 Pre-adult stages mortality Offspring mortality at each developmental stage (eggs, larvae, pupae) was calculated and the results were expressed as a percentage. 2.5.3 Oviposition period The duration of this period was calculated by counting the number of days between the first and last oviposition of females. 2.5.4 Female and male longevity Female and male longevity was determined by counting the number of days between emergence to death time of adult wasps. Dead insects were removed from the containers and then their sex was determined. 2.5.5 Number of eggs laid (=Fecundity) The number of eggs on the larvae under each glass was examined daily at a specific time (every morning) and the number of eggs laid in each replicate was counted and recorded separately using a stereomicroscope. 2.5.6 Percentage of egg hatching After 2 days of egg hatching, the larvae were counted and this number was considered as hatched eggs. 2.5.7 Sex ratio of offspring After emerging of adult wasps developed from offspring, the number of female and male adults in each replication was counted. Finally, the total number of female adults produced was divided by the total number of adults produced by each individual, and the results were calculated as a percentage. 2.5.8 Adult emergence The total number of adult insects in each generation was counted in each replication and divided by the total number of pupae and the results were expressed as a percentage. 2.5.9 Parasitism The number of parasitized larvae in each replicate and each generation was counted every day and the total number of parasitized larvae was divided over the lifespan of the wasp and the results were considered as the average of daily parasitism per female. 2.5.10 Paralysis rate of host larvae The number of paralyzed larvae per day was counted in each generation and the total number of paralyzed larvae was divided over the lifespan of the wasp and was considered as the average of daily paralysis per female. 2.6 Statistical analysis IAEES www.iaees.org Arthropods, 2022, 11(1): 1-17 5 The present study was conducted as a completely randomized design. Analysis of variance of data for this experiment was performed by SAS statistical software (version 9.4). Also, SPSS 16 software was used to normalize the data and to draw graphs. Comparisons of means of data were performed using Duncan's multiple range test at one and five percent probability levels. Data were converted to square root to normalize them. 3 Results 3.1 Pre-adult stages longevity The results of the analysis of variance showed that successive rearing in different generations had a significant effect on the mean longevity of pre-adult stages at the level of one percent probability (Table 1). The highest longevity with averages of 12.14 and 11.82 days belonged to the eighth and tenth generations (G8 and G10), respectively, which did not differ significantly together but had significant differences with other generations (Table 2). The lowest longevity of pre-adult stages was seen in the first, third, and fourth generations (G1, G3, and G4), which were significantly different from other generations but were not significantly different together. Also, no significant difference was observed between the longevity of pre-adult stages in the second and fifth generations (G2 and G5) together and the sixth, seventh and ninth generations (G6, G7, and G9) together. 3.2 Pre-adult stages mortality The results of the analysis of variance showed that the effect of different generations on mortality of eggs, larvae, and pupae of H. hebetor was significant at the level of 1% probability (Table 1). The highest mean percentage of egg mortality was seen in the tenth generation (G10) with an average of 38.68%, which was not significantly different from that in the seventh generation (G7) at the level of one percent probability but was significantly different from the percentage of egg mortality in the other generations (Table 2). The lowest percentage of egg mortality was observed in the second generation (G2) with an average of 9.13%, which was significantly different from the percentage of mortality in the sixth, seventh, eighth, ninth, and tenth generations (G6, G7, G8, G9, and G10), but no significant difference was observed between this generation and other generations (Table 2). Also, the highest percentage of larval mortality was in the first and tenth generations (G1 and G10) with averages of 1.20 and 1.27%, which were not significantly different from each other and the fifth generation (G5), but a significant difference was observed between the first and tenth generations with other generations (Table 2). The lowest percentage of larval mortality belonged to the fourth and eighth generations (G4 and G8) with the same average of 0.82%, which did not differ significantly together but had a significant difference with the percentage of larval mortality obtained in the first, second, fifth, ninth and tenth generations (Table 2). The highest percentage of pupal mortality occurred in the first generation (G1) with an average of 10.83%, which was not significantly different from the percentage of mortality obtained in the second, third, and tenth generations (G2, G3, and G10), but was significantly different from that in other generations. The lowest percentage of pupal mortality belonged to the fifth, seventh, and sixth generations (G5, G7, and G6) with averages of 0.88, 1.35, and 1.52%, respectively, which were not significantly different together, but differed significantly with mortality obtained in other generations at a 1% probability level (Table 2). 3.3 Oviposition period Based on the results of the analysis of variance, the effect of different generations on the oviposition period of adult females was significant at the level of 5% probability (Table 1). Based on the results of the mean comparison, the longest oviposition period was recorded in the ninth generation (G9) with an average of 18.88 days, which was significantly different from the mentioned period in the first, second, third, sixth, and tenth generations, but was not significantly different from mean obtained in the other generations (Table 2). The lowest duration of this period was observed in the sixth generation (G6) with an average of 10.41 days, which IAEES www.iaees.org Arthropods, 2022, 11(1): 1-17 6 was significantly different from the average obtained in the fourth, fifth, seventh, and ninth generations, but was not significantly different from the oviposition period in other treatments (Table 2). Table 1 Results of variance analysis of the effect of long-term mass rearing (different generations) on the pre-adult stages longevity and pre-adult stages mortality, the oviposition period, and the female and male longevity of H. hebetor. Parameters Df Mean square F Pre-adult stages longevity 9 42.52 84.59** Egg 9 23.58 15.11** Larva 9 0.73 17.95** Pupa 9 13.11 20.53** Oviposition period 9 2.70 1.71* Female longevity 9 2.09 1.54* Male longevity 9 1.34 1.75* Pre-adult stages mortality * and ** indicate the significant differences between parameters obtained in different generations reared at the levels of 5% and 1% probability, respectively. 3.4 Female and male longevity The results of the analysis of variance showed that the effect of rearing generation on the female and male longevity was significant at the level of 5% probability (Table 1). The mean female longevity in the ninth generation (G9) with an average of 21.88 days was higher than that in other treatments, which was significantly different from the mean obtained in the first, second, and sixth generations at the level of 5% probability. The lowest female longevity was related to the sixth generation (G6) with an average of 13.86 days, which was significantly different from the average longevity recorded in the ninth and tenth generations (G9 and G10) but was not significantly different from averages obtained in the other generations (Table 2). The highest and lowest male longevity with averages of 17.60 and 11.16 days belonged to the tenth and first generations (G10 and G1), respectively, which had significant differences together (Table 2). The male longevity obtained in the first generation (G1) showed a significant difference with values recorded in the eighth, ninth, and tenth generations at the level of 5% probability and the longevity obtained in the tenth generation (G10) had a significant difference with those in the first, second, third and fourth generations at the level of 5% probability. IAEES www.iaees.org Arthropods, 2022, 11(1): 1-17 7 Table 2 Comparison of mean the pre-adult stages longevity and pre-adult stages mortality, the oviposition period and the female and male longevity of H. hebetor in different generation reared. Parameters Generation reared (G) G1 G2 G3 G4 G5 G6 G7 G8 G9 G10 9.86e 10.50d 9.54e 9.78e 10.62d 11.26c 11.40c 12.14a 11.60bc 11.82ab 16.42de 9.13e 16.24de 17.89de 13.12e 24.16bc d 32.76ab 23.39cd 28.52bc 38.68a 1.20a 1.00bc 0.85cd 0.82d 1.13ab 0.87cd 0.84cd 0.82d 1.03b 1.27a 10.83a 8.94ab c 9.74ab 6.72bc 0.88d 1.52d 1.35d 6.81bc 5.90c 8.12abc Oviposition period (day) 12.92bc 12.56b c 13.20bc 16.29ab 15.92ab 10.41c 16.36ab 14.72ab c 18.88a 12.32bc female longevity (day) 15.08bc 15.60b c 16.44ab c 17.00ab c 18.48ab c 13.86c 19.44ab c 19.88ab c 21.88a 20.48ab male longevity (day) 11.16c 13.24b c 13.52bc 13.60bc 14.36ab c 13.84ab c 14.76ab c 15.60ab 16.68ab 17.60a Pre-adult stages longevity (day) Pre-adult stages mortality (%) Egg Larva Pupa * Different letters in each row indicate a statistically significant difference (p <0.05). 3.5 Number of eggs laid (=Fecundity) The results of variance analysis showed that the effect of generation on fecundity and egg production of females H. hebetor was significant at the level of 1% probability (Table 3). The highest number of eggs laid was obtained in the fifth generation (G5) with a daily average of 14.24 eggs per female, which was significantly different from the values obtained in the eighth and tenth generations (G8 and G9). The lowest average number of eggs laid belonged to the eighth generation (G8) with an average of 8.43 eggs per female, which was significantly different from fecundity calculated in the third, fourth, fifth, and sixth generations, but was not significantly different from that in the other generations (Table 4). 3.6 Percentage of egg hatching Based on the results obtained from the analysis of variance, the effect of successive generations on the percentage of egg hatching was significant at the level of 1% probability (Table 3). Based on the mean comparison results, the highest mean percentage of egg hatching was related to the second generation (G2) with an average of 89.29%, which was significantly different from means obtained in the sixth, eighth, ninth, and tenth generations, but significant difference was not observed between the percentage of egg hatching in the second generation and other generations (Table 4). The lowest mean percentage of egg hatching was IAEES www.iaees.org Arthropods, 2022, 11(1): 1-17 8 related to the tenth and ninth generations (G10 and G9) with averages of 59.96 and 70.24%, respectively, which were significantly different from each other and values obtained in other generations (Table 4). 3.7 Sex ratio of offspring Based on the analysis of variance related to the sex ratio of offspring, it was found that the average percentage of sex ratio of offspring in different generations had a significant difference at the level of 1% probability (Table 3). The highest percentage of sex ratio was related to the eighth generation (G8) with an average of 66.48 (female/female+male) which was significantly different from the sex ratio calculated in the second and ninth generations (48.86 and 44.57, respectively) at the level of 1% probability. The lowest percentage of sex ratio was related to the ninth generation (G9) with an average of 44.57% (female/female+male), which was not significantly different from the values obtained in the second and fifth generations, but there was no significant difference between the percentage of sex ratio in the ninth generation and other generations (Table 4). Table 3 Results of variance analysis of the effect of long-term mass rearing (different generations) on the fecundity, percentage of egg hatching, the sex ratio, and the adult emergence of H. hebetor. Parameters Df Mean square F Fecundity (the number of eggs laid 9 3.09 3.30** Percentage of egg hatching 9 1856.38 12.42** Sex ratio of offsprings 9 1158.99 4.41** Adult emergence 9 330.77 15.63** ** indicates the significant differences between parameters obtained in different generations reared at the level of 1% probability. Table 4 Comparison of mean the fecundity, percentage of egg hatching, the sex ratio, and the adult emergence of H. hebetor in different generations reared. Parameters Generation reared (G) G1 G2 G3 G4 G5 G6 G7 G8 G9 G10 Fecundity (egg/female) 12.66ab 12.19a b 13.00a 12.79a 14.24a 13.22a 10.38ab 8.43b 12.14ab 8.44b Percentage of egg hatching (%) 85.62a 89.29a 82.66ab c 81.88ab c 86.26a 75.16cd 84.82ab 77.54bc d 70.24d 59.96e Sex ratio of offsprings (female/femal e+male) 63.20a 48.86b c 60.46ab 66.01a 57.07ab c 62.29ab 58.62ab 66.48a 44.57c 57.82ab Adult emergence (%) 89.28d 91.06b cd 90.26cd 92.56bc d 99.56a 98.50a 98.65a 93.19bc 94.60b 91.88bc d * Different letters in each row indicate a statistically significant difference (p <0.05) IAEES www.iaees.org Arthropods, 2022, 11(1): 1-17 9 3.8 Adult emergence Based on the results obtained from the analysis of variance, the effect of successive generations on the percentage of adult emergence of H. hebetor was significant at the level of 1% probability (Table 3). The highest mean adult emergence was related to the fifth, sixth, and seventh generations with averages of 99.56, 98.50, and 98.65%, respectively, which were not significantly different from each other but were significantly different from values recorded in other generations (Table 4). The lowest mean adult emergence was obtained in the first generation with an average of 89.28%, which had a significant difference with the percentage of adult emergence in the eighth and ninth generations, but a significant difference was not observed between the first generation and other generations (Table 4). 3.9 Parasitism The results obtained from the analysis of variance showed that the effect of generations reared on the daily mean parasitism was significant at the level of 5% probability (Table 5). The highest daily mean parasitism was related to the fifth generation (G5) with an average of 3.98 (larvae/day), which was significantly different from the level of parasitism in the sixth, eighth and tenth generations, but was not significantly different from parasitism rates in other generations (Table 6). The lowest daily mean parasitism belonged to the tenth generation (G10) with an average of 2.12 (larvae/day), which had a significant difference from the mean parasitism in the first, third, fourth, fifth, and ninth generations (Table 6). Table 5 Results of variance analysis of the effect of long-term mass rearing (different generations) on the parasitism and the paralysis rate of host larvae by H. hebetor. Parameters Df Mean square F Parasitism 9 8.02 4.87* Paralysis rate of host larvae 9 18.35 8.79** * and ** indicate the significant differences between parameters obtained in different generations reared at the levels of 5% and 1% probability, respectively. Table 6 Comparison of mean the parasitism and the paralysis rate of host larvae by H. hebetor in different generations reared. Parameters Generation reared (G) G1 G2 G3 G4 G5 G6 G7 G8 G9 G10 Parasitism (larvae/day) 3.25ab 3.11ab cd 3.20abc 3.35ab 3.98a 2.61bcd 3.00abc d 2.31cd 3.36ab 2.12d Paralysis rate of host larvae (larvae/day) 8.19ab 7.75ab 8.26ab 8.59a 8.72a 7.90ab 8.23ab 7.09b 8.42a 5.84c * Different letters in each row indicate a statistically significant difference (p<0.05) IAEES www.iaees.org 10 Arthropods, 2022, 11(1): 1-17 3.10 Paralysis rate of host larvae Based on the results of the analysis of variance, the effect of successive generations on the daily paralysis rate of Mediterranean flour moth larvae was significant at the level of 1% probability (Table 5). The results of the mean comparison showed that the highest mean daily paralysis of larvae was related to the fourth, fifth, and ninth generations with the averages of 8.59, 8.59, and 8.42 (larvae/day), respectively, which had a significant difference with the means obtained in the eighth and fourth generations, but did not have significant differences with each other and other generations (Table 6). The lowest mean daily paralysis of host larvae was observed in the tenth generation (G10) with an average of 5.84 (larvae/day), which was significantly different from the averages obtained in other generations (Table 6). 4 Discussion Parasitic wasps represent an alternative and environmentally friendly approach in postharvest systems for the management of pest populations because parasitoids are environmentally safe and do not negatively impact humans or beneficial organisms (Mbata and Warsi, 2019). H. hebetor is an important ectoparasitoid that has already been demonstrated to have biocontrol potentials and can regulate a wide range of stored product moth pests including P. interpunctella (Warsi et al., 2018) and E. kuehniella (Ghimire and Phillips, 2014). Lack of information on the quality and performance of natural enemies reared continuously for many generations is one of the most important issues which should be considered when establishing a biological control program, as successive mass rearing of natural enemies on factitious hosts in laboratories or insectaries may decrease the performance of the reared insects under field conditions (Bertin et al., 2017). Determining the performance of natural enemies under sequential mass rearing can provide insights into the most efficient generation of these beneficial insects and to what generation we can use natural enemies reared under these conditions without any loss in their performance (Badran et al., 2020). In the present study, we evaluated the quality of H. hebetor reared on E. kuehniella over 10 sequential generations using some biological parameters as our measurement of parasitoid quality. We found that these parameters of H. hebetor varied significantly among generations. The results of other researches also showed fluctuations in the values of biological parameters among generations reared (Badran et al., 2020; Ghaemmaghami et al., 2021a). According to the results of the present study, an increased trend was observed during the pre-adult stages longevity with increasing number of generations, although, between the first and fourth generations, the longest period of this period was related to the second generation (G2). Badran et al. (2020) reported the longest pre-adult period of H. hebetor after 20 generations reared on E. kuehniella in the second generation (G2) with an average of 12.15 days. In the study of these researchers, the duration of the pre-adult period increased from the fourth to the fifteenth generation (G4 to G15). In the present study, although the longest duration of this period was observed in the eighth generation (G8) with an average of 12.14 days, the results of the two studies were similar in the duration of this period. Also, in both studies, an increasing trend was observed during the period with increasing the number of rearing generations. The results of the present study are inconsistent with some other studies. For example, Borzoui et al. (2016) showed a decrease during the larval stage of H. hebetor reared on E. kuehniella. Also, Magro and Parra (2004) reported a shorter larval stage of H. hebetor on E. kuehniella with increasing number of generations. According to the results, mortality at the egg stage increased with increasing number of generations. At this development stage, the second and fifth generations had the lowest mortality with averages of 9.13 and 13.12%, respectively. In the larval stage, the mortality rate decreased from the first to the eighth generation (except for the fifth generation) with increasing the number of generations, but the mortality rate increased in the ninth and tenth generations. In the pupal stage, the mortality rate was high in the early generations (G1 to IAEES www.iaees.org Arthropods, 2022, 11(1): 1-17 11 G4), but this rate decreased with the increase in the number of generations to the seventh generation (G7), but it increased after the seventh generation (G7). Badran et al. (2020) showed that the survival rate of larvae and pupae increases with increasing the number of generations to the fifteenth generation (G15), in other words, the mortality rate decreases with increasing number of generations. This indicates that in insect rearing, the mortality rate in the early generations is high due to the imposition of artificial conditions, but this rate decreases after a few generations and adaptation to laboratory conditions. Based on the results of the present study, the highest and lowest mean duration of the oviposition period were observed in the ninth and sixth generations (G9 and G6), respectively. Duration of the oviposition period increased when the number of generations increased to the fourth generation, but it decreased from the fourth to the sixth generation. The duration of this period fluctuated from the sixth generation to the next, but the oviposition period in the tenth generation was shorter than that in the first generation. Badran et al. (2020) reported that the oviposition period of H. hebetor gradually decreased with increasing generations, which this trend was observed in some generations in the present study. The researchers observed the longest oviposition period in the second and fourth generations and the shortest period in the 15th and 20th generations. A decrease in the duration of the oviposition period with increasing number of generations was also reported in other studies. For example, Ghaemmaghami et al. (2021a) stated that the oviposition period of T. brassicae on S. cerealella increased from 4.65 days in the fifth generation to 2.55 days in the 45th generation. Lü et al. (2017) also showed that the biological parameters of Trichogramma dendrolimi decreased after 20 generations. The results showed that the highest and lowest mean female longevity was observed in the ninth and sixth generations, respectively, and for male longevity in the tenth and first generations, respectively. The study of this parameter in the 10 generations studied shows that the female and male longevity increased with increasing number of generations. The results of other researchers are different from the results of the present study. Badran et al. (2020) showed that female longevity decreased from the second generation (G2) to the twentieth generation (G20). Host size, age, and species can affect parasitoid longevity (Milonas, 2005; Charles and Paine, 2016). Badran et al. (2020) stated that the quality of Ephestia larvae may decrease with successive rearing, which may reduce the longevity of H. hebetor. The researchers also reported that the initial egg density of the parasitoid could affect the longevity of the female. An increase in the number of eggs laid by the female during rearing may decrease female longevity (Milonas, 2005; Charles and Paine 2016). These results may have been inconsistent due to the differences in the populations studied, the number of rearing generations, and the experimental conditions. The results showed that the highest and lowest number of eggs laid occurred in the fifth and eighth generations, respectively. The findings of the present study showed that the reproduction rate of this parasitoid increased from the first to the fifth generation and then decreased. This result is consistent with the results of Attaran (1996) on H. hebetor, Benson (1974) on H. hebetor, Takahashi quoted by Benson (1974) on H. hebetor, Gündüz and Gülel (2005) and Jooyandeh (2008) on H. hebetor, Rojas et al. (1995) on Bracon thurberiphage, Thomson and Hoffman (2002) on Trichogramma carverae. However, it was different from the results of Badran et al. (2020) on H. hebetor and Ghaemmaghami et al. (2021a, b) on T. brassicae. Badran et al. (2020) reported that the highest and lowest fecundity occurred in the second and twentieth generations, respectively, and the reproduction rate decreased with increasing number of generations. The fecundity obtained in the present study (8.43-24.24 eggs/female) was less than the values obtained in the study of Badran et al. (2020) with an average of 49.98 -136.07 eggs/female. Ghaemmaghami et al. (2021b) reported that the highest and lowest fecundity of T. brassicae were found in G5 (47.41 eggs/female) and G45 (20.25 eggs/female), respectively. Amir-Maafi and Chi (2006) and Farag et al. (2015) reported the average of fecundity 78.3 eggs/female and 6.9 eggs/female. Nasab et al. (2014) estimated the mean fecundity of this IAEES www.iaees.org 12 Arthropods, 2022, 11(1): 1-17 parasitoid on E. kuehniella between 14.94-24.46 egg/female, while the fecundity on hosts reared on wheat, soybeans, and wheat + soybean decreased over three generations, but this parameter increased on hosts reared on barley and rice over three generations. The discrepancy between the present study and the mentioned research could be due to Wolbachia infection, the difference in the duration of the oviposition period, using feral populations of H. hebetor with different genetic patterns instead of laboratory or insectary lines, using mature pyralid larvae as hosts in the experiment or supplying adult wasps with honey as a carbohydrate source (Ghimire and Phillips, 2010, 2014; Bagheri et al., 2019). Also, the decrease of host genetic diversity of E. kuehniella due to inbreeding over successive rearing on a standard diet may reduce host quality and thus lead to a reduction in H. hebetor fecundity over generations (Bertin et al., 2017). Based on the results of the mean comparison, the highest mean percentage of egg hatching was observed in the second generation (G2) and the lowest mean percentage of egg hatching was recorded in the tenth and ninth generations (G10 and G9). Also, the percentage of egg hatching over 10 generations had a decreasing trend despite observing fluctuations. Badran et al. (2020) reported that the gross reproduction rate (GRR) and net reproduction rate (NRR) of H. hebetor, which represent the number of offspring produced per person excluding mortality and normal mortality, respectively, increased by increasing the number of generations to the tenth generation, which was different from the results of the present study. On the other hand, Ghaemmaghami et al. (2021a) stated that the values of the above two parameters for T. brassicae decreased with increasing number of generations, which is consistent with the results of the present study. Sex ratio is an economically important trait in parasitoids that affects the financial profitability of mass rearing (Badran et al., 2020; Ghaemmaghami et al., 2021a). The highest and lowest sex ratio was related to the eighth generation (G8) and the ninth generation (G9), respectively. The results showed that the sex ratio decreased in the ninth and tenth generations compared to the first generation (G1). Badran et al. (2020) reported that the sex ratio (female/female+male) in H. hebetor under successive rearing showed an increasing trend from the second generation (17%) to the tenth generation (60%) and then this ratio gradually decreased after the tenth generation. They also reported some fluctuations in the sex ratio of H. hebetor under successive rearing. Parasitoids reared for many successive generations usually show some fluctuations in sex ratio (Wylie, 1979). In the present study, fluctuations in sex ratio were observed during ten generations. Mahdi Nasab et al. (2014) reported the increased trend in the sex ratio of H. hebetor on E. kuehniella reared on wheat and barley over three generations and decreased trend on rice. The researchers also confirmed some fluctuations in sex ratio over three rearing generations. Ghaemmaghami et al. (2021a) also found some fluctuations in the sex ratio of T. beassicae during long-term mass rearing on S. cerealella. The above studies as well as the findings of Pratissoli et al. (2004) and Lü et al. (2015) who showed fluctuations in the sex ratio of parasitoids experiencing successive rearing is in the agreement with the results of the present study. The association of some bacterial endosymbionts, such as Wolbachia, with H. hebetor could be the reason for more female production in H. hebetor. The important role of Wolbachia in shifting the sex ratio towards more female production in parasitoids has been confirmed by various researchers (Rousset et al., 1992; Pintureau et al., 2000; Tagami et al., 2001; Karimi et al., 2012; Weinert et al., 2015; Badran et al., 2020; Ghaemmaghami et al., 2021a). The results of the present study showed that the highest mean adult emergence was related to the fifth, sixth, and seventh generations, and the lowest mean was observed in the first generation. According to the results, the percentage of adult emergence increased up to the fifth generation (G5). The highest percentage of adult emergence was seen in the fifth generation, but then gradually decreased. This result was similar to the results of Prezotti et al. (2004) on T. pretiosum, Nordlund et al. (1997) on T. minutum. Taghikhani et al. (2019) recorded the highest and lowest adult emergence of T. brassicae for the fifth and second generations, IAEES www.iaees.org Arthropods, 2022, 11(1): 1-17 13 respectively in the laboratory. They stated that the highest rate of adult emergence was recorded for the first three generations in the insectarium, while the last five generations showed a significant decrease in the rate of adult emergence. The researchers reported that the emergence rate of insectary-reared wasps was lower than the rate obtained in the laboratory, which may highlight the unsuitability of the rearing method, the factitious host, and/or the climatic conditions. According to Soares et al. (2012), the emergence rate of parasitoid wasps, especially Trichogramma, may depend on the size and quality of the host egg, the number of parasitoids that develop per egg, and temperature. According to the results, the highest and lowest daily mean parasitism belonged to the fifth and tenth generations, respectively. The results of the parasitism showed that the rate of parasitism increased to the fifth generation and then decreased with increasing generation. This result was similar to the results of Benson (1974) on H. hebetor and Prezotti et al. (2004) on T. pretiosum but was different from the results of Nordlund et al. (1997) on T. minutum. Ghaemmaghami et al. (2021a) reported that the rate of T. brassicae parasitism decreased with increasing number of rearing generations. Taghikhani et al. (2019) stated that there was a significant difference in the rate of parasitism among the generations of T. brassicae. The maximum rate of parasitism was observed in the third generation (G3), followed by a decreasing trend until the eighth generation (G8). The results of the mean comparison showed that the highest mean daily paralysis of larvae was recorded for the fourth and fifth generation (G4 and G5), while the lowest was observed in the tenth generation (G10). According to the results, the paralysis rate of larvae increased from the first to the fifth generation, followed by fluctuations, so that it reached its lowest value in the tenth generation (G10). Our results showed that a lack of information about the consequences of long-term mass rearing of H. hebetor may lead to the failure of biological control programs. We also showed that mass rearing in continuous generations had an effect on the biological parameters of H. hebetor and the fifth generation (G5) had the highest quality compared to other generations. Due to reduced female fecundity, after these generations, the colony needs to be rejuvenated by adding natural individuals. Acknowledgments We would like to thank Eng. Kurdi (the respectable director of the insectarium of Gorgan city) and all the colleagues of this center who helped us in carrying out this research. References Ahmad N, Jam NA. 2015. Insect pest management in conservation agriculture. In: Conservation Agriculture (Farooq M, Siddique K, eds). 133-155, Springer, Dordrecht, Netherlands Amir-Maafi M, Chi H. 2006. Demography of Habrobracon hebetor (Hymenoptera: Braconidae) on Two Pyralid Hosts (Lepidoptera: Pyralidae). Annals of the Entomological Society of America, 99(1): 84-90 Ardeh MJ, Ghazavi M. 2010. Principle of biocontrol for greenhouses production. Iranian Research Institute of Plant Protection Press, Tehran, Iran Attaran MR. 1996. Effects of laboratory hosts on the biological characteristics of the parasitoid Habrabracon hebetor. MSc Thesis, Faculty of Agriculture, Tarbiat Modares University, Iran Ba MN, Baoua IB, Kabore A, Amadou L, Oumarou N, Dabire-Binso C, Sanon A. 2014. Augmentative onfarm delivery methods for the parasitoid Habrobracon hebetor Say (Hymenoptera: Braconidae) to control the millet head miner Heliocheilus albipunctella (de Joannis) (Lepidoptera: Noctuidae) in Burkina Faso and Niger. BioControl, 59(6): 689-696 IAEES www.iaees.org 14 Arthropods, 2022, 11(1): 1-17 Badran F, Fathipour Y, Bagheri AN, Attaran MR, Reddy GVP. 2020. Effect of prolonged mass rearing on life history traits of Habrobracon hebetor (Hymenoptera: Braconidae). International Journal of Pest Management, 1-10 Bagheri Z, Talebi AA, Asgari S, Mehrabadi M. 2019. Wolbachia induce cytoplasmic incompatibility and affect mate preference in Habrobracon hebetor to increase the chance of its transmission to the next generation. Journal of Invertebrate Pathology, 163: 1-7 Baoua IB, Ba MN, Amadou L, Kabore A, Dabire-Binso CL. 2018. Field dispersal of the parasitoid wasp Habrobracon hebetor (Hymenoptera: Braconidae) following augmentative release against the millet head miner Heliocheilus albipunctella (Lepidoptera: Noctuidae) in the Sahel. Biocontrol Science and Technology, 28(4): 404-415 Belda C, Riudavets J. 2013. Natural enemies associated with lepidopteran pests in food and feed processing companies. Journal of Stored Products Research, 53: 54-60 Benson JF. 1974. population dynamics of Bracon hebetor say (Hym: Braconidae) and Ephestia cautella walker (Lep: Phycitidae) in a laboratory ecosystem. Journal of Animal Ecology, 43(1): 71-86 Bertin A, Pavinato VAC, Parra GRP. 2017. Fitness-related changes in laboratory populations of the egg parasitoid Trichogramma galloi and the implications of rearing on factitious hosts. BioControl, 62(4): 435444 Borzoui E, Naseri B, Mohammadzadeh-Bidarani M. 2016. Adaptation of Habrobracon hebetor (Hymenoptera: Braconidae) to Rearing on Ephestia kuehniella (Lepidoptera: Pyralidae) and Helicoverpa armigera (Lepidoptera: Noctuidae). Journal of Insect Science, 16(1): 12-17 Brower JH, Press JW. 1990. Interaction of Bracon hebetor (Hymenoptera: Braconidae) and Trichogramma pretiosum (Hymenoptera: Trichogrammatidae) in suppressing stored-product populations in small in shell peanut storages. Journal of Economic Entomology, 83(3): 1096-1101 Brower JH, Smith L, Vail PV, Flinn PW. 1996. Biological control. In: Integrated Management of Insects In Stored Products (Subramanyam B, Hagstrum DW, eds). 223-286, Marcel Dekker, New York, USA Bueno E, Romero A, Osorio I, Zaviezo T. 2017. Laboratory rearing methods that promote inbreeding have a negative impact on the fitness of Mastrus ridens Horstmann (Hymenoptera: Ichneumonidae), a parasitoid used to control the codling moth. Chilean Journal of Agricultural Research, 77(4): 413-419 Carabajal Paladino L, Papeschi A, Lanzavecchia S, Cladera J, Bressa MJ. 2013. Cytogenetic characterization of Diachasmimorpha longicaudata (Hymenoptera: Braconidae), a parasitoid wasp used as a biological control agent. European Journal of Entomology, 110(3): 401-409 Centrella ML, Shaw SR. 2010. A new species of phytophagous braconid Allorhogas minimus (Hymenoptera: Braconidae: Doryctinae) reared from fruit galls on Miconia longifolia (Melastomataceae) in Costa Rica. International Journal of Tropical Insect Science, 30(2): 101-107 Charles JJ, Paine TD. 2016. Fitness effects of food resources on the polyphagous aphid parasitoid, Aphidius colemani Viereck (Hymenoptera: Braconidae: Aphidiinae). PLoS One, 11(1): e0147551 Cline DL, Press JW. 1990. Reduction in almond moth (Lepidoptera: Pyralidae) infestations using commercial packaging of foods in combination with the parasitic wasp, Bracon hebetor (Hymenoptera: Braconidae). Journal of Economic Entomology, 83(3): 1110-1113 Dabhi MR, Korat DM. 2010. Copulation and paralyzing behaviour of Bracon hebetor Say. Insect Environment, 16(2): 58 IAEES www.iaees.org Arthropods, 2022, 11(1): 1-17 15 Darwish E, El-Shazly M, El-Sherif H. 2003. The choice of probing sites by Bracon hebetor Say (Hymenoptera: Braconidae) foraging for Ephestia kuehniella Zeller (Lepidoptera: Pyralidae). Journal of Stored Products Research, 39 (3): 265-276 Dubey NK, Srivastava B, Kumar A. 2008. Current status of plant products as botanical pesticides in storage pest management. Journal of Biopesticides, 1: 182-186 Dutton A, Bigler F. 1995. Flight activity assessment of the egg parasitoid Trichogramma brassicae (Hym.: Trichogrammatidae) in laboratory and field condition. Entomophaga, 40(2): 223-233 El-Aziz SE. 2011. Control strategies of stored product pests. Journal of Entomology, 8: 101-122 Farag NA, Ismail IA, Elbehery HHA, Abdel-Rahman RS, Abdel-Raheem MA. 2015. Life table of Bracon hebetor say. (Hymenoptera: Braconidae) reared on different hosts. International Journal of ChemTech Research, 8(9): 123-130 Ghaemmaghami E, Fathipour Y, Bagheri A, Talebi AA, Reddy GVP. 2021a. Quality control of the parasitoid wasp Trichogramma brassicae (Hymenoptera: Trichogrammatidae) over 45 generations of rearing on Sitotroga cerealella. Insect Science, 28: 180-190 Ghaemmaghami E, Fathipour Y, Bagheri A, Talebi AA, Reddy GVP. 2021b. Continuous rearing on Ephestia kuehniella reshaped quality of the parasitoid wasp Trichogramma brassicae (Hymenoptera: Trichogrammatidae). Journal of Asia-Pacific Entomology, 24: 166-174 Ghimire MN, Phillips TW. 2010. Mass rearing of Habrobracon hebetor Say (Hymenoptera: Braconidae) on larvae of the Indian meal moth, Plodia interpunctella (Lepidoptera: Pyralidae): effects of host density, parasitoid density, and rearing containers. Journal of Stored Products Research, 46(4): 214-220 Ghimire MN, Phillips TW. 2014. Oviposition and reproductive performance of Habrobracon hebetor (Hymenoptera: Braconidae) on six different pyralid host species. Annals of the Entomological Society of America, 107: 809-817 Godfray HCJ. 1994. Parasitoids: behavioral and evolutionary ecology. Princeton University Press, Princeton, NJ, USA Gunduz EA, Gulel A. 2005. Investigation of fecundity and sex ratio in the parasitoid Bracon hebetor Say (Hym: Braconidae) in relation to parasitoid age. Turkish Journal of Zoology, 29: 291-294 Jooyandeh A. 2008. Evaluation of the effect of parasitoid age, Habrabracon hebetor Say. (Hym: Braconidae) on its fecundity rate and sex ratio. Proceeding of the 18th Iranian Plant Protection Congress, August 24 2008, Bu-Ali Sina University, Hamadan, Iran Kamkar B, Mahdavi Damghani AM. 2013. Principles of Sustainable Agriculture. Ferdowsi University Press, Mashhad, Iran Karimi J, Darsouei R, Hosseini M, Stouthamer R. 2012. Molecular characterization of Iranian Trichogrammatids (Hymenoptera: Trichogrammatidae) and their Wolbachia endosymbiont. Journal of Asia-Pacific Entomology, 15: 73-77 Lü X, Han S, Li L. 2015. Biochemical analyses of Trichogramma dendrolimi (Hymenoptera: Trichogrammatidae) in vitro and in vivo rearing for 10 generations. Florida Entomologist, 98: 911-915 Lü X, Han S, Li Z, Li L. 2017. Biological characters of Trichogramma dendrolimi (Hymenoptera: Trichogrammatidae) reared in vitro versus in vivo for thirty generations. Scientific Reports, 7: 17928 Magro SR, Parra JRP. 2004. Comparison of artificial diets for rearing Bracon hebetor Say (Hymenoptera: Braconidae). Biological Control, 29(3): 341-347 Mahdi Nasab Z, Faal-Mohammad-Ali H, Shishehbor P. 2014. The effect of diets of Ephestia kuehniella Zeller on biological characteristics and life table of Habrobracon hebetor (Hymenoptera: Braconidae) in laboratory condition. Journal of Plant Protection Research, 37(3): 82-96 IAEES www.iaees.org 16 Arthropods, 2022, 11(1): 1-17 Mbata GN, Warsi S. 2019. Habrobracon hebetor and Pteromalus cerealellae as tools in the postharvest integrated pest management. Insects, 10: 85 Milonas PG. 2005. Influence of initial egg density and host size on the development of the gregarious Barcon hebetor on three different host species. Biocontrol, 50(3): 415-428 Nordlund DA, Wu ZX, Greenberg SM. 1997. In vitro rearing of Trichogramma minutum Riley (Hymenoptera: Trichogrammatidae) for ten generations with quality assessment comparisons of in vitro and in vivo reared adults. Biological Control, 9: 201-207 Pintureau B, Grenier S, Boleat B, Lassabliere F, Heddi A, Khatchadourian C. 2000. Dynamics of Wolbachia populations in transfected lines of Trichogramma. Journal of Invertebrate Pathology, 76: 20-25 Pratissoli D, Oliveira HN, Goncalves JR, Zanuncio JC, Holtz AM. 2004. Changes in biological characteristics of Trichogramma pretiosum (Hym: Trichogrammatidae) reared on eggs of Anagasta kuehniella (lep: pyralidae) for 23 generations. Biocontrol Science and Technology, 14(3): 313-319 Prezotti L, Para JRP, Vencovsky R, Coelho ASG, Cruz I. 2003. Effect of the size of the founder population on the quality of sexual population on the quality of sexual population of Trichogramma pretiosum in laboratory. Biological Control, 30: 174-180 Rojas MG, Vinson SB, Williams HJ. 1995. Supplemental feeding increase the utilization of a factititous host for rearing Bracon thurberiphagae Muesebeck (Hymenoptera: Braconidae) a parasitoid of Anthonomus grandis (Coleoptera: curculionidae). Biological Control, 5: 591-597 Rousset F, Bouchon D, Pintureau B, Juchault P, Solignac M. 1992. Wolbachia endosymbionts responsible for various alterations of sexuality in arthropods. Proceeding of the Royal Society B: Biological Sciences, 250: 91-98 Salmanova LM. 1991. Changes of Trichogramma cultures in permanent rearing on the labolatory host Sitotroga cerealella Oliv. Dissertation biological faculty of Lomonosow University, Moscow, Russia SAS Institute. 2013. PROC User’s Manual, Version 9.4. SAS Institute, Cary, NC, USA Schöller M. 2010a. Prospects for biological control of stored-product pests. Julius-Kühn-Archiv, 429: 25-31 Schöller M. 2010b. Biological control of stored-product insects in commodities, food processing facilities and museums. 10th International Working Conference on Stored Product Protection, Julius-Kühn-Archiv, 425: 596-606 Soares MA, Leite GLD, Zanuncio JC, Sá VGMD, Ferreira CS, Rocha SL, Pires EM, Serrão JE. 2012. Quality control of Trichogramma atopovirilia and Trichogramma pretiosum (Hym.: Trichogrammatidae) adults reared under laboratory conditions. Brazilian Archives of Biology and Technology, 55(2): 305-311 Solá Cassi M. 2017. Approaches for the biological control of stored product pests. PhD thesis, Universitat Autonoma de Barcelona, Spain SPSS-Inc. 2009. PASW Statistics for Windows (16.0 ed). SPSS Inc, Chicago, USA Tagami Y, Miura K, Stouthamer R. 2001. How does infection with parthenogenesis-inducing Wolbachia reduce the fitness of Trichogramma? Journal of Invertebrate Pathology, 78: 267-271 Taghikhani N, Fathipour Y, Bagheri AN, Talebi AA. 2019. Generation-dependent fitness of the parasitoid wasp Trichogramma brassicae (Hymenoptera: Trichogrammatidae) in laboratory and insectarium condition. Journal of Crop Protection, 8(3): 351-359 Thomson LJ, Hoffman AA. 2002. Laboratory Fecundity as Predictor of Field Success in Trichogramma carverae (Hymenoptera: Trichogrammatidae). Journal of Economic Entomology, 95(5): 912-917 Ueno T, Ueno K. 2007. The effects of host-feeding on synovigenic egg development in an endoparasitic wasp, Itoplectis naranyae. Journal of Insect Science, 7: 46 IAEES www.iaees.org Arthropods, 2022, 11(1): 1-17 17 Van Lenteren JC. 2003. Quality Control and Production of Biological Control Agent: Theory and Testing Procedures. CABI, UK Wang ZZ, Liu YQ, Shi M, Huang JH, Chen XX. 2019. Parasitoid wasps as effective biological control agents. Journal of integrative agriculture, 18(4): 705-715 Warsi S, Mbata GN, Payton ME. 2018. Improvement of reproductive performance of Habrobracon hebetor: Consideration of diapausing and non-diapausing larvae of Plodia interpunctella. Biological Control, 118: 32-36 Weinert LA, Araujo-Jnr EV, Ahmed MZ, Welch JJ. 2015. The incidence of bacterial endosymbionts in terrestrial arthropods. Proceeding of the Royal Society B: Biological Sciences, 282: 1-6 Wylie HG. 1979. Sex ratio variability of Muscidifurax zaraptor (Hymenoptera: Pteromalidae). The Canadian Entomologist, 111(1): 105-109 Yazdanian M. 2001. Evaluation of the developmental rate and fecundity of the Mediterranean flour moth, Ephestia kuehniella Zeller on several diets prepared from wheat bran flour in dry and wet form. MSc Thesis in Agricultural Entomology. Faculty of Agriculture, University of Tabriz, Tabriz, Iran IAEES www.iaees.org Arthropods, 2022, 11(1): 18-55 Article Diversity of spiders (Chelicerata: Araneae) in Uttar Pradesh and Uttarakhand, India Rajendra Singh, Garima Singh Department of Zoology, Deendayal Upadhyaya University of Gorakhpur, Gorakhpur, Uttar Pradesh, India Email: rsinghgpu@gmail.com Received 9 October 2021; Accepted 20 November 2021; Published 1 March 2022 Abstract An updated checklist of faunal biodiversity of the spiders, in two norther states of India, Uttar Pradesh and Uttarakhand is presented herewith. A total of 520 species of spiders described under 236 genera belonging to 50 families were recorded in both the states of north India. The biodiversity of spiders is more in Uttar Pradesh (284 species, 146 genera, 36 families) than Uttarakhand (373 species, 202 genera, 45 families). However, most of the areas in both the states are still virgin regarding the faunal survey programmes and need intensive and extensive survey in those areas by enthusiastic workers. Keywords Uttar Pradesh; Uttarakhand; spiders; Araneae; checklist; faunal distribution. Arthropods ISSN 2224­4255 URL: http://www.iaees.org/publications/journals/arthropods/online­version.asp RSS: http://www.iaees.org/publications/journals/arthropods/rss.xml E­mail: arthropods@iaees.org Editor­in­Chief: WenJun Zhang Publisher: International Academy of Ecology and Environmental Sciences 1 Introduction Spiders (Arachnida: Araneae) are one of the most hatred chelicerate arthropods by the humans in spite of their significant role in keeping down the insect pest population in agriculture as they lavishly feed on insect pests. The position of order Araneae is seventh in global diversity of animals after the five largest insect orders (Coleoptera, Lepidoptera, Hymenoptera, Diptera, Hemiptera) and one arachnid order (Acari) in terms of species diversity (Sharma et al., 2020). The world spider catalog (WSC, 2021) took account of 49,783 species in 4234 genera belonging to 129 families. Out of them, only 1877 species belonging to 479 genera in 60 families are reported in India (Caleb and Sankaran, 2021), though in recent updates, 2344 species under 596 genera grouped into 65 families are recorded in India (Singh and Singh, 2021a). In this list, several species were considered cases of misidentification by the authors. However, there exist many species in the wild and museums that still await description and classification. Despite recent research works on the diversity and distribution of spiders in India, their number is insufficient as compared to the other parts of the world. For the sustainable management and conservation of biodiversity of the animal species of any region of the world, their proper documentation is vital as it helps in monitoring the rate of loss of species. Preparation of checklist of species is an essential component of systematic documentation. Hence, in view of increasing IAEES www.iaees.org Arthropods, 2022, 11(1): 18-55 19 intensity of anthropogenic threats to biodiversity, a cataloguing and appropriate documentation of biodiversity, especially on ignored groups like spiders, is desirable immediately. So far, very few attempts were made to study the spider fauna of two north Indian states, Uttar Pradesh and Uttarakhand. For continuation of state fauna series of spiders of India (Singh and Singh, 2021b, c, d), in the present article the updated checklist of spider fauna of abovementioned two north Indian states is provided. Family-wise placement and the latest zoological names are mentioned according to WSC (2021). 2 Materials and Methods The present checklist is based on the published literature on the spiders recorded from these two north Indian states, Uttar Pradesh and Uttarakhand, in recent past books, book chapters, journals, proceedings, few authentic theses, websites, and World Species Catalog (WSC, 2021) up to November 25, 2021. In most of the literature published earlier, several errors crept in the scientific names of the spiders even in the recent publications. It happens because such contents become outdated quickly and, due to their perceived comprehensiveness, readers sometimes overlook newer sources of data. The spider taxonomists are continually describing new taxa, modifying their status, and other nomenclatural decisions and clarifications (WSC, 2021; Singh and Singh, 2020). In the present checklist, the scientific names of the spiders are corrected following WSC (2021). If a spider species is identified only up to a generic level, it was considered as species only if no other species of that genus is reported within that state. In few cases, the locations of spider species are corrected, particularly of those spiders that were described/recorded during the British period and even after the independence of India (1947) till the carving of Uttarakhand from Uttar Pradesh. For their synonymy, WSC (2021) may be consulted. 3 Results and Discussion A total of 520 species of spiders were reported from these two northern states out of which 113 species were recorded in both states. A total of 284 species of spiders belonging to 146 genera and 36 families were reported from Uttar Pradesh while 373 species of spiders described under 202 genera belonging to 45 families were recorded from Uttarakhand. Four families of spiders (Barychelidae, Desidae, Ischnothelidae, Stenochilidae) recorded in Uttar Pradesh were not reported from Uttarakhand. Similarly, 13 families of spiders (Anyphaenidae, Atypidae, Cybaeidae, Filistatidae, Idiopidae, Mimetidae, Nesticidae, Oecobiidae, Palpimanidae, Pimoidae, Psechridae, Segestriidae, Trochanteriidae) recorded from Uttarakhand were not reported from Uttar Pradesh. However, most of the areas in both states are still virgin regarding the faunal survey programmes and need an intensive and extensive survey programmes in those areas by enthusiastic workers. 3.1 Uttar Pradesh Uttar Pradesh (Coordinates: 26.8467° N, 80.9462° E) is one of the largest north Indian states and was created in 1937 as United Province and renamed Uttar Pradesh in 1950. In 2000, a new state Uttarakhand (previously Uttaranchal) was carved from the Himalayan hill region of the state. Uttar Pradesh is presently divided into 75 districts (Table 1, Fig. 1). Uttar Pradesh is bordered by Bihar to the east, Rajasthan to the west, Haryana, Himachal Pradesh and Delhi to the northwest, Uttarakhand and Nepal to the north, Madhya Pradesh and Chhattisgarh to the south, and Jharkhand to the southeast (Fig. 1). Uttar Pradesh is the fourth-largest state of India by area (243,290 km2). The state is inundated by several rivers, prominently two rivers, the Ganges and Yamuna, other prominent rivers are Betwa, Gomti, Rapti and Saryu (Ghaghra). North of the state is bordered by the Himalayas having high mountains, but the larger areas are Gangetic Plain including the Ganga-Yamuna Doab, the Ghaghra plains, the Ganges plains and the Terai. In the south, there is a smaller IAEES www.iaees.org Arthropods, 2022, 11(1): 18-55 20 Vindhya Range and plateau region characterized by hard rock strata and varied topography of hills, plains, valleys and plateaus. The Terai area is covered with tall elephant grass and thick forests mixed with marshes and swamps. The forest cover is only 6.9% while the cultivable area is 82%. The flora and fauna of the northern belt of Uttar Pradesh is highly rich as compared with the southern part. Summers are extreme with temperatures fluctuating anywhere between 0 °C and 50 °C in parts of the state along with dry hot winds. The Gangetic plain varies from semiarid to sub-humid and the mean annual rainfall ranges from 65 cm in the southwest corner of the state to 100 cm in the eastern and the southeastern parts of the state. Table 1 Number of species of spiders recorded in different districts of Uttar Pradesh. Code Districts No. of Code Districts species AG Agra AL No. of Code Districts species No. of species 93 FR Farrukhabad 1 MG Maharajganj Aligarh 0 FT Fatehpur 0 MH Mahoba 0 AM Amethi 0 GB G. Buddha Nagar 1 MI Mirzapur 2 AN Ambedkar Nagar 0 GN Gonda 61 MO Moradabad 0 AR Amroha 1 GP Ghazipur 1 MP Mainpuri 0 AU Auraiya 0 GR Gorakhpur 57 MT Mathura 63 AY Ayodhya 3 GZ Ghaziabad 0 MU Muzaffarnagar 0 AZ Azamgarh 20 HA Hapur 0 PG Prayagraj 9 BB Barabanki 1 HM Hamirpur 0 PI Pilibhit 0 BD Badaun 0 HR Hardoi 0 PR Pratapgarh 0 BG Bagpat 0 HT Hathras 0 RA Rampur 0 BH Bahraich 0 JH Jhansi 0 RB Rae Bareli 0 BI Bijnor 2 JL Jalaun 0 SA Saharanpur 9 BL Ballia 1 JU Jaunpur 11 SH Shamli 0 BN Banda 0 KD Kanpur Dehat 2 SI Sitapur 0 BP Balrampur 0 KG Kasganj 0 SJ Shahjahanpur 1 BR Bareilly 1 KJ Kannauj 0 SK St. Kabir Nagar 0 BS Basti 0 KN Kanpur Nagar 7 SM Sambhal 3 BU Bulandshahr 0 KS Kaushambi 0 SN Siddharthnagar CD Chandauli 12 KU Kushinagar 57 SO Sonbhadra 0 CT Chitrakoot 1 LA Lalitpur 0 SR St. Ravidas Nagar 0 DE Deoria 42 LK Lakhimpur Kheri 178 SU Sultanpur 0 ET Etah 0 LU Lucknow 61 SV Shravasti 0 EW Etawah 0 MB Mau 4 UN Unnao 49 FI Firozabad 0 ME Meerut 4 VA Varanasi 28 IAEES 61 61 www.iaees.org Arthropods, 2022, 11(1): 18-55 21 Fig. 1 Number of species of spiders described/recorded from different districts of Uttar Pradesh. District codes are given in Table 1. Regarding the records of spiders in Uttar Pradesh, Blackwall (1867) was probably the first who had described three species (Crossopriza lyoni, Drassodes delicatus, Hippasa greenalliae) and reported one species (Artema atlanta Walckenaer, 1837) from Meerut and Agra region. Later, Pocock (1900, 1901) described one species (Oxyopes ryvesi Pocock, 1901) from Prayagraj and recorded other 6 species from Bareilly (Selenops radiatus Latreille, 1819), Meerut (Artema atlanta Walckenaer, 1837; Crossopriza lyoni (Blackwall, 1867)) and Prayagraj (Cyrtophora cicatrosa (Stoliczka, 1869); Hersilia savignyi Lucas, 1836; Palystes flavidus Simon, 1897) and Shahjahanpur (Chilobrachys hardwickei (Pocock, 1895)) districts of Uttar Pradesh. Among the Indian authors during post-independent period, Basu (1965) was the first to describe a species of Thomisidae, Pistius bhadurii from Saharanpur district of Uttar Pradesh. Later on, many authors have reported several species from different districts of Uttar Pradesh. Regarding the faunal survey, Hore and Uniyal (2008a, b, c) and Uniyal and Hore (2009) recorded more than one hundred species of spiders belonging to several families from Terai Conservation Area, particularly Lakhimpur Kheri district. Lawania and Mathur (2014a, b, c, d) recorded 56 species of spiders from Agra and Mathura districts of Uttar Pradesh. Singh and Singh (2014) reported 58 species belonging to 28 genera and 10 families while Sharma and Singh (2018a, b) accounted 63 species of spiders in five districts of northeast Uttar Pradesh (Kushinagar, Deoria, Gorakhpur, Maharajganj, Siddharthnagar). Later on, Kumar et al. (2017a, b) catalogued 44 species from Lucknow and IAEES www.iaees.org 22 Arthropods, 2022, 11(1): 18-55 Unnao districts, and Mishra and Rastogi (2020) recorded 14 species of spiders from Varanasi district. Recently, Singh et al. (2021) recorded 61 species of spiders belonging to 12 families in Parvati Aranga Bird Sanctuary, Gonda. Most of the national parks and sanctuaries, forest areas, agricultural fields of the states still await intensive and extensive survey programmes to record these predatory chelicerates. In the present compilation, a total of 286 species described under 147 genera belonging to 36 families were enlisted that have been recorded/described from only 36 districts of Uttar Pradesh giving up-to-date information in the light of modern taxonomic concepts. Out of 75 districts of Uttar Pradesh, maximum number of species of spiders were recorded from Lakhimpur Kheri (178 species) followed by Agra (93 species), Mathura (63 species), Lucknow and Gonda (61 species each), Maharajganj (61 species), Siddharthnagar (61 species), Gorakhpur and Kushinagar (57 species each), Unnao (49 species), and Deoria (42 species) (Table 1). No faunal survey of spiders so far conducted in 41 districts of Uttar Pradesh (Table 1; Fig. 1). Hence, an intensive and extensive faunal survey is required in these areas. Following is the familywise list of species of spiders recorded/described from different districts of Uttar Pradesh. 3.1.1 Family Agelenidae • Agelena gautami Tikader, 1962 (Uniyal and Hore, 2009) • Agelena inda Simon, 1897 (Uniyal and Hore, 2009) • Agelena sp. (Hore and Uniyal, 2008a, b; Lawania and Mathur, 2014a, b, c) • Tegenaria domestica (Clerck, 1757) (Lawania and Mathur, 2014a) • Tegenaria sp. (Anjali and Prakash, 2012) 3.1.2 Family Amaurobiidae • Amaurobius jugorum L. Koch, 1868 (Marusik et al., 2012) 3.1.3 Family Araneidae • Anepsion maritatum (Pickard-Cambridge, 1877) (Lawania and Mathur, 2014a) • Arachnura melanura Simon, 1867 (Hore and Uniyal, 2008a, b; Uniyal and Hore, 2009) • Araneus bilunifer Pocock, 1900 (Hore and Uniyal, 2008a, b; Uniyal and Hore, 2009) • Araneus diadematus Clerck, 1757 (Yadav and Prakash, 2021) • Araneus ellipticus (Tikader and Bal, 1981) (Singh and Singh, 2014; Sharma and Singh, 2018a, b; Singh et al., 2021) • Araneus mitificus (Simon, 1886) (Lawania and Mathur, 2014a, c; Singh and Singh, 2014; Kumar et al., 2017a, b; Sharma and Singh, 2018a, b; Singh et al., 2021) • Araneus sp. (Hore and Uniyal, 2008b; Uniyal and Hore, 2009; Anjali and Prakash, 2012; Singh et al., 2013; Chandra et al., 2017; Sharma and Singh, 2018a; Sujayanand et al., 2021) • Argiope aemula (Walckenaer, 1837) (Lawania and Mathur, 2014a, b, c, d; Singh and Singh, 2014; Chaubey, 2017a; Sharma and Singh, 2018a, b; Chandra et al., 2021; Singh et al., 2021) • Argiope anasuja Thorell, 1887 (Hore and Uniyal, 2008a, b; Uniyal and Hore, 2009; Anjali and Prakash, 2012; Lawania and Mathur, 2014a, b, c, d; Kumar et al., 2017a, b; Sharma and Singh, 2018a, b; Singh et al., 2021) • Argiope aurantia Lucas, 1833 (Anjali and Prakash, 2012) • Argiope catenulata (Doleschall 1859) (Singh and Singh, 2014; Sharma and Singh, 2018a, b; Singh et al., 2021) • Argiope luzona (Walckenaer, 1837) (Sharma and Singh, 2018a, b; Singh et al., 2021) • Argiope pulchella Thorell, 1881 (Biswas and Biswas, 2006; Hore and Uniyal, 2008a, b, c; Uniyal and Hore, IAEES www.iaees.org Arthropods, 2022, 11(1): 18-55 23 2009; Lawania and Mathur, 2014a, b, c, d; Kumar et al., 2017a, b; Sharma and Singh, 2018a, b; Anjali et al., 2019; Singh et al., 2021) • Chorizopes anjanes Tikader, 1965 (Chandra et al., 2021) • Cyclosa bifida (Doleschall, 1859) (Singh and Singh, 2014; Sharma and Singh, 2018a, b; Singh et al., 2021) • Cyclosa confraga (Thorell,1892) (Hore and Uniyal, 2008a, b, c; Kumar et al., 2017a, b; Uniyal and Hore, 2009; Kumar et al., 2017a) • Cyclosa hexatuberculata Tikader, 1982 (Chandra et al., 2021) • Cyclosa insulana (Costa, 1834) (Lawania and Mathur, 2014a; Kumar et al., 2017a, b; Sharma and Singh, 2018a, b; Singh et al., 2021) • Cyclosa mulmeinensis (Thorell, 1887) (Hore and Uniyal, 2008a, b; Uniyal and Hore, 2009; Singh and Singh, 2014; Sharma and Singh, 2018a, b; Singh et al., 2021) • Cyclosa simoni Tikader, 1982 (Hore and Uniyal, 2008a, b; Uniyal and Hore, 2009) • Cyclosa sp. (Hore and Uniyal, 2008a, b, c; Uniyal and Hore, 2009; Lawania and Mathur, 2014a, b, c, d) • Cyphalonotus sp. (Hore and Uniyal, 2008a, c) • Cyrtarachne sp. (Kumar et al., 2017a, b) • Cyrtophora bidenta Tikader, 1970 (Kumar et al., 2017a, b; Uniyal and Hore, 2009; Kumar et al., 2017a) • Cyrtophora cicatrosa (Stoliczka, 1869) (Pocock, 1900; Tikader, 1982; Biswas and Biswas, 1992; Uniyal and Hore, 2009; Anjali and Prakash, 2012; Lawania and Mathur, 2014a, b, c, d; Chaubey and Mishra, 2016) • Cyrtophora citricola (Forsskål, 1775) (Hore and Uniyal, 2008a, b; Uniyal and Hore, 2009; Anjali and Prakash, 2012; Lawania and Mathur, 2014a, b, c, d; Singh and Singh, 2014; Chaubey and Mishra, 2017a; Sharma and Singh, 2018a, b; Mishra and Rastogi, 2020; Singh et al., 2021) • Cyrtophora exanthematica (Doleschall, 1859) (Sharma and Singh, 2018a, b; Singh et al., 2021) • Cyrtophora feae Thorell, 1887 (Lawania and Mathur, 2014a, b) • Cyrtophora jabalpurensis Gajbe and Gajbe, 1999 (Hore and Uniyal, 2008a, b) • Cyrtophora ksudra Sherriffs, 1928 (Hore and Uniyal, 2008a, b; Uniyal and Hore, 2009) • Cyrtophora moluccensis (Doleschall, 1857) (Uniyal and Hore, 2009; Lawania and Mathur, 2014a, b) • Cyrtophora unicolor (Doleschall, 1857) (Hore and Uniyal, 2008c) • Cyrtophora sp. (Hore and Uniyal, 2008a, b, c; Uniyal and Hore, 2009) • Eriophora sp. (Kumar et al., 2017a, b) • Eriovixia excelsa (Simon, 1889) (Hore and Uniyal, 2008a, b; Singh and Singh, 2014; Sharma and Singh, 2018a, b; Singh et al., 2021) • Eriovixia laglaizei (Simon, 1877) (Hore and Uniyal, 2008a, b; Singh and Singh, 2014; Sharma and Singh, 2018a, b; Singh et al., 2021) • Gasteracantha dalyi Pocock, 1900 (Hore and Uniyal, 2008a, b; Uniyal and Hore, 2009) • Gasteracantha diadesmia Thorell, 1887 (Singh and Singh, 2014; Sharma and Singh, 2018a, b) • Gasteracantha geminata (Fabricius, 1798) (Hore and Uniyal, 2008a, b; Uniyal and Hore, 2009) • Gasteracantha kuhli C.L. Koch, 1837 (Singh and Singh, 2014; Sharma and Singh, 2018a, b; Singh et al., 2021) • Gasteracantha sp. (Hore and Uniyal, 2008a, b, c) • Gea subarmata Thorell, 1890 (Hore and Uniyal, 2008a, b; Uniyal and Hore, 2009; Singh and Singh, 2014; Sharma and Singh, 2018a, b; Singh et al., 2021) • Gea sp. (Hore and Uniyal, 2008a, b, c) • Larinia chloris Audoin (1826) (Uniyal and Hore, 2009) • Larinia emertoni Gajbe and Gajbe, 2004 (Lawania and Mathur, 2014a; Sharma and Singh, 2018b; Singh et IAEES www.iaees.org 24 Arthropods, 2022, 11(1): 18-55 al., 2021) • Larinia kanpurae Patel and Nigam, 1994 (Patel and Nigam, 1994; Sharma and Singh, 2018a) • Larinia phthisica (L. Koch, 1871) (Singh and Singh, 2014; Sharma and Singh, 2018a, b; Chandra et al., 2021; Singh et al., 2021) • Larinia sp. (Hore and Uniyal, 2008a, b; Uniyal and Hore, 2009) • Macracantha hasselti (C. L. Koch, 1837) (Hore and Uniyal, 2008a, b; Uniyal and Hore, 2009; Sharma and Singh, 2018a, b; Singh et al., 2021) • Neoscona adianta (Walckenaer, 1802) (Mishra et al., 2012a) • Neoscona biswasi Bhandari and Gajbe, 2001 (Hore and Uniyal, 2008a, b; Uniyal and Hore, 2009) • Neoscona crucifera (Lucas, 1838) (Mishra et al., 2012a) • Neoscona dhruvai Patel and Nigam, 1994 (Patel and Nigam, 1994; Sharma and Singh, 2018a, b; Singh et al., 2021) • Neoscona inusta (L. Koch, 1871) (Khan and Misra, 2003; Chandra et al., 2017) • Neoscona molemensis Tikader and Bal, 1981 (Singh and Singh, 2014; Sharma and Singh, 2018a, b; Singh et al., 2021) • Neoscona mukerjei Tikader, 1980 (Hore and Uniyal, 2008a, b; Uniyal and Hore, 2009; Kumar et al., 2017a, b; Mishra and Rastogi, 2020; Chandra et al., 2021) • Neoscona nautica (L. Koch, 1875) (Mishra et al., 2012b; Singh and Singh, 2014; Sharma and Singh, 2018a, b; Singh et al., 2021; Yadav and Prakash, 2021) • Neoscona odites (Simon, 1906) (Hore and Uniyal, 2008a, b; Uniyal and Hore, 2009; Halder et al., 2012) • Neoscona sinhagadensis (Tikader, 1975) (Tandon and Lal, 1983) • Neoscona theisi (Walckenaer, 1837) (Hore and Uniyal, 2008a, b; Uniyal and Hore, 2009; Singh and Singh, 2014; Chandra et al., 2017; Kumar et al., 2017a, b; Sharma and Singh, 2018a, b; Mishra and Rastogi, 2020; Mishra et al., 2021; Singh et al., 2021; Yadav and Prakash, 2021) • Neoscona vigilans (Blackwall, 1865) (Hore and Uniyal, 2008a, b, c; Uniyal and Hore, 2009) • Neoscona sp. (Lawania and Mathur, 2014a) • Nephila kuhlii (Doleschall, 1859) (Lawania and Mathur, 2014a, b) • Nephila pilipes (Fabricius, 1793) (Tikader, 1982; Hore and Uniyal, 2008a, b, c; Uniyal and Hore, 2009; Lawania and Mathur, 2014a, b; Sharma and Singh, 2018a, b; Singh et al., 2021) • Nephilengys malabarensis (Walckenaer, 1841) (Singh and Singh, 2014; Sharma and Singh, 2018a, b; Singh et al., 2021) • Parawixia dehaani (Doleschall, 1859) (Hore and Uniyal, 2008a, b; Uniyal and Hore, 2009; Singh and Singh, 2014; Kumar et al., 2017a, b; Sharma and Singh, 2018a, b) • Parawixia sp. (Hore and Uniyal, 2008a, b; Uniyal and Hore, 2009) • Poltys illepidus C.L. Koch, 1843 (Uniyal and Hore, 2009; Singh and Singh, 2014; Sharma and Singh, 2018a, b) • Poltys sp. (Hore and Uniyal, 2008a, b) • Trichonephila clavata (L. Koch, 1878) (Kumar et al., 2017a, b; Yadav, 2018) • Zygiella sp. (Hore and Uniyal, 2008a, b; Uniyal and Hore, 2009) 3.1.4 Family Barychelidae • Sason robustum (Pickard-Cambridge, 1883) (Hore and Uniyal, 2008a, b; Uniyal and Hore, 2009) • Sasonichus sullivani Pocock, 1900 (Hore and Uniyal 2008a, b; Uniyal and Hore, 2009) 3.1.5 Family Cheiracanthiidae IAEES www.iaees.org Arthropods, 2022, 11(1): 18-55 25 • Cheiracanthium adjacens O. Pickard-Cambridge, 1885 (Hore and Uniyal, 2008a, b; Uniyal and Hore, 2009) • Cheiracanthium danieli Tikader, 1975 (Tandon and Lal, 1983) • Cheiracanthium sp. (Lawania and Mathur, 2014a) 3.1.6 Family Clubionidae • Clubiona boxaensis Biswas and Biswas, 1992 (Hore and Uniyal, 2008a, b, c) • Clubiona deletrix Pickard-Cambridge, 1885 (Hore and Uniyal, 2008a, b) • Clubiona drassodes Pickard-Cambridge, 1874 (Singh and Singh, 2014; Sharma and Singh, 2018a, b; Singh et al., 2021) • Clubiona filicata O. P.-Cambridge, 1874 (Hore and Uniyal, 2008a, b, c) • Clubiona japonicola Bösenberg and Strand, 1906 (Khan and Misra, 2003; Singh and Singh, 2014; Chandra et al., 2017; Sharma and Singh, 2018a, b; Singh et al., 2021) • Clubiona sp. (Hore and Uniyal, 2008a, b; Sharma and Singh, 2018a; Sujayanand et al., 2021) 3.1.7 Family Corinnidae • Castianeira zetes Simon, 1897 (Yadav and Prakash, 2021) • Castianeira sp. (Lawania and Mathur, 2014a) 3.1.8 Family Ctenidae • Nilus sp. (Kumar et al., 2017a, b) 3.1.9 Family Desidae • Desis inermis Gravely, 1927 (Hore and Uniyal, 2008a, b) • Desis sp. (Hore and Uniyal, 2008a, b) 3.1.10 Family Dictynidae • Dictyna turbida Simon, 1905 (Hore and Uniyal, 2008a) • Nigma albida (O. Pickard-Cambridge, 1885) (Hore and Uniyal, 2008a) • Nigma shiprai (Tikader, 1966) (Lawania and Mathur, 2014a) 3.1.11 Family Eresidae • Stegodyphus sarasinorum Karsch, 1892 (Tandon and Lal, 1983; Kumar et al., 2017a, b; Mishra and Rastogi, 2020; Chandra et al., 2021) 3.1.12 Family Gnaphosidae • Callilepis lambai Tikader and Gajbe, 1977 (Lawania and Mathur, 2014a, b, c, d) • Callilepis rukminiae Tikader and Gajbe, 1977 (Lawania and Mathur, 2014a, b, c, d) • Drassodes delicatus (Blackwall, 1867) (Blackwall, 1867) • Drassodes gangeticus Tikader and Gajbe, 1975 (Hore and Uniyal, 2008a, b, c; Uniyal and Hore, 2009) • Drassodes himalayensis Tikader and Gajbe, 1975 (Gajbe, 1988) • Drassodes luridus (Pickard-Cambridge, 1874) (Hore and Uniyal, 2008a, b; Uniyal and Hore, 2009) • Drassodes parvidens Caporiacco, 1935 (Hore and Uniyal, 2008a, b; Uniyal and Hore, 2009) • Drassodes sp (Anjali and Prakash, 2012; Lawania and Mathur, 2014a, b, c, d) • Eilica kandarpae Nigam and Patel, 1996 (Nigam and Patel, 1996) • Gnaphosa kailana Tikader, 1966 (Hore and Uniyal, 2008a, b; Uniyal and Hore, 2009) • Gnaphosa poonaensis Tikader, 1973 (Tikader, 1982; Gajbe, 1988) IAEES www.iaees.org 26 Arthropods, 2022, 11(1): 18-55 • Gnaphosa stoliczkai Pickard-Cambridge, 1885 (Hore and Uniyal, 2008a, b, c; Uniyal and Hore, 2009) • Gnaphosa sp. (Hore and Uniyal, 2008a, b) • Haplodrassus ambalaensis Gajbe, 1992 (Hore and Uniyal, 2008b; Uniyal and Hore, 2009) • Haplodrassus bengalensis Gajbe, 1992 (Hore and Uniyal, 2008a) • Haplodrassus morosus (Pickard-Cambridge, 1872) (Uniyal and Hore, 2009) • Haplodrassus tehriensis Tikader and Gajbe, 1977 (Uniyal and Hore, 2009) • Haplodrassus sp. (Hore and Uniyal, 2008a, b; Yadav & Prakash, 2021) • Herpyllus sp. (Hore and Uniyal, 2008a, b) • Ladissa sp. (Hore and Uniyal, 2008b) • Marinarozelotes jaxartensis (Kroneberg, 1875) (Tikader and Gajbe, 1975; Tikader, 1982; Gajbe, 1988) • Prodidomus saharanpurensis (Tikader, 1982) (Tikader, 1982) • Scotophaeus sp. (Hore and Uniyal, 2008a, b) • Setaphis browni (Tucker, 1923) (Gajbe, 1988) • Urozelotes rusticus (L. Koch, 1872) (Singh and Singh, 2014; Sharma and Singh, 2018b; Singh et al., 2021) • Zelotes calcuttaensis (Biswas, 1984) (Uniyal and Hore, 2009) • Zelotes chandosiensis Tikader and Gajbe, 1976 (Tikader and Gajbe, 1976a; Tikader, 1982) • Zelotes nainitalensis Tikader and Gajbe, 1976 (Tikader and Gajbe, 1976a; Uniyal and Hore, 2009) • Zelotes pexus (Simon, 1885) (Uniyal and Hore, 2009) • Zelotes sataraensis Tikader and Gajbe, 1979 (Gajbe, 1988) • Zelotes sp. (Hore and Uniyal, 2008a, b, c) 3.1.13 Family Hahniidae • Hahnia mridulae Tikader, 1970) (Hore and Uniyal, 2008a) • Hahnia sp. (Hore and Uniyal, 2008a) • Neoantistea maxima (Caporiacco, 1935) (Hore and Uniyal, 2008a) 3.1.14 Family Hersiliidae • Hersilia savignyi Lucas, 1836 (Pocock, 1900; Biswas and Biswas, 1992; Uniyal and Hore, 2009; Lawania and Mathur, 2014a, b; Sharma and Singh, 2018a, b; Anjali and Prakash, 2019; Singh et al., 2021) • Hersilia sp. (Anjali and Prakash, 2012; Kumar et al., 2017a, b) • Neotama punctigera Baehr and Baehr, 1993 (Uniyal and Hore, 2009) 3.1.15 Family Ischnothelidae • Indothele mala Coyle, 1995 (Hore and Uniyal 2008a) • Indothele rothi Coyle, 1995 (Hore and Uniyal, 2008a, b) 3.1.16 Family Linyphiidae • Atypena sp. (Chandra et al., 2017; Kumar et al., 2017a, b) • Erigone rohtangensis Tikader, 1981 (Hore and Uniyal, 2008a) • Erigone sp. (Kumar et al., 2017a, b) • Lepthyphantes peramplus (Pickard–Cambridge, 1885) (Uniyal and Hore, 2009) • Lepthyphantes stramineus (Pickard–Cambridge, 1885) (Uniyal and Hore, 2009) • Linyphia sikkimensis Tikader, 1970 (Uniyal and Hore, 2009) • Linyphia sp. (Tandon and Lal, 1983; Hore and Uniyal, 2008a; Lawania and Mathur, 2014a, b, c, d; Kumar et al., 2017a, b) IAEES www.iaees.org Arthropods, 2022, 11(1): 18-55 27 • Oedothorax globiceps Thaler, 1987 (Hore and Uniyal, 2008a; Uniyal and Hore, 2009) • Oedothorax sp. (Hore and Uniyal, 2008a; Uniyal and Hore, 2009) 3.1.17 Family Liocranidae • Oedignatha indica (Tikader, 1981) (Hore and Uniyal, 2008a) • Oedignatha sp. (Hore and Uniyal, 2008a) 3.1.18 Family Lycosidae • Arctosa himalayensis Tikader and Malhotra, 1980 (Biswas and Biswas, 1992; Khan and Misra, 2003) • Arctosa indica Tikader and Malhotra, 1980 (Hore and Uniyal, 2008a, b, c) • Arctosa sp. (Hore and Uniyal, 2008a, b, c; Uniyal and Hore, 2009) • Evippa solanensis Tikader and Malhotra, 1980 (Uniyal and Hore, 2009) • Hippasa agelenoides (Simon, 1884) (Kumar et al., 2017a, b; Yadav & Prakash, 2021) • Hippasa greenalliae (Blackwall, 1867) (Blackwall, 1867; Mishra and Rastogi, 2020) • Hippasa himalayensis Gravely, 1924 (Uniyal and Hore, 2009) • Hippasa holmerae Thorell, 1895 (Biswas and Biswas, 1992; Yadav et al., 2012a; Chandra et al., 2017; Sharma and Singh, 2018a, b; Singh et al., 2021) • Hippasa olivacea (Thorell, 1887) (Chandra et al., 2021) • Hippasa partita (O. Pickard-Cambridge, 1876) (Singh and Singh, 2014; Sharma and Singh, 2018a, b; Singh et al., 2021) • Hippasa pisaurina Pocock, 1900 (Hore and Uniyal, 2008a, b, c) • Hippasa sp. (Hore and Uniyal, 2008a, b; Lawania and Mathur, 2014a) • Lycosa carmichaeli Gravely, 1924 (Gravely, 1924) • Lycosa mackenziei Gravely, 1924 (Singh and Singh, 2014; Lawania and Mathur, 2014a, b, c; Sharma and Singh, 2018a, b; Singh et al., 2021; Yadav & Prakash, 2021) • Lycosa nigrotibialis Simon, 1884 (Chandra et al., 2021) • Lycosa pictula Pocock, 1901 (Anjali and Prakash, 2012; Lawania and Mathur, 2014a, b, c, d) • Lycosa prolifica Pocock, 1901 (Biswas and Biswas, 2006) • Lycosa tista Tikader, 1970 (Uniyal and Hore, 2009; Anjali and Prakash, 2012; Kumar et al., 2017a, b; Yadav & Prakash, 2021) • Lycosa sp. (Agrawal et al., 2010) • Pardosa heterophthalma (Simon, 1898) (Biswas and Biswas, 2010) • Pardosa kupupa (Tikader, 1970) (Hore and Uniyal, 2008a, b) • Pardosa minuta Tikader and Malhotra, 1976 (Hore and Uniyal, 2008a, b) • Pardosa pseudoannulata (Bösenberg and Strand, 1906) (Khan and Misra, 2003; Anjali and Prakash, 2012; Lawania and Mathur, 2014a; Singh and Singh, 2014; Chandra et al., 2017; Sharma and Singh, 2018a, b; Singh et al., 2021) • Pardosa songosa Tikader and Malhotra, 1976 (Tikader and Malhotra, 1976) • Pardosa sumatrana (Thorell, 1890) (Singh and Singh, 2014; Kumar et al., 2017a, b; Sharma and Singh, 2018a, b; Mishra and Rastogi, 2020; Mishra et al., 2021; Singh et al., 2021) • Pardosa timidula (Roewer, 1951) (Uniyal and Hore, 2009) • Pardosa sp. (Hore and Uniyal, 2008a, b; Lawania and Mathur, 2014a, b, c; Sharma and Singh, 2018a; Agrawal et al., 2010) • Trochosa himalayensis Tikader and Malhotra, 1980 (Hore and Uniyal, 2008a, b) • Trochosa punctipes (Gravely, 1924) (Gravely, 1924; Tikader, 1966a) IAEES www.iaees.org 28 Arthropods, 2022, 11(1): 18-55 • Trochosa sp. (Hore and Uniyal, 2008a, b; Kumar et al., 2017a, b) • Trochosa urbana O. Pickard-Cambridge, 1876 (Lawania and Mathur, 2014a, d; Kumar et al., 2017b) • Wadicosa fidelis (O. Pickard-Cambridge, 1872) (Biswas and Biswas, 1992; Khan and Misra, 2003; Hore and Uniyal, 2008a, b, c; Anjali and Prakash, 2012; Lawania and Mathur, 2014a, b, c; Singh and Singh, 2014; Chandra et al., 2017; Sharma and Singh, 2018a, b; Singh et al., 2021) 3.1.19 Family Oonopidae • Gamasomorpha clypeolaria Simon, 1907 (Uniyal and Hore, 2009) • Gamasomorpha sp. (Uniyal and Hore, 2009) • Triaeris nagarensis Tikader and Malhotra, 1974 (Chandra et al., 2021) 3.1.20 Family Oxyopidae • Hamataliwa sp. (Kumar et al., 2017a, b) • Oxyopes assamensis Tikader, 1969 (Lawania and Mathur, 2014a, b) • Oxyopes birmanicus Thorell, 1887 (Hore and Uniyal, 2008a, b; Anjali and Prakash, 2012; Kumar et al., 2017a, b; Lawania and Mathur, 2014a, b, c, d ) • Oxyopes elongatus Biswas et al., 1996 (Hore and Uniyal, 2008a, b; Uniyal and Hore, 2009; Mishra and Rastogi, 2020) • Oxyopes hindostanicus Pocock, 1901 (Sherriffs, 1951) • Oxyopes indicus (Walckenaer, 1805) (Anjali and Prakash, 2012) • Oxyopes javanus Thorell, 1887 (Khan and Misra, 2003; Lawania and Mathur, 2014a, b, c, d; Singh and Singh, 2014; Kumar et al., 2017a, b; Sharma and Singh, 2018a, b; Mishra and Rastogi, 2020; Mishra et al., 2021; Singh et al., 2021) • Oxyopes ketani Gajbe and Gajbe, 1999 (Gajbe, 1992a) • Oxyopes lineatipes (C.L.Koch, 1847) (Halder et al., 2012) • Oxyopes pandae Tikader, 1969 (Tikader, 1969a; Gajbe, 1999; Khan and Misra, 2003) • Oxyopes pankaji Gajbe and Gajbe, 2000 (Lawania and Mathur, 2014a, b, c, d) • Oxyopes pawani Gajbe, 1992 (Gajbe, 1992a) • Oxyopes quadrifasciatus L. Koch, 1878 (Chaubey, 2019a) • Oxyopes ratnae Tikader, 1970 (Khan and Misra, 2003; Anjali and Prakash, 2012; Lawania and Mathur, 2014a, b, c, d) • Oxyopes ryvesi Pocock, 1901 (Pocock, 1901; Sherriffs, 1951; Gajbe, 2008) • Oxyopes salticus Hentz, 1845 (Anjali and Prakash, 2012) • Oxyopes sertatus L. Koch, 1878 (Anjali and Prakash, 2012; Lawania and Mathur, 2014a) • Oxyopes shweta Tikader 1970 (Hore and Uniyal, 2008a, b; Uniyal and Hore, 2009; Anjali and Prakash, 2012; Lawania and Mathur, 2014a, b, c, d; Sharma and Singh, 2018a, b; Singh et al., 2021) • Oxyopes sp. (Hore and Uniyal, 2008a, b; Lawania and Mathur, 2014a, b, c, Dubey et al., 2021; Sujayanand et al., 2021) • Peucetia ketani Gajbe, 1992 (Gajbe, 1992a) • Peucetia yogeshi Gajbe, 1999 (Chandra et al., 2021) • Peucetia sp. (Hore and Uniyal, 2008a, b; Kumar et al., 2017a, b) 3.1.21 Family Philodromidae • Philodromus bhagirathai Tikader, 1966 (Tikader, 1966b, 1971) • Philodromus pali Gajbe and Gajbe, 2000 (Uniyal and Hore, 2009) IAEES www.iaees.org Arthropods, 2022, 11(1): 18-55 29 • Philodromus sp. (Hore and Uniyal, 2008a, b; Anjali and Prakash, 2012; Lawania and Mathur, 2014a; Kumar et al., 2017a, b) 3.1.22 Family Pholcidae • Artema atlanta Walckenaer, 1837 (Blackwall, 1867; Pocock, 1900; Lawania and Mathur, 2014a, b, c, d; Anjali and Prakash, 2019) • Artema sp. (Hore and Uniyal, 2008a; Kumar et al., 2017a) • Crossopriza lyoni (Blackwall, 1867) (Blackwall, 1867; Pocock, 1900; Hore and Uniyal, 2008a; Lawania and Mathur, 2014a, b, c, d; Uniyal and Hore, 2009; Kumar et al., 2017a, b; Sharma and Singh, 2018a, b; Anjali and Prakash, 2019; Singh et al., 2021) • Pholcus phalangioides (Fuesslin, 1775) (Lawania and Mathur, 2014a, b, d; Singh and Singh, 2014; Kumar et al., 2017a, b; Sharma and Singh, 2018a, b; Singh et al., 2021) • Pholcus sp. (Lawania and Mathur, 2014a, b) • Smeringopus pallidus (Blackwall, 1858) (Uniyal and Hore, 2009) 3.1.23 Family Pisauridae • Nilus albocinctus (Doleschall, 1859) (Uniyal and Hore, 2009) • Nilus decorata (Patel and Reddy, 1990) (Hore and Uniyal, 2008a; Uniyal and Hore, 2009) • Pisaura sp. (Hore and Uniyal, 2008a, b; Uniyal and Hore, 2009; Kumar et al., 2017a, b) 3.1.24 Family Salticidae • Afrafacila sp. (Yadav and Prakash, 2021) • Asemonea tenuipes (Pickard-Cambridge, 1869) (Lawania and Mathur, 2014a) • Bavia sp. (Lawania and Mathur, 2014a) • Carrhotus viduus (C.L. Koch, 1846) (Uniyal and Hore, 2009) • Carrhotus sp. (Kumar et al., 2017a, b) • Cosmophasis umbratica Simon, 1903 (Lawania and Mathur, 2014a) • Epocilla aurantiaca (Simon, 1885) (Anjali et al., 2019) • Epocilla sp. (Yadav and Prakash, 2021) • Harmochirus brachiatus (Thorell, 1877) (Singh and Singh, 2014; Sharma and Singh, 2018a, b; Singh et al., 2021) • Hasarius adansoni (Audouin, 1826) (Anjali and Prakash, 2012; Lawania and Mathur, 2014a; Singh and Singh, 2014; Sharma and Singh, 2018a, b; Singh et al., 2021) • Hyllus semicupreus (Simon, 1885) (Lawania and Mathur, 2014a, b, c, d; Yadav and Prakash, 2021) • Hyllus sp. (Kumar et al., 2017b) • Marengo crassipes Peckham and Peckham, 1892 (Sharma and Singh, 2018a, b; Singh et al., 2021) • Marpissa sp. (Tandon and Lal, 1983; Hore and Uniyal, 2008a; Halder et al., 2012) • Menemerus bivittatus (Dufour, 1831) (Chaubey et al., 2019a; Yadav and Prakash, 2021) • Menemerus semilimbatus (Hahn, 1829) (Chaubey and Yadav, 2017a; Anjali and Prakash, 2019; Anjali et al., 2019; Yadav and Prakash, 2021) • Myrmaplata plataleoides (O. Pickard-Cambridge, 1869) (Mishra and Rastogi, 2020) • Myrmarachne himalayensis Narayan, 1915 (Uniyal and Hore, 2009) • Myrmarachne melanocephala MacLeay, 1839 (Lawania and Mathur, 2014a; Kumar et al., 2017a, b; Sharma and Singh, 2018a, b; Anjali et al., 2019; Chaubey, 2019b; Singh et al., 2021) • Myrmarachne sp. (Hore and Uniyal, 2008a, b, c; Uniyal and Hore, 2009; Sharma and Singh, 2018a) IAEES www.iaees.org 30 Arthropods, 2022, 11(1): 18-55 • Opisthoncus sp. (Chaubey, 2019c) • Phlegra sp. (Kumar et al., 2017a, b) • Phidippus audax (Hentz, 1845) (Chaubey, 2017b) • Phidippus clarus Keyserling, 1885 (Anjali and Prakash, 2012) • Phidippus sp. (Tandon and Lal, 1983; Chandra et al., 2017) • Phidippus yashodharae Tikader, 1977 (Anjali and Prakash, 2012; Lawania and Mathur, 2014a, b, c) • Phintella bifurcata Proszyński, 1992 (Uniyal and Hore, 2009) • Phintella sp. (Hore and Uniyal, 2008a, b; Kumar et al., 2017a, b) • Phintella vittata (C L Koch, 1846) (Lawania and Mathur, 2014a, b, c; Kumar et al., 2017a; Mishra and Rastogi, 2020) • Plexippus calcutaensis (Tikader, 1974) (Singh and Singh, 2014; Sharma and Singh, 2018a, b; Singh et al., 2021) • Plexippus paykulli (Audouin, 1826) (Hore and Uniyal, 2008a, b; Uniyal and Hore, 2009; Anjali and Prakash, 2012; Lawania and Mathur, 2014a, b, c, d; Singh and Singh, 2014; Chaubey and Yadav, 2017b; Kumar et al., 2017a, b; Sharma and Singh, 2018a, b; Mishra and Rastogi, 2020; Singh et al., 2021) • Plexippus petersi (Karsch, 1878) (Singh and Singh, 2014; Chaubey, 2017c; Sharma and Singh, 2018a, b; Anjali et al., 2019; Singh et al., 2021) • Plexippus redimitus Simon, 1906 (Uniyal and Hore, 2009) • Plexippus sp. (Hore and Uniyal, 2008a, b; Kumar et al., 2017a) • Portia albimana (Simon, 1900) (Uniyal and Hore, 2009) • Portia assamensis Wanless, 1978 (Lawania and Mathur, 2014a, b, c, d) • Portia sp. (Anjali and Prakash, 2012) • Rhene flavigera (C.L. Koch, 1846) (Kumar et al., 2017a, b) • Rhene indica Tikader, 1973 (Uniyal and Hore, 2009) • Rhene sp. (Hore and Uniyal, 2008a, b; Uniyal and Hore, 2009; Kumar et al., 2017a) • Salticus sp. (Khan and Misra, 2003; Kumar et al., 2017a, b) • Siler sp. (Kumar et al., 2017a, b) • Stenaelurillus lesserti Reimoser, 1934 (Sharma and Singh, 2018a, b; Singh et al., 2021) • Telamonia dimidiata (Simon, 1899) (Khan and Misra, 2003; Anjali and Prakash, 2012; Lawania and Mathur, 2014a, b, c, d; Chaubey, 2019d; Mishra and Rastogi, 2020; Anjali et al., 2019) • Telamonia festiva Thorell, 1887 (Uniyal and Hore, 2009) • Telamonia sp. (Hore and Uniyal, 2008a, b) • Thiania sp. (Kumar et al., 2017a, b) • Thyene imperialis (Rossi 1846) (Yadav and Prakash, 2021) • Zenodorus sp. (Chaubey et al., 2019b) 3.1.25 Family Scytodidae • Scytodes pallida Doleschall, 1859 (Uniyal and Hore, 2009) 3.1.26 Family Selenopidae • Selenops radiatus Latreille, 1819 (Pocock, 1900; Kumar et al., 2017a) • Selenops sp. (Anjali and Prakash, 2012; Lawania and Mathur, 2014a) 3.1.27 Family Sparassidae • Heteropoda fabrei Simon, 1885 (Hore and Uniyal, 2008a, b; Uniyal and Hore, 2009) IAEES www.iaees.org Arthropods, 2022, 11(1): 18-55 31 • Heteropoda leprosa Simon, 1884 (Biswas and Biswas, 1992) • Heteropoda nilgirina Pocock, 1901 (Sethi and Tikader, 1988) • Heteropoda sp. (Kumar et al., 2017a) • Heteropoda venatoria (Linnaeus 1767) (Hore and Uniyal, 2008a, b; Uniyal and Hore, 2009; Singh and Singh, 2014; Kumar et al., 2017a, b; Sharma and Singh, 2018a, b; Singh et al., 2021) • Olios obesulus (Pocock, 1901) (Gravely, 1931; Sethi and Tikader, 1988) • Olios milleti (Pocock, 1901) (Mishra and Rastogi, 2020) • Olios punctipes Simon, 1884 (Gravely, 1931; Sethi and Tikader, 1988) • Olios tikaderi Kundu et al., 1999 (Hore and Uniyal, 2008a, b; Uniyal and Hore, 2009) • Olios sp. (Yadav and Prakash, 2021) • Palystes flavidus Simon, 1897 (Pocock, 1900; Tikader and Sethi, 1990) • Spariolenus buxa (Saha et al., 1995) (Hore and Uniyal, 2008a, b; Uniyal and Hore, 2009) 3.1.28 Family Stenochilidae • Stenochilus crocatus Simon, 1884 (Chandra et al., 2021) 3.1.29 Family Tetrablemmidae • Tetrablemma deccanense (Tikader, 1976) (Uniyal and Hore, 2009) • Tetrablemma sp. (Hore and Uniyal, 2008a, b, c) 3.1.30 Family Tetragnathidae • Guizygiella indica (Tikader and Bal, 1980) (Hore and Uniyal, 2008b; Anjali et al., 2019; Yadav and Prakash, 2021) • Guizygiella melanocrania (Thorell, 1887) (Chaubey, 2017d; Chandra et al., 2021) • Leucauge celebesiana (Walckenaer, 1841) (Hore and Uniyal, 2008a, b, c; Uniyal and Hore, 2009; Kumar et al., 2017a, b; Singh and Singh, 2014; Sharma and Singh, 2018a, b; Singh et al., 2021) • Leucauge decorata (Blackwall, 1864) (Hore and Uniyal, 2008a, b; Uniyal and Hore, 2009; Anjali and Prakash, 2012; Yadav et al., 2012b; Lawania and Mathur, 2014a, b, c, d; Singh and Singh, 2014; Kumar et al., 2017a, b; Sharma and Singh, 2018a, b; Singh et al., 2021) • Leucauge sp. (Hore and Uniyal, 2008a, b; Uniyal and Hore, 2009) • Meta sp. (Hore and Uniyal, 2008a, b; Uniyal and Hore, 2009) • Opadometa fastigata (Simon, 1877) (Gravely, 1921) • Tetragnatha ceylonica Pickard-Cambridge 1869 (Sharma and Singh, 2018a, b; Singh et al., 2021) • Tetragnatha chamberlini (Gajbe, 2004) (Hore and Uniyal, 2008a, b, c; Uniyal and Hore, 2009; Lawania and Mathur, 2014a; Chaubey and Mishra, 2017b) • Tetragnatha javana (Thorell, 1890) (Khan and Misra, 2003; Singh and Singh, 2014; Chandra et al., 2017; Sharma and Singh, 2018a, b; Mishra et al., 2021; Singh et al., 2021) • Tetragnatha keyserlingi Simon, 1890 (Singh and Singh, 2014; Sharma and Singh, 2018a, b; Singh et al., 2021) • Tetragnatha mandibulata Walckenaer, 1842 (Khan and Misra, 2003; Singh and Singh, 2014; Chandra et al., 2017; Kumar et al., 2017a, b; Sharma and Singh, 2018a, b; Singh et al., 2021) • Tetragnatha sp. (Khan and Misra, 2003) • Tylorida sp. (Hore and Uniyal, 2008a) • Tylorida ventralis (Thorell, 1877) (Hore and Uniyal, 2008a, b; Uniyal and Hore, 2009) 3.1.31 Family Theraphosidae IAEES www.iaees.org 32 Arthropods, 2022, 11(1): 18-55 • Chilobrachys hardwickei (Pocock, 1895) (Pocock, 1900; Siliwal et al., 2011) • Chilobrachys sp. (Hore and Uniyal 2008a, b) • Haplocosmia himalayana (Pocock, 1899) (Uniyal and Hore, 2009) 3.1.32 Family Theridiidae • Achaearanea budana Tikader, 1970 (Hore and Uniyal, 2008a, b, c; Uniyal and Hore, 2009) • Achaearanea sp. (Hore and Uniyal, 2008a, b, c; Uniyal and Hore, 2009; Gupta and Siliwal, 2012) • Achaearanea triangularis (Patel, 2005) (Hore and Uniyal, 2008a, b; Uniyal and Hore, 2009) • Argyrodes cyrtophorae Tikader, 1963 (Hore and Uniyal, 2008a, b; Uniyal and Hore, 2009) • Argyrodes fissifrons Pickard-Cambridge, 1869 (Hore and Uniyal, 2008a, b; Uniyal and Hore, 2009) • Chikunia nigra (O. Pickard-Cambridge, 1880) (Hore and Uniyal, 2008a, b; Uniyal and Hore, 2009; Lawania and Mathur, 2014a) • Chrysso urbasae (Tikader, 1970) (Uniyal and Hore, 2009) • Chrysso sp. (Hore and Uniyal, 2008a, b; Uniyal and Hore, 2009) • Dipoenura fimbriata Simon, 1909 (Sharma and Singh, 2018b; Singh et al., 2021) • Meotipa picturata Simon, 1895 (Hore and Uniyal, 2008a, b, c; Uniyal and Hore, 2009) • Meotipa pulcherrima (Mello-Leitão, 1917) (Lawania and Mathur, 2014a) • Molione triacantha Thorell, 1892 (Sharma and Singh, 2018a, b; Singh et al., 2021) • Nihonhimea mundula (L. Koch, 1872) (Lawania and Mathur, 2014a; Anjali and Prakash, 2012) • Steatoda sp. (Lawania and Mathur, 2014a) • Theridion incertum Pickard-Cambridge, 1885 (Uniyal and Hore, 2009) • Theridion manjithar Tikader, 1970 (Uniyal and Hore, 2009) • Theridion sp. (Hore and Uniyal, 2008a, b; Uniyal and Hore, 2009; Chandra et al., 2021) • Thwaitesia margaritifera Pickard-Cambridge, 1881 (Sharma and Singh, 2018a, b; Singh et al., 2021) 3.1.33 Family Thomisidae • Camaricus formosus Thorell, 1887 (Singh and Singh, 2014; Sharma and Singh, 2018a, b; Singh et al., 2021) • Camaricus sp. (Singh and Singh, 2014; Kumar et al., 2017a, b; Sharma and Singh, 2018a, b) • Diaea subdola Pickard-Cambridge, 1885 (Hore and Uniyal, 2008a, b; Uniyal and Hore, 2009) • Indoxysticus minutus (Tikader, 1960) (Lawania and Mathur, 2014a; Kumar et al., 2017a, b) • Lysiteles sp. (Kumar et al., 2017a) • Mastira menoka (Tikader, 1963) (Sharma and Singh, 2018a, b; Singh et al., 2021) • Misumena indra Tikader, 1963 (Hore and Uniyal, 2008a, b; Uniyal and Hore, 2009) • Misumena mridulai Tikader, 1962 (Hore and Uniyal, 2008a, b; Uniyal and Hore, 2009) • Misumena vatia (Clerck, 1757) (Kumar et al., 2017a, b) • Misumena sp. (Yadav and Prakash, 2021) • Ozyptila chandosiensis Tikader, 1980 (Tikader, 1980) • Ozyptila manii Tikader, 1961 (Hore and Uniyal, 2008a, b; Uniyal and Hore, 2009) • Ozyptila sp. (Hore and Uniyal, 2008a, b; Uniyal and Hore, 2009) • Pistius bhadurii Basu, 1965 (Basu, 1965; Tikader, 1971) • Runcinia insecta (L. Koch, 1875) (Hore and Uniyal, 2008a, b; Uniyal and Hore, 2009) • Runcinia roonwali Tikader, 1965 (Uniyal and Hore, 2009) • Runcinia sp. (Hore and Uniyal, 2008a, b) • Thomisus lobosus Tikader, 1965 (Lawania and Mathur, 2014a, b, c, d) • Thomisus pooneus Tikader, 1965 (Chandra et al., 2021) IAEES www.iaees.org Arthropods, 2022, 11(1): 18-55 33 • Thomisus projectus Tikader, 1960 (Lawania and Mathur, 2014a, b, c, d) • Thomisus pugilis Stoliczka, 1869 (Hore and Uniyal, 2008a, b; Uniyal and Hore, 2009) • Thomisus sorajaii Basu, 1963 (Chandra et al., 2021) • Thomisus unidentatus Dippenaar-Schoeman and van Harten, 2007 (Diksha et al., 2018) • Thomisus sp. (Khan and Misra, 2003; Kumar et al., 2017a; Chandra et al., 2021; Sujayanand et al., 2021) • Xysticus sp. (Anjali and Prakash, 2012; Kumar et al., 2017a, b) 3.1.34 Family Trachelidae • Trachelas himalayensis Biswas, 1993 (Hore and Uniyal, 2008a, b) 3.1.35 Family Uloboridae • Miagrammopes gravelyi Tikader, 1971 (Uniyal and Hore, 2009) • Miagrammopes indicus Tikader, 1971 (Uniyal and Hore, 2009) • Uloborus danolius Tikader, 1969 (Hore and Uniyal, 2008a, b; Uniyal and Hore, 2009; Lawania and Mathur, 2014a, b, c, d) • Uloborus sp. (Tandon and Lal, 1983; Hore and Uniyal, 2008a, b; Uniyal and Hore, 2009) • Zosis geniculata (Olivier, 1789) (Kumar et al., 2017a) 3.1.36 Family Zodariidae • Tropizodium bengalensis (Tikader and Patel, 1975) (Uniyal and Hore, 2009) • Tropizodium sp. (Hore and Uniyal, 2008a, b, c) 3.2 Uttarakhand Uttarakhand (formerly known as Uttaranchal) (Coordinates: 30.0668° N, 79.0193° E), known for the natural environment of the Himalayas, the Bhabar and the Terai regions and is bordered by Tibet Autonomous Region of China to the north; Nepal to the east; Uttar Pradesh to the south and Himachal Pradesh to the west and north-west. Administratively, it is divided into 13 districts (Fig. 2). Uttarakhand has a total area of 53,483 km2 of which about 86% is mountainous and 65% is covered by forest. Most of the northern part of the state is covered by high Himalayan peaks and glaciers. Two of the most pious rivers, Ganges and Yamuna originate in the Gangotri and Yamunotri glaciers of Uttarakhand and are fed by numerous lakes, glacial melts and streams. There are several parks and sanctuaries, Jim Corbett National Park is one of the oldest national parks in India. Being southern slope of the Himalaya range, the climate and vegetation vary greatly with elevation, from glaciers at the highest elevations which are covered by ice and bare rock to subtropical forests at the lower elevations. Depending upon the location and altitude, temperature varies from -4°C to 43°C. The flora and fauna of Uttarakhand are very rich. The perusal of literature reveals that Simon (1889) was probably the first who had described/recorded 24 species of spiders in Uttarakhand. Later, Simon (1897) again added 9 more species of spiders in Uttarakhand. Thereafter, Pocock (1899, 1900) described/recorded additional 9 species of spiders from the state. In the 20th century before independence of India (1947), few more workers recorded 25 more species of spiders, for example, Leardi in Airaghi (1901), 8 species; Simon (1906), 1 species; Strand (1907, 1909), 2 species; Gravely (1921, 1924, 1931), 8 species and Fage (1946), 4 species. After independence, Basu (1964) was probably the first who described 2 species of spiders from the Dehradun district, namely, Massuria roonwali and Pistius kanikae. Later, Basu (1965) again described 3 more species, Pistius barchensis, Pistius gangulyi and Pistius robustus. Thereafter, several workers sporadically described/recorded about a hundred of the species of spiders. The first faunistic survey of spiders was conducted by Biswas and Biswas (2010) who recorded and compiled IAEES www.iaees.org Arthropods, 2022, 11(1): 18-55 34 125 species belonging to 49 genera under 17 families in Uttarakhand. Later, Uniyal et al. (2011) recorded 88 species of spiders in Nanda Devi Biosphere Reserve situated in the Chamoli district. After one year, Gupta and Siliwal (2012) prepared a checklist of spiders of Wildlife Institute of India campus, Dehradun district of Uttarakhand. In recent years, Pooja et al. (2019) and Siddhu et al. (2020) recorded 31 and 27 species of spiders from Navdanya Biodiversity Farm located in Dehradun district and Nainital district of Uttarakhand, respectively. However, most of the national parks and sanctuaries, forest areas, agricultural fields of the states still await intensive and extensive survey programmes to record these spiders. Fig. 2 Number of species of spiders described/recorded from different districts of Uttarakhand. In the present compilation, a total of 373 species described under 202 genera belonging to 45 families were enlisted that have been recorded/described from 11 districts of Uttarakhand out of 13 districts giving up-todate information in the light of modern taxonomic concepts. The maximum number of species of spiders were recorded from the district Dehradun (240 species) followed by Chamoli (188 species), Pithoragarh (76 species), Almora (64 species), Nainital (50 species), Hardwar (43 species), Pauri Garhwal (30 species), Tehri Garhwal (16 species), Uttarkashi (9 species), Bageshwar (3 species), and Rudra Prayag (2 species). No faunal survey of spiders was so far conducted in 2 districts, Champawat and Udham Singh Nagar of Uttarakhand (Fig. 2). Hence, the intensive and extensive faunal survey is required in these areas. Following is the familywise list of species of spiders recorded/described from different districts of Uttarakhand. 3.2.1 Family Agelenidae • Agelena sp. (Quasin and Uniyal, 2013) IAEES www.iaees.org Arthropods, 2022, 11(1): 18-55 35 • Draconarius harduarae (Biswas and Roy, 2008) (Biswas and Roy, 2008) • Draconarius joshimath Quasin et al., 2017 (Quasin et al., 2017a) • Draconarius sp. (Uniyal et al., 2011) 3.2.2 Family Amaurobiidae • Amaurobius sp. (Uniyal et al., 2011) • Himalmartensus mussooriensis (Biswas and Roy, 2008) (Biswas and Roy, 2008) • Himalmartensus nandadevi Quasin et al., 2015 (Quasin et al., 2015) • Himalmartensus sp. (Uniyal et al., 2011) 3.2.3 Family Anyphaenidae • Anyphaena soricina Simon, 1889 (Simon, 1889) • Anyphaena sp. (Uniyal et al., 2011) 3.2.4 Family Araneidae • Araneus bilunifer Pocock, 1900 (Uniyal et al., 2011) • Araneus camilla (Simon, 1889) (Simon, 1889) • Araneus ellipticus (Tikader and Bal, 1981) (Uniyal et al., 2011; Quasin and Uniyal, 2013) • Araneus himalayanus (Simon, 1889) (Simon, 1889) • Araneus minutalis (Simon, 1889) (Simon, 1889) • Araneus mitificus (Simon, 1886) (Biswas and Biswas, 2010; Uniyal et al., 2011; Gupta and Siliwal, 2012; Siddhu et al., 2020) • Araniella nympha (Simon, 1889) (Simon, 1889; Uniyal et al., 2011; Quasin and Uniyal, 2013) • Araneus sp. (Quasin and Uniyal, 2010, 2011a; Uniyal et al., 2011; Pooja et al., 2019) • Araniella cucurbitina (Clerck, 1757) (Rajpoot et al., 2018) • Araniella maasdorpi Zamani and Marusik, 2020 (Zamani and Marusik, 2020) • Araniella sp. (Quasin and Uniyal, 2011a; Uniyal et al., 2011) • Argiope aemula (Walckenaer, 1837) (Biswas and Biswas, 2010; Gupta and Siliwal, 2012) • Argiope anasuja Thorell, 1887 (Biswas and Biswas, 2010; Uniyal et al., 2011; Gupta and Siliwal, 2012) • Argiope catenulata (Doleschall 1859) (Biswas and Biswas, 2010) • Argiope minuta Karsch, 1879 (Biswas and Biswas, 2010) • Argiope pulchella Thorell, 1881 (Uniyal and Hore, 2006; Biswas and Biswas, 2010; Gupta and Siliwal, 2012; Siddhu et al., 2020) • Argiope trifasciata (Forsskål, 1775) (Biswas and Biswas, 2010) • Argiope sp. (Quasin and Uniyal, 2010; Uniyal et al., 2011) • Cercidia punctigera Simon, 1889 (Simon, 1889) • Chorizopes sp. (Uniyal et al., 2011) • Cyclosa bifida (Doleschall, 1859) (Gupta and Siliwal, 2012; Siddhu et al., 2020) • Cyclosa confraga (Thorell, 1892) (Quasin and Uniyal, 2010, 2011a, 2013; Uniyal et al., 2011) • Cyclosa gossypiata Keswani, 2013 (Siddhu et al., 2020) • Cyclosa hexatuberculata Tikader, 1982 (Uniyal et al., 2011) • Cyclosa insulana (Costa, 1834) (Quasin and Uniyal, 2011a; Uniyal et al., 2011) • Cyclosa quinqueguttata (Thorell, 1881) (Simon, 1889) • Cyclosa simoni Tikader, 1982 (Biswas and Biswas, 2010) • Cyclosa spirifera Simon, 1889 (Simon, 1889; Biswas and Biswas, 2010) IAEES www.iaees.org 36 Arthropods, 2022, 11(1): 18-55 • Cyclosa sp. (Uniyal et al., 2011) • Cyrtarachne raniceps Pocock, 1900 (Gupta and Siliwal, 2012) • Cyrtarachne sp. (Uniyal et al., 2011) • Cyrtophora citricola (Forsskål, 1775) (Siddhu et al., 2020) • Cyrtophora moluccensis (Doleschall, 1857) (Quasin and Uniyal, 2011a, 2013; Uniyal et al., 2011) • Cyrtophora sp. (Quasin and Uniyal, 2010; Uniyal et al., 2011) • Eriophora sp. (Uniyal et al., 2011; Uniyal et al., 2011; Quasin and Uniyal, 2013) • Eriovixia excelsa (Simon, 1889) (Simon, 1889) • Eriovixia laglaizei (Simon, 1877) (Simon, 1889; Pocock, 1900; Gupta and Siliwal, 2012; Pooja et al., 2019; Siddhu et al., 2020) • Eriovixia poonaensis (Tikader and Bal, 1981) (Gupta and Siliwal, 2012) • Eriovixia sp. (Uniyal et al., 2011) • Gasteracantha unguifera Simon, 1889 (Simon, 1889; Pocock, 1900; Tikader, 1982) • Gea sp. (Gupta and Siliwal, 2012; Pooja et al., 2019) • Gea subarmata Thorell, 1890 (Tikader, 1982; Levi, 1983; Uniyal and Hore, 2006, 2009) • Larinia chloris Audoin (1826) (Biswas and Biswas, 2010) • Larinia phthisica (L. Koch, 1871) (Biswas and Biswas, 2010) • Larinia sp. (Quasin and Uniyal, 2011a; Gupta and Siliwal, 2012) • Lipocrea fusiformis (Thorell, 1877) (Simon, 1889) • Neoscona achine (Simon, 1906) (Quasin and Uniyal, 2011a; Uniyal et al., 2011) • Neoscona bengalensis Tikader and Bal, 1981 (Quasin and Uniyal, 2011a; Gupta and Siliwal, 2012) • Neoscona biswasi Bhandari and Gajbe, 2001 (Quasin and Uniyal, 2011a; Uniyal et al., 2011) • Neoscona chrysanthusi Tikader and Bal, 1981 (Gupta and Siliwal, 2012) • Neoscona inusta (L. Koch, 1871) (Quasin and Uniyal, 2013) • Neoscona mukerjei Tikader, 1980 (Biswas and Biswas, 2010; Uniyal and Hore, 2006; Quasin and Uniyal, 2010, 2011a, 2013; Uniyal et al., 2011; Pooja et al., 2019) • Neoscona nautica (L. Koch, 1875) (Biswas and Biswas, 2010; Quasin and Uniyal, 2010, 2011a; Uniyal et al., 2011; Gupta and Siliwal, 2012; Siddhu et al., 2020) • Neoscona odites (Simon, 1906) (Biswas and Biswas, 2010) • Neoscona pavida (Simon, 1906) (Biswas and Biswas, 2010) • Neoscona shillongensis Tikader and Bal, 1981 (Biswas and Biswas, 2010; Quasin and Uniyal, 2011a; Uniyal et al., 2011) • Neoscona sinhagadensis (Tikader, 1975) (Gupta and Siliwal, 2012) • Neoscona theisi (Walckenaer, 1837) (Biswas and Biswas, 2010; Quasin and Uniyal, 2011a, 2013; Uniyal et al., 2011; Gupta and Siliwal, 2012; Pooja et al., 2019; Siddhu et al., 2020) • Neoscona vigilans (Blackwall, 1865) (Biswas and Biswas, 2010; Quasin and Uniyal, 2011a; Uniyal et al., 2011; Gupta and Siliwal, 2012) • Neoscona sp. (Uniyal et al., 2011; Pooja et al., 2019) • Nephila dirangensis Biswas and Biswas, 2006 (Biswas and Biswas, 2010) • Nephila kuhlii (Doleschall, 1859) (Biswas and Biswas, 2010) • Nephila pilipes (Fabricius, 1793) (Simon, 1889; Pocock, 1900; Biswas and Biswas, 2010; Gupta and Siliwal, 2012; Siddhu et al., 2020) • Nephilengys malabarensis (Walckenaer, 1841) (Biswas and Biswas, 2010) • Parawixia dehaani (Doleschall, 1859) (Uniyal and Hore, 2006; Quasin and Uniyal, 2010, 2011a; Uniyal et IAEES www.iaees.org Arthropods, 2022, 11(1): 18-55 37 al., 2011; Gupta and Siliwal, 2012) • Parawixia sp. (Uniyal et al., 2011) • Plebs himalayaensis (Tikader, 1975) (Uniyal et al., 2011; Pooja et al., 2019) • Thelacantha brevispina (Doleschall, 1857) (Pocock, 1900; Simon, 1897; Tikader, 1982; Quasin and Uniyal, 2011a; Uniyal et al., 2011) • Trichonephila clavata (L. Koch, 1878) (Biswas and Biswas, 2010; Uniyal et al., 2011) 3.2.5 Family Atypidae • Atypus wii Siliwal et al., 2014 (Siliwal et al., 2014) 3.2.6 Family Cheiracanthiidae • Cheiracanthium danieli Tikader, 1975 (Quasin and Uniyal, 2011a) • Cheiracanthium himalayense Gravely, 1931 (Majumder and Tikader, 1991; Biswas and Biswas, 2010) • Cheiracanthium melanostomum (Thorell, 1895) (Biswas and Biswas, 2010; Pooja et al., 2019) • Cheiracanthium pauriense Majumder and Tikader, 1991 (Majumder and Tikader, 1991; Biswas and Biswas, 2010) • Cheiracanthium rupicola (Thorell, 1897) (Uniyal et al., 2011) • Cheiracanthium sadanai Tikader, 1976 (Biswas and Biswas, 2010) • Cheiracanthium sp. (Quasin and Uniyal, 2010; Uniyal et al., 2011; Gupta and Siliwal, 2012; Pooja et al., 2019) • Cheiracanthium triviale (Thorell, 1895) (Biswas and Biswas, 2010) 3.2.7 Family Clubionidae • Clubiona chakrabartei Majumder and Tikader, 1991 (Majumder and Tikader, 1991; Biswas and Biswas, 2010) • Clubiona diversa O. P.-Cambridge, 1862 (Biswas and Biswas, 2010) • Clubiona drassodes Pickard-Cambridge, 1874 (Majumder and Tikader, 1991; Biswas and Biswas, 2010) • Clubiona hysgina Simon, 1889 (Simon, 1889; Biswas and Biswas, 2010) • Clubiona shillongensis Majumder and Tikader, 1991 (Biswas and Biswas, 2010) • Clubiona tikaderi Majumder and Tikader, 1991 (Biswas and Biswas, 2010) • Clubiona sp. (Quasin and Uniyal, 2011a; Gupta and Siliwal, 2012; Quasin and Uniyal, 2010; Quasin and Uniyal, 2013) 3.2.8 Family Corinnidae • Apochinomma dolosum Simon, 1897 (Simon, 1897; Majumder and Tikader, 1991; Biswas and Biswas, 2010) • Castianeira zetes Simon, 1897 (Uniyal et al., 2011) • Corinnomma severum (Thorell, 1877) (Simon, 1897) • Castianeira sp. (Gupta and Siliwal, 2012) 3.2.9 Family Ctenidae • Amauropelma staschi Jager, 2012 (Jäger, 2012) • Anahita smythiesi (Simon, 1897) (Simon, 1897; Gravely, 1931; Tikader and Malhotra, 1981) • Nilus sp. (Jithin et al., 2021) 3.2.10 Family Cybaeidae • Cedicus bucculentus Simon, 1889 (Simon, 1889) IAEES www.iaees.org 38 Arthropods, 2022, 11(1): 18-55 3.2.11 Family Dictynidae • Dictyna sp. (Quasin and Uniyal, 2010; Gupta and Siliwal, 2012) 3.2.12 Family Eresidae • Stegodyphus sarasinorum Karsch, 1892 (Biswas and Biswas, 2010) 3.2.13 Family Filistatidae • Pritha sp. (Gupta and Siliwal, 2012) 3.2.14 Family Gnaphosidae • Callilepis rajani Gajbe, 1983 (Gajbe, 1983) • Camillina smythiesi (Simon, 1897) (Simon, 1897; Tikader, 1982) • Drassodes deoprayagensis Tikader and Gajbe, 1975 (Tikader and Gajbe, 1975; Tikader, 1982; Gajbe, 1988) • Drassodes gangeticus Tikader and Gajbe, 1975 (Tikader and Gajbe, 1975; Tikader, 1982) • Drassodes himalayensis Tikader and Gajbe, 1975 (Tikader and Gajbe, 1975; Tikader, 1982; Gajbe, 1988, 2005) • Drassodes luridus (Pickard-Cambridge, 1874) (Biswas and Biswas, 2010) • Drassodes sirmourensis (Tikader and Gajbe, 1977) (Biswas and Biswas, 2010) • Drassodes sitae Tikader and Gajbe, 1975 (Tikader and Gajbe, 1975; Tikader, 1982; Gajbe, 1988) • Drassodes viveki (Gajbe, 1992) (Gajbe, 1992b) • Drassodes sp. (Quasin and Uniyal, 2011a, 2013; Uniyal et al., 2011) • Gnaphosa kankhalae Biswas and Roy, 2008 (Biswas and Roy, 2008) • Gnaphosa pauriensis Tikader and Gajbe, 1977 (Tikader and Gajbe, 1977a; Tikader, 1982; Gajbe, 1988) • Gnaphosa poonaensis Tikader, 1973 (Biswas and Biswas, 2010; Uniyal et al., 2011) • Gnaphosa sp. (Quasin and Uniyal, 2010, 2011a; Uniyal et al., 2011) • Haplodrassus dumdumensis Tikader, 1982 (Biswas and Biswas, 2010) • Haplodrassus sataraensis Tikader and Gajbe, 1977 (Gajbe, 1988; Biswas and Biswas, 2010) • Haplodrassus tehriensis Tikader and Gajbe, 1977 (Tikader and Gajbe, 1977b; Gajbe, 1988) • Herpyllus sp. (Uniyal et al., 2011; Pooja et al., 2019) • Megamyrmaekion pritiae (Tikader, 1982) (Biswas and Biswas, 2010) • Poecilochroa kuljitae (Tikader, 1982) (Biswas and Biswas, 2010) • Poecilochroa sedula (Simon, 1897) (Simon, 1897; Tikader, 1982) • Scotophaeus madalasae Tikader and Gajbe, 1977 (Tikader and Gajbe, 1977c; Gajbe, 1988) • Scotophaeus sp. (Quasin and Uniyal, 2011a; Uniyal et al., 2011) • Setaphis subtilis (Simon, 1897) (Biswas and Biswas, 2010) • Zelotes mandlaensis Tikader and Gajbe, 1976 (Biswas and Biswas, 2010) • Zelotes nainitalensis Tikader and Gajbe, 1976 (Tikader and Gajbe, 1976a; Tikader, 1982; Biswas and Biswas, 2010) • Zelotes sataraensis Tikader and Gajbe, 1979 (Gajbe, 1988) • Zelotes surekhae Tikader and Gajbe, 1976 (Biswas and Biswas, 2010) • Zelotes sp. (Quasin and Uniyal, 2010, 2011a; Uniyal et al., 2011) 3.2.15 Family Hahniidae • Hahnia sp. (Uniyal et al., 2011) 3.2.16 Family Hersiliidae IAEES www.iaees.org Arthropods, 2022, 11(1): 18-55 39 • Hersilia savignyi Lucas, 1836 (Gajbe, 2007; Biswas and Biswas, 2010; Gupta and Siliwal, 2012; Pooja et al., 2019) • Hersilia sp. (Uniyal et al., 2011; Pooja et al., 2019) 3.2.17 Family Idiopidae • Heligmomerus wii Siliwal et al., 2020 (Siliwal et al., 2020) 3.2.18 Family Linyphiidae • Agyneta sp. (Uniyal et al., 2011) • Anguliphantes nepalensis Tanasevitch, 2011 (Tanasevitch, 2011) • Atypena adelinae Barrion and Litsinger, 199 (Uniyal et al., 2011) • Bathyphantes sp. (Uniyal et al., 2011) • Caviphantes pseudosaxetorum Wunderlich, 1979 (Tanasevitch, 2011) • Cresmatoneta leucophthalma (Fage, 1946) (Fage, 1946) • Erigone sp. (Quasin and Uniyal, 2011a) • Gongylidiellum confusum Thaler, 1987 (Tanasevitch, 2011) • Gongylidioides pectinatus Tanasevitch, 2011 (Tanasevitch, 2011) • Lepthyphantes rudrai Tikader, 1970 (Biswas and Biswas, 2010) • Linyphia sikkimensis Tikader, 1970 (Biswas and Biswas, 2010) • Linyphia sp. (Quasin and Uniyal, 2010, 2011a; Uniyal et al., 2011; Gupta and Siliwal, 2012) • Microbathyphantes palmarius (Marples, 1955) (Tanasevitch, 2011) • Microlinyphia sp. (Uniyal et al., 2011) • Neriene birmanica (Thorell, 1887) (Pooja et al., 2019; Siddhu et al., 2020) • Neriene macella (Thorell, 1898) (Tanasevitch, 2017) • Neriene sp. (Quasin and Uniyal, 2010, 2011a; Uniyal et al., 2011) • Neriene sundaica (Simon, 1905) (Gupta and Siliwal, 2012) • Pelecopsis indus Tanasevitch, 2011 (Tanasevitch, 2011) • Pityohyphantes sp. (Uniyal et al., 2011) • Scotargus pilosus Simon, 1913 (Tanasevitch, 2011) • Tiso incisus Tanasevitch, 2011 (Tanasevitch, 2011) 3.2.19 Family Liocranidae • Agroeca gangotrae Biswas and Roy, 2008 (Biswas and Roy, 2008) • Oedignatha procerula Simon, 1897 (Simon, 1897; Majumder and Tikader, 1991; Biswas and Biswas, 2010) • Oedignatha sp. (Uniyal et al., 2011) • Paratus indicus Marusik, Zheng and Li, 2008 (Marusik et al., 2008) •Sphingius nainitalensis (Gajbe, 1979) (Gajbe, 1979; Biswas and Biswas, 2010) 3.2.20 Family Lycosidae • Arctosa himalayensis Tikader and Malhotra, 1980 (Tikader and Malhotra, 1980) • Arctosa indica Tikader and Malhotra, 1980 (Biswas and Biswas, 2010) • Arctosa khudiensis (Sinha, 1951) (Uniyal and Hore, 2006) • Arctosa mulani (Dyal, 1935) (Biswas and Biswas, 2010) • Arctosa sp. (Gupta and Siliwal, 2012) • Draposa atropalpis (Gravely, 1924) (Biswas and Biswas, 2010) IAEES www.iaees.org 40 Arthropods, 2022, 11(1): 18-55 • Draposa burasantiensis (Tikader and Malhotra, 1976) (Biswas and Biswas, 2010) • Draposa lyrivulva (Bösenberg and Strand, 1906) (Biswas and Biswas, 2010) • Draposa oakleyi (Gravely, 1924) (Biswas and Biswas, 2010) • Evippa rajasthanea Tikader and Malhotra, 1980 (Uniyal and Hore, 2006) • Evippa sohani Tikader and Malhotra, 1980 (Uniyal and Hore, 2006) • Hippasa agelenoides (Simon, 1884) (Simon, 1897; Tikader and Malhotra, 1980; Biswas and Biswas, 2010; Quasin and Uniyal, 2010, 2011a) • Hippasa greenalliae (Blackwall, 1867) (Biswas and Biswas, 2010; Quasin and Uniyal, 2013) • Hippasa holmerae Thorell, 1895 (Tikader and Malhotra, 1980; Biswas and Biswas, 2010) • Hippasa loundesi Gravely, 1924 (Biswas and Biswas, 2010) • Hippasa lycosina Pocock, 1900 (Tikader and Malhotra, 1980; Biswas and Biswas, 2010) • Hippasa madraspatana Gravely, 1924 (Biswas and Biswas, 2010) • Hippasa olivacea (Thorell, 1887) (Leardi in Airaghi, 1901; Bastawade and Borkar, 2008) • Hippasa pisaurina Pocock, 1900 (Uniyal and Hore, 2006) • Hogna himalayensis (Gravely, 1924) (Biswas and Biswas, 2010) • Hogna stictopyga (Thorell, 1895) (Leardi in Airaghi, 1901) • Lycosa carmichaeli Gravely, 1924 (Gravely, 1924; Tikader and Malhotra, 1980; Bastawade and Borkar, 2008) • Lycosa chaperi Simon, 1885 (Biswas and Biswas, 2010) • Lycosa fuscana Pocock, 1901 (Biswas and Biswas, 2010) • Lycosa indagatrix Walckenaer, 1837 (Biswas and Biswas, 2010) • Lycosa iranii Pocock, 1901 (Biswas and Biswas, 2010) • Lycosa lambai Tikader and Malhotra, 1980 (Biswas and Biswas, 2010) • Lycosa madani Pocock, 1901 (Biswas and Biswas, 2010) • Lycosa mahabaleshwarensis Tikader and Malhotra, 1980 (Biswas and Biswas, 2010) • Lycosa nigrotibialis Simon, 1884 (Biswas and Biswas, 2010) • Lycosa phipsoni Pocock, 1899 (Biswas and Biswas, 2010) • Lycosa pictula Pocock, 1901 (Biswas and Biswas, 2010) • Lycosa prolifica Pocock, 1901 (Tikader and Malhotra, 1980; Biswas and Biswas, 2010) • Lycosa shillongensis Tikader and Malhotra, 1980 (Biswas and Biswas, 2010) • Lycosa tista Tikader, 1970 (Uniyal et al., 2011; Biswas and Biswas, 2010; Siddhu et al., 2020) • Lycosa sp. (Quasin and Uniyal, 2010, 2011a; Uniyal et al., 2011; Pooja et al., 2019) • Pardosa algoides Schenkel, 1963 (Biswas and Biswas, 2010) • Pardosa altitudis Tikader and Malhotra, 1980 (Tikader and Malhotra, 1980) • Pardosa fletcheri (Gravely, 1924) (Tikader and Malhotra, 1980) • Pardosa heterophthalma (Simon, 1898) (Biswas and Biswas, 2010) • Pardosa kupupa (Tikader, 1970) (Biswas and Biswas, 2010) • Pardosa minuta Tikader and Malhotra, 1976 (Biswas and Biswas, 2010) • Pardosa mukundi Tikader and Malhotra, 1980 (Biswas and Biswas, 2010) • Pardosa pseudoannulata (Bösenberg and Strand, 1906) (Biswas and Biswas, 2010; Quasin and Uniyal, 2011a; Siddhu et al., 2020) • Pardosa rhenockensis (Tikader, 1970) (Biswas and Biswas, 2010) • Pardosa shyamae (Tikader, 1970) (Biswas and Biswas, 2010; Pooja et al., 2019) • Pardosa songosa Tikader and Malhotra, 1976 (Tikader and Malhotra, 1976; Gupta and Siliwal, 2012; Pooja IAEES www.iaees.org Arthropods, 2022, 11(1): 18-55 41 et al., 2019) • Pardosa sumatrana (Thorell, 1890) (Biswas and Biswas, 2010; Quasin and Uniyal, 2010, 2011a; Siddhu et al., 2020) • Pardosa sp. (Quasin and Uniyal, 2010; Pooja et al., 2019) • Trochosa himalayensis Tikader and Malhotra, 1980 (Tikader and Malhotra, 1980; Biswas and Biswas, 2010) • Trochosa punctipes (Gravely, 1924) (Biswas and Biswas, 2010) • Trochosa sp. (Uniyal et al., 2011) • Wadicosa fidelis (O. Pickard-Cambridge, 1872) (Tikader and Malhotra, 1980; Biswas and Biswas, 2010; Gupta and Siliwal, 2012) • Wadicosa quadrifera (Gravely, 1924) (Biswas and Biswas, 2010) 3.2.21 Family Mimetidae • Mimetus sp. (Uniyal et al., 2011) 3.2.22 Family Nesticidae • Nesticella nepalensis (Hubert, 1973) (Lin et al., 2016) 3.2.23 Family Oecobiidae • Oecobius sp. (Uniyal et al., 2011; Gupta and Siliwal, 2012) 3.2.24 Family Oonopidae • Camptoscaphiella fulva Caporiacco, 1935 (Baehr and Ubick, 2010) • Camptoscaphiella glenniei (Fage, 1946) (Fage, 1946; Grismado et al., 2014) • Dysderoides typhlos Fage, 1946 (Fage, 1946; Grismado et al., 2014) • Pelicinus lachivala Platnick et al., 2012 (Platnick et al., 2012) 3.2.25 Family Oxyopidae • Hamadruas sikkimensis (Tikader, 1970) (Uniyal and Hore, 2006) • Hamataliwa sp. (Uniyal et al., 2011; Pooja et al., 2019) • Oxyopes javanus Thorell, 1887 (Quasin and Uniyal, 2010; Uniyal et al., 2011; Gupta and Siliwal, 2012; Pooja et al., 2019) • Oxyopes kusumae Gajbe, 1999 (Pooja et al., 2019) • Oxyopes pankaji Gajbe and Gajbe, 2000 (Siddhu et al., 2020) • Oxyopes shweta Tikader 1970 (Biswas and Biswas, 2010; Uniyal et al., 2011; Gupta and Siliwal, 2012; Pooja et al., 2019) • Oxyopes sp. (Quasin and Uniyal, 2011a; Uniyal et al., 2011; Gupta and Siliwal, 2012; Pooja et al., 2019; Siddhu et al., 2020) • Peucetia latikae Tikader, 1970 (Biswas and Biswas, 2010) • Peucetia viridana (Stoliczka, 1869) (Simon, 1889; Biswas and Biswas, 2010; Gupta and Siliwal, 2012) • Peucetia sp. (Uniyal et al., 2011) 3.2.26 Family Palpimanidae • Palpimanus sp. (Uniyal et al., 2011) 3.2.27 Family Philodromidae • Philodromus chambaensis Tikader, 1980 (Tikader, 1980; Quasin and Uniyal, 2010; Uniyal et al., 2011) IAEES www.iaees.org 42 Arthropods, 2022, 11(1): 18-55 • Philodromus sp. (Quasin and Uniyal, 2011a; Uniyal et al., 2011) • Tibellus elongatus Tikader, 1960 (Gupta and Siliwal, 2012) • Tibellus sp. (Quasin and Uniyal, 2011a) 3.2.28 Family Pholcidae • Artema atlanta Walckenaer, 1837 (Gupta and Siliwal, 2012) • Crossopriza lyoni (Blackwall, 1867) (Biswas and Biswas, 2010; Gupta and Siliwal, 2012; Siddhu et al., 2020) • Pholcus djelalabad Senglet, 2008 (Huber, 2011) • Pholcus phalangioides (Fuesslin, 1775) (Quasin and Uniyal, 2010, 2011a) • Pholcus sp. (Gupta and Siliwal, 2012) 3.2.29 Family Pimoidae • Pimoa crispa (Fage, 1946) (Fage, 1946) • Pimoa nainital Zhang and Li, 2021 (Lin et al., 2021) 3.2.30 Family Pisauridae • Nilus albocinctus (Doleschall, 1859) (Pooja et al., 2019) • Perenethis dentifasciata (O. Pickard-Cambridge, 1885) (Gupta and Siliwal, 2012) • Perenethis sp. (Uniyal et al., 2011) • Perenethis venusta L. Koch, 1878 (Gupta and Siliwal, 2012) • Pisaura mirabilis (Clerck, 1757) (Uniyal et al., 2011) • Pisaura sp. (Quasin and Uniyal, 2013; Uniyal et al., 2011; Gupta and Siliwal, 2012) 3.2.31 Family Psechridae • Psechrus himalayanus Simon, 1906 (Simon, 1906; Levi, 1982; Quasin and Uniyal, 2010, 2011a; Uniyal et al., 2011; Bayer, 2012) • Psechrus torvus (O. Pickard-Cambridge, 1869) (Gupta and Siliwal, 2012) 3.2.32 Family Salticidae • Aelurillus quadrimaculatus Simon, 1889 (Simon, 1889) • Asemonea tenuipes (Pickard-Cambridge, 1869) (Gupta and Siliwal, 2012) • Bianor albobimaculatus (Lucas, 1846) (Logunov, 2019) • Bianor angulosus (Karsch, 1879) (Siddhu et al., 2020) • Bianor balius Thorell, 1890 (Pooja et al., 2019) • Bianor narmadaensis (Tikader, 1975) (Biswas and Biswas, 2010) • Bianor pashanensis (Tikader, 1975) (Biswas and Biswas, 2010) • Bianor sp. (Gupta and Siliwal, 2012) • Brettus anchorum Wanless, 1979 (Gupta and Siliwal, 2012; Pooja et al., 2019) • Carrhotus erus Jastrzębski, 1999 (Logunov, 2021) • Carrhotus sannio (Thorell, 1877) (Logunov, 2021) • Carrhotus sp. (Quasin and Uniyal, 2010; Uniyal et al., 2011) • Chrysilla volupe (Karsch, 1879) (Caleb et al., 2018a) • Cosmophasis sp. (Quasin and Uniyal, 2010) • Epeus indicus Prószyński, 1992 (Gupta and Siliwal, 2012) • Epocilla aurantiaca (Simon, 1885) (Gupta and Siliwal, 2012) • Evarcha pococki Zabka, 1985 (Gupta and Siliwal, 2012) IAEES www.iaees.org Arthropods, 2022, 11(1): 18-55 43 • Evarcha sp. (Pooja et al., 2019; Siddhu et al., 2020) • Hasarius adansoni (Audouin, 1826) (Gupta and Siliwal, 2012) • Heliophanus curvidens (O. P.-Cambridge. 1872) (Uniyal et al., 2011) • Hyllus semicupreus (Simon, 1885) (Biswas and Biswas, 2010; Quasin and Uniyal, 2010; Pooja et al., 2019) • Hyllus sp. (Uniyal et al., 2011) • Marengo crassipes Peckham and Peckham, 1892 (Gupta and Siliwal, 2012) • Marpissa pauariensis Biswas and Roy, 2008 (Biswas and Roy, 2008) • Menemerus bivittatus (Dufour, 1831) (Gupta and Siliwal, 2012) • Menemerus sp. (Gupta and Siliwal, 2012; Siddhu et al., 2020) • Myrmaplata plataleoides (O. Pickard-Cambridge, 1869) (Pooja et al., 2019) • Myrmarachne melanocephala MacLeay, 1839 (Biswas and Biswas, 2010; Quasin and Uniyal, 2011a; Uniyal et al., 2011; Yadav and Prakash, 2021) • Myrmarachne prava (Karsch, 1880) (Biswas and Biswas, 2010) • Myrmarachne sp. (Uniyal et al., 2011; Gupta and Siliwal, 2012; Quasin and Uniyal, 2013) • Nandicius frigidus (Pickard-Cambridge, 1885) (Simon, 1889) • Nandicius mussooriensis (Prószyński, 1992) (Prószyński, 1992) • Nandicius vallisflorum Caleb et al., 2018 (Caleb et al., 2018b) • Onomastus sp. (Gupta and Siliwal, 2012) • Orientattus aurantius (Kanesharatnam and Benjamin, 2018) (Caleb and Acharya, 2019) • Pancorius sp. (Gupta and Siliwal, 2012) • Pellenes himalaya Caleb et al., 2018 (Caleb et al., 2018b) • Pellenes sp. (Uniyal et al., 2011) • Phintella vittata (C L Koch, 1846) (Sherriffs, 1931; Gupta and Siliwal, 2012; Pooja et al., 2019) • Phintella sp. (Uniyal et al., 2011) • Phintelloides versicolor (C. L. Koch, 1846) (Gupta and Siliwal, 2012) • Phlegra dhakuriensis (Tikader, 1974) (Biswas and Biswas, 2010) • Phlegra sp. (Uniyal et al., 2011) • Plexippus calcutaensis (Tikader, 1974) (Biswas and Biswas, 2010) • Plexippus paykulli (Audouin, 1826) (Simon, 1889; Biswas and Biswas, 2010; Uniyal et al., 2011; Gupta and Siliwal, 2012; Pooja et al., 2019; Siddhu et al., 2020; Logunov, 2021) • Plexippus sp. (Quasin and Uniyal, 2010, 2013; Uniyal et al., 2011; Gupta and Siliwal, 2012; Pooja et al., 2019) • Portia albimana (Simon, 1900) (Simon, 1900; Sherriffs, 1931; Wanless, 1978) • Portia sp. (Gupta and Siliwal, 2012) • Pseudicius sp. (Uniyal et al., 2011) • Rhene danieli Tikader, 1973 (Quasin and Uniyal, 2010; Uniyal et al., 2011) • Rhene flavicomans Simon, 1902 (Uniyal et al., 2011; Gupta and Siliwal, 2012) • Rhene flavigera (C.L. Koch, 1846) (Quasin and Uniyal, 2010; Pooja et al., 2019) • Rhene mus (Simon, 1889 (Simon, 1889) • Rhene rubrigera (Thorell, 1887) (Gupta and Siliwal, 2012) • Rhene sp. (Quasin and Uniyal, 2011a; Uniyal et al., 2011; Pooja et al., 2019) • Salticus sp. (Uniyal et al., 2011) • Siler sp. (Quasin and Uniyal, 2010, 2011a; Uniyal et al., 2011) • Stenaelurillus sp. (Uniyal et al., 2011; Siddhu et al., 2020) IAEES www.iaees.org 44 Arthropods, 2022, 11(1): 18-55 • Synagelides martensi Bohdanowicz, 1987 (Logunov and Hereward, 2006) • Telamonia dimidiata (Simon, 1899) (Biswas and Biswas, 2010; Gupta and Siliwal, 2012; Pooja et al., 2019; Siddhu et al., 2020) • Thiania bhamoensis Thorell, 1887 (Gupta and Siliwal, 2012) • Thiania sp. (Uniyal et al., 2011) • Thyene bivittata Xie and Peng, 1995 (Logunov, 2021) • Thyene imperialis (Rossi 1846) (Logunov, 2021) • Thyene sp. (Gupta and Siliwal, 2012) 3.2.33 Family Scytodidae • Scytodes propinqua Stoliczka, 1869 (Simon, 1897) • Scytodes thoracica (Latreille, 1802) (Uniyal et al., 2011) • Scytodes sp. (Uniyal et al., 2011) 3.2.34 Family Segestriidae • Segestria sp. (Uniyal et al., 2011) 3.2.35 Family Selenopidae • Makdiops agumbensis (Tikader, 1969) (Biswas and Biswas, 2010) • Makdiops montigena (Simon, 1889) (Simon, 1889; Pocock, 1900; Gravely, 1931; Crews and Harvey, 2011; Sankaran et al., 2020a) • Selenops radiatus Latreille, 1819 (Leardi in Airaghi, 1901; Quasin and Uniyal, 2010; Uniyal et al., 2011; Kumar et al., 2017a) • Selenops sp. (Quasin and Uniyal, 2011a, 2013) 3.2.36 Family Sparassidae • Heteropoda bhaikakai Patel and Patel, 1973 (Biswas and Biswas, 2010) • Heteropoda kandiana Pocock, 1899 (Biswas and Biswas, 2010) • Heteropoda kuluensis Sethi and Tikader, 1988 (Biswas and Biswas, 2010) • Heteropoda leprosa Simon, 1884 (Sethi and Tikader, 1988) • Heteropoda nilgirina Pocock, 1901 (Sethi and Tikader, 1988) • Heteropoda pedata Strand, 1907 (Strand, 1907, 1909) • Heteropoda phasma Simon, 1897 (Pocock, 1900; Sethi and Tikader, 1988; Biswas and Biswas, 2010) • Heteropoda venatoria (Linnaeus 1767) (Leardi in Airaghi, 1901; Biswas and Biswas, 2010; Quasin and Uniyal, 2010, 2011a, 2013; Uniyal et al., 2011; Siddhu et al., 2020) • Heteropoda sp. (Uniyal et al., 2011; Gupta and Siliwal, 2012) • Olios milleti (Pocock, 1901) (Gupta and Siliwal, 2012) • Olios punctipes Simon, 1884 (Strand, 1909) • Olios rosettii (Leardi in Airaghi, 1901) (Leardi in Airaghi, 1901) • Olios sanguinifrons (Simon, 1906) (Uniyal et al., 2011; Pooja et al., 2019) • Olios sp. (Quasin and Uniyal, 2010, 2011a; Uniyal et al., 2011) • Pseudopoda casaria (Simon, 1897) (Gravely, 1931; Jäger, 2001) • Pseudopoda lutea (Thorell, 1895) (Leardi in Airaghi, 1901) • Pseudopoda prompta (Pickard-Cambridge, 1885) (Pocock, 1900; Strand, 1909; Sethi and Tikader, 1988; Jäger, 2001; Quasin and Uniyal, 2010, 2011a; Uniyal et al., 2011) IAEES www.iaees.org Arthropods, 2022, 11(1): 18-55 45 • Pseudopoda sp. (Uniyal et al., 2011) • Spariolenus tigris Simon, 1880 (Biswas and Biswas, 2010) 3.2.37 Family Tetrablemmidae • Tetrablemma loebli Bourne, 1980 (Bourne, 1980; Sankaran and Sebastian, 2016) 3.2.38 Family Tetragnathidae • Dyschiriognatha sp. (Uniyal et al., 2011) • Guizygeilla sp. (Uniyal et al., 2011) • Guizygiella indica (Tikader and Bal, 1980) (Hore and Uniyal, 2008a; Uniyal and Hore, 2009; Quasin and Uniyal, 2013) • Guizygiella sp. (Uniyal et al., 2011) • Leucauge celebesiana (Walckenaer, 1841) (Biswas and Biswas, 2010; Quasin and Uniyal, 2011a; Uniyal et al., 2011) • Leucauge decorata (Blackwall, 1864) (Simon, 1889; Gravely, 1921; Tikader, 1982; Uniyal and Hore, 2006; Gupta and Siliwal, 2012; Biswas and Biswas, 2010; Quasin and Uniyal, 2010, 2011a, 2013; Uniyal et al., 2011; Pooja et al., 2019; Siddhu et al., 2020) • Leucauge parangscipinia Barrion and Litsinger, 1995 (Gupta and Siliwal, 2012) • Leucauge tessellata (Thorell, 1887) (Biswas and Biswas, 2010) • Leucauge sp. (Uniyal et al., 2011) • Metellina sp. (Uniyal et al., 2011) • Opadometa fastigata (Simon, 1877) (Gravely, 1921; Tikader, 1982; Gupta and Siliwal, 2012) • Tetragnatha andamanensis Tikader, 1977 (Biswas and Biswas, 2010) • Tetragnatha javana (Thorell, 1890) (Biswas and Biswas, 2010; Siddhu et al., 2020) • Tetragnatha keyserlingi Simon, 1890 (Uniyal et al., 2011; Gupta and Siliwal, 2012) • Tetragnatha mandibulata Walckenaer, 1842 (Gupta and Siliwal, 2012) • Tetragnatha sp. (Quasin and Uniyal, 2010, 2011a; Uniyal et al., 2011; Gupta and Siliwal, 2012; Siddhu et al., 2020) • Tylorida striata (Thorell, 1877) (Siddhu et al., 2020) • Tylorida ventralis (Thorell, 1877) (Siddhu et al., 2020) 3.2.39 Family Theraphosidae • Haplocosmia himalayana (Pocock, 1899) (Pocock, 1899, 1900; Siliwal et al., 2011; Gupta and Siliwal, 2012) • Chilobrachys himalayensis (Tikader, 1977) (Biswas and Biswas, 2010) • Chilobrachys khasiensis (Tikader, 1977) (Biswas and Biswas, 2010) • Lyrognathus saltator Pocock, 1900 (Siddhu et al., 2020) • Poecilotheria regalis Pocock, 1899 (Biswas and Biswas, 2010) 3.2.40 Family Theridiidae • Achaearanea durgae Tikader, 1970 (Biswas and Biswas, 2010) • Achaearanea sp. (Quasin and Uniyal, 2013) • Argyrodes argentatus Pickard-Cambridge, 1880 (Siddhu et al., 2020) • Argyrodes gazedes Tikader, 1970 (Uniyal et al., 2011) • Argyrodes sp. (Quasin and Uniyal, 2010, 2011; Uniyal et al., 2011; Pooja et al., 2019) • Chrysso nigriceps Keyserling, 1884 (Gupta and Siliwal, 2012) IAEES www.iaees.org 46 Arthropods, 2022, 11(1): 18-55 • Chrysso sp. (Uniyal et al., 2011; Gupta and Siliwal, 2012) • Dipoenura fimbriata Simon, 1909 (Gupta and Siliwal, 2012) • Enoplognatha sp. (Uniyal et al., 2011) • Episinus affinis Bösenberg and Strand, 1906 (Quasin et al., 2011; Uniyal et al., 2011) • Euryopis sp. (Uniyal et al., 2011; Pooja et al., 2019) • Molione triacantha Thorell, 1892 (Gupta and Siliwal, 2012) • Nihonhimea mundula (L. Koch, 1872) (Quasin and Uniyal, 2010; Pooja et al., 2019) • Parasteatoda sp. (Quasin and Uniyal, 2011a; Uniyal et al., 2011) • Phylloneta impressa (L. Koch, 1881) (Uniyal et al., 2011; Quasin and Uniyal, 2011b) • Phylloneta sp. (Uniyal et al., 2011) • Ruborridion musivum (Simon, 1873) (Quasin et al., 2017b) • Steatoda cingulata (Thorell, 1890) (Quasin et al., 2019) • Steatoda sp. (Uniyal et al., 2011) • Theridion subvittatum Simon, 1889 (Simon, 1889; Prasad et al., 2019) • Theridion sp. (Quasin and Uniyal, 2011a, 2013; Uniyal et al., 2011; Gupta and Siliwal, 2012; Pooja et al., 2019) • Thwaitesia margaritifera Pickard-Cambridge, 1881 (Gupta and Siliwal, 2012) 3.2.41 Family Thomisidae • Amyciaea forticeps (O.Pickard- Cambridge, 1873) (Gupta and Siliwal, 2012) • Bomis sp. (Gupta and Siliwal, 2012) • Camaricus formosus Thorell, 1887 (Biswas and Biswas, 2010; Gupta and Siliwal, 2012) • Camaricus sp. (Uniyal et al., 2011) • Diaea sp. (Quasin and Uniyal, 2010; Uniyal et al., 2011) • Henriksenia hilaris (Thorell, 1877) (Tikader, 1965; Biswas and Biswas, 2010; Uniyal et al., 2011) • Heriaeus horridus Tyschchenko, 1965 (Tikader, 1980) • Indoxysticus minutus (Tikader, 1960) (Uniyal et al., 2011; Gupta and Siliwal, 2012) • Lysiteles brunettii (Tikader, 1962) (Uniyal et al., 2011) • Lysiteles niger Ono, 1979 (Uniyal et al., 2011) • Lysiteles sp. (Quasin and Uniyal, 2010; Uniyal et al., 2011) • Massuria roonwali (Basu, 1964) (Basu, 1964; Tikader, 1971; Gupta and Siliwal, 2012) • Mastira menoka (Tikader, 1963) (Uniyal et al., 2011; Pooja et al., 2019) • Misumena mridulai Tikader, 1962 (Uniyal et al., 2011) • Misumena sp. (Quasin and Uniyal, 2011a, 2013; Uniyal et al., 2011; Gupta and Siliwal, 2012) • Misumenoides naginae Biswas and Roy, 2008 (Biswas and Roy, 2008) • Misumenops sp. (Uniyal et al., 2011) • Monaeses sp. (Gupta and Siliwal, 2012) • Oxytate elongata (Tikader, 1980) (Gupta and Siliwal, 2012) • Ozyptila sp. (Uniyal et al., 2011) • Pistius barchensis Basu, 1965 (Basu, 1965; Tikader, 1971) • Pistius bhadurii Basu, 1965 (Gupta and Siliwal, 2012) • Pistius gangulyi Basu, 1965 (Basu, 1965; Tikader, 1971) • Pistius kanikae Basu, 1964 (Basu, 1964; Tikader, 1971) • Pistius robustus Basu, 1965 (Basu, 1965; Tikader, 1971) • Runcinia insecta (L. Koch, 1875) (Gupta and Siliwal, 2012) IAEES www.iaees.org Arthropods, 2022, 11(1): 18-55 47 • Runcinia sp. (Quasin and Uniyal, 2013; Uniyal and Hore, 2006; Uniyal et al., 2011) • Synema decoratum Tikader, 1960 (Uniyal et al., 2011) • Thomisus dentiger (Thorell, 1887) (Leardi in Airaghi, 1901) • Thomisus lobosus Tikader, 1965 (Gupta and Siliwal, 2012; Pooja et al., 2019) • Thomisus onustus Walckenaer, 1805 (Uniyal et al., 2011) • Thomisus projectus Tikader, 1960 (Biswas and Biswas, 2010) • Thomisus sp. (Uniyal and Hore, 2006; Quasin and Uniyal, 2010; Pooja et al., 2019) • Xysticus croceus Fox, 1937 (Quasin and Uniyal, 2010; Uniyal et al., 2011) • Xysticus jaharai Basu, 1979 (Basu, 1979) • Xysticus joyantius Tikader, 1966 (Uniyal et al., 2011) • Xysticus kali Tikader and Biswas, 1974 (Uniyal et al., 2011) • Xysticus shyamrupus Tikader, 1966 (Biswas and Biswas, 2010) • Xysticus sp. (Quasin and Uniyal, 2011a, 2013; Uniyal et al., 2011; Gupta and Siliwal, 2012) 3.2.42 Family Trachelidae • Trachelas chamoli Quasin, Siliwal and Uniyal, 2018 (Quasin et al., 2018) • Trachelas sp. (Uniyal et al., 2011) 3.2.43 Family Trochanteriidae • Plator himalayaensis Tikader and Gajbe, 1976 (Tikader and Gajbe, 1976b; Sankaran et al., 2020b) • Plator indicus Simon, 1897 (Pocock, 1899, 1900; Uniyal et al., 2011) • Plator pandeae Tikader, 1969 (Tikader, 1969b) 3.2.44 Family Uloboridae • Hyptiotes sp. (Uniyal et al., 2011) • Miagrammopes extensus Simon, 1889 (Simon, 1889) • Miagrammopes sp. (Uniyal et al., 2011; Gupta and Siliwal, 2012) • Uloborus danolius Tikader, 1969 (Biswas and Biswas, 2010; Gupta and Siliwal, 2012) • Uloborus krishnae Tikader, 1970 (Uniyal et al., 2011; Pooja et al., 2019) • Uloborus sp. (Quasin and Uniyal, 2010, 2011a; Uniyal et al., 2011; Gupta and Siliwal, 2012) • Zosis geniculata (Olivier, 1789) (Leardi in Airaghi, 1901; Quasin and Uniyal, 2010; Uniyal et al., 2011; Pooja et al., 2019; Siddhu et al., 2020) 3.2.45 Family Zodariidae • Hermippus sp. (Gupta and Siliwal, 2012) • Lutica sp. (Quasin & Uniyal, 2013) • Zodarion sp. (Uniyal et al., 2011) Acknowledgements We are grateful of Dr. J.T.B. Caleb, Zoological Survey of India, Kolkata for providing few relevant literatures. References Agrawal N, Srivastava M, Tripathi A, Singh A. 2010. Survey and monitoring of pests, parasites and predators of pulse crops in central and eastern Uttar Pradesh. The Journal of Plant Protection Sciences, 1: 45-52 IAEES www.iaees.org 48 Arthropods, 2022, 11(1): 18-55 Anjali, Prakash S. 2012. Diversity of spiders (Araneae) from semi arid habitat of Agra (India). Indian Journal of Arachnology, 1(2): 66-72 Anjali, Prakash S. 2019. Some adaptive pattern of behaviour in spiders of semi-arid regions. Journal of Entomology and Zoology Studies, 7(2): 1118-1122 Anjali, Jindal V, Prakash S. 2019. Species richness and diversity of spiders in the semiarid habitats of north India. Indian Journal of Entomology, 81(4): 783-787 Baehr BC, Ubick D. 2010. A review of the Asian goblin spider genus Camptoscaphiella (Araneae: Oonopidae). American Museum Novitates, 3697: 1-65 Bastawade DB, Borkar M. 2008. Arachnida (orders Scorpiones, Uropygi, Amblypygi, Araneae and Phalangida). In: State Fauna Series-16. Fauna of Goa (Ed. Director), Zoological Survey of India, Kolkata, 211-242 Basu BD. 1964. Diagnosis of two new species of Pistius (Thomisidae: Araneae: Arachnida) from India. Journal of the Bengal Natural History Society, 32: 104-109 Basu, BD. 1965. Four new species of the spider genus Pistius Simon (Arachnida: Araneae: Thomisidae) from India. Proceedings of the Zoological Society, Calcutta, 18: 71-77 Basu KC. 1979. On a new spider of the genus Xysticus Koch, 1835 (Thomisidae: Arachnida) from Nainital, India. Journal of the Zoological Society of India, 28: 149-150 Bayer S. 2012. The lace-sheet-weavers-a long story (Araneae: Psechridae: Psechrus). Zootaxa, 3379: 1-170 Biswas B, Biswas K. 1992. Araneae: Spiders. State Fauna Series 3: Fauna of West Bengal, 3: 357-500 Biswas B, Biswas K. 2006. Araneae: Spiders.In: Fauna of Arunachal Pradesh, State Fauna Series. Zoological Survey of India, Kolkata, 13(2): 491-518 Biswas B, Biswas K. 2010. Araneae: Spider. In. Fauna of Uttarakhand, State Fauna Series, 18 (Part-3) (The Director, ed). 243-282, Zoological Survey of India, Kolkata, India Biswas B, Roy R. 2008. Description of six new species of spiders of the genera Lathys (Family: Dictynidae), Marpissa (Family: Salticidae), Misumenoides (Family: Thomisidae), Agroeca (Family: Clubionidae), Gnaphosa (Family: Gnaphosidae) and Flanona (Family: Lycosidae) from India. Records of the Zoological Survey of India, 108: 43-57 Blackwall J. 1867. Descriptions of several species of East Indian spiders, apparently to be new or little known to arachnologists. Annals and Magazine of Natural History, 19: 387-394 Bourne JD. 1980. New armored spiders of the family Tetrablemmidae from New Ireland and northern India (Araneae). Revue Suisse de Zoologie, 87: 301-317 Caleb JTD, Acharya S. 2019. First record of the genus Schenkelia Lessert, 1927 (Araneae: Salticidae) from India. Acta Arachnologica, 68(2): 73-75 Caleb JTD, Sankaran PM. 2021. Araneae of India, version 2021. https://indianspiders.in/. Accessed on October 10 2021 Caleb JTD, Sanap RV, Patel KG, Sudhin PP, Nafin KS, Sudhikumar AV. 2018a. First description of the female of Chrysilla volupe (Karsch, 1879) (Araneae: Salticidae: Chrysillini) from India, with notes on the species’ distribution and life history. Arthropoda Selecta, 27(2): 143-153 Caleb JTD, Sajan SK, Kumar V. 2018b. New jumping spiders from the alpine meadows of the Valley of Flowers, western Himalayas, India (Araneae, Salticidae). ZooKeys, 783: 113-124 Chandra K, Bharti D, Kumar S, Raghunathan C, Gupta D, Alfred JRB, Chowdhury BR. 2021. Faunal Diversity in Ramsar Wetlands of India. Zoological Survey of India, Kolkata, India Chandra U, Singh IB, Singh HM. 2017. Studies on Population dynamics of spider in rice crop regarding biocontrol. International Journal of Current Microbiology and Applied Sciences, Special Issue, 4: 116-124 IAEES www.iaees.org Arthropods, 2022, 11(1): 18-55 49 Chaubey SN. 2017a. Studies on habit and habitat, external morphology, feeding capacity and prey preference of true worb-weaving spider, Argiope aemula (Walckenaer). Indian Journal of Scientific Research, 15(1): 30-34 Chaubey SN. 2017b. Studies on habit and habitat, external morphology, feeding capacity and prey preference of jumping spider Phidippus audax (Koch). Journal of Applied Bioscience, 43(2): 84-89 Chaubey SN. 2017c. Studies on habit and habitat, external morphology, feeding capacity and prey preference of zebra jumper Plexippus petersi (Karsch). Indian Journal of Scientific Research, 15(1): 64-68 Chaubey SN. 2017d. Studies on habit, habitat, external morphology, feeding capcity and prey preference of worb-weaving spider, Guizygiella melanocrania. Flora and Fauna, 23(2): 439-444 Chaubey SN. 2019a. Studies on habit and habitat, external morphology, feeding capacity and prey preference of a lynx spider, Oxyopes quadrifasciatus. Journal of Applied Bioscience, 45(1-2): 24-28 Chaubey SN. 2019b. Studies on habit and habitat, external morphology, feeding capacity and prey preference of ant spider, Myrmarachne orientalis (Tikader). Indian Journal of Scientific Research, 10(1): 179-182 Chaubey SN. 2019c. Studies on habit and habitat, external morphology, feeding capacity and prey preference of garden jumping spider, Opisthoncus species. Indian Journal of Scientific research, 10(1): 29-33 Chaubey SN. 2019d. Studies on habit and habitat, external morphology, feeding capacity and prey preference of two striped spider, Telamonia dimidiata. Flora & Fauna, 25(2): 204-208 Chaubey SN, Mishra RS. 2016. Habit and habitat, morphology, feeding capacity and prey preference of six humped dome spider, Cyrtophora cicatrosa Stoliczka. Journal of Applied Bioscience, 42(2): 109-113 Chaubey SN, Mishra RS. 2017a. Studies on the habit and habitat, morphology, feeding capacity and prey preference of six humped dome spider Cyrtophora citricola Simon. Indian Journal of Scientific Research, 7(2): 39-45 Chaubey SN, Mishra RS. 2017b. Study on the morphology, feeding capacity and prey preference of long jawed spider, Eucta chamberlini (Simon). Journal of Experimental Zoology, India, 20(1): 61-65 Chaubey SN, Yadav PR. 2017a. Studies on habit and habitat, external morphology, feeding capacity and prey preference of jumping spider Menemerus semilimbatus (Simon). Journal of Experimental Zoology, India, 20(2): 729-732 Chaubey SN, Yadav PR. 2017b. Studies on habit and habitat, external morphology, feeding capacity and prey preference of zebra jumper spider Plexippus paykuli (Audeuin) Journal of Experimental Zoology India, 20(2): 901-905 Chaubey SN, Mishra RS, Yadav PR. 2019a. Studies on habit and habitat, external morphology, feeding capacity and prey preference of gray wall jumper spider, Menemerus bivittatus (Dufour). Journal of Experimental Zoology, India, 22(2): 813-816 Chaubey SN, Mishra RS, Yadav PR. 2019b. Studies on habit and habitat, external morphology, feeding capacity and prey preference of Zenodorus (Peckham and Peckham) spider species. Journal of Experimental Zoology, India, 22(2): 809-812 Crews SC, Harvey MS. 2011. The spider family Selenopidae (Arachnida, Araneae) in Australasia and the Oriental region. ZooKeys, 99: 1-103 Diksha Khan RA, Sultana A, Das SK. 2018. A new spider record of genus Thomisus Walckenaer, 1805 (Araneae: Thomisidae) from India. Serket, 16(2): 96-99 Fage L. 1946. Araignées cavernicoles de l'Inde. Bulletin du Muséum National d'Histoire Naturelle de Paris, 18: 382-388 Gajbe UA. 1979. Studies on some spiders of the genus Sosticus from India (Araneae: Gnaphosidae). Bulletin of the Zoological Survey of India, 2: 69-74 IAEES www.iaees.org 50 Arthropods, 2022, 11(1): 18-55 Gajbe UA. 1983. On three new species of spiders of the genus Callilepis Westring (Family: Gnaphosidae) from India. Records of the Zoological Survey of India, 81: 127-133 Gajbe UA. 1988. On a collection of spiders of the family Gnaphosidae from India (Araneae: Arachnida). Records of Zoological Survey of India, 85(1): 59-74 Gajbe UA. 1992a. A new species of Oxyopes Latreille and one of Peucetia Thorell from Uttar Pradesh, India (Araneae, family: Oxyopidae). Records of the Zoological Survey of India, 91: 389-393 Gajbe UA. 1992b. On two new species of Haplodrassus spiders from India (Araneae: Gnaphosidae). Records of the Zoological Survey of India, 91: 313-317 Gajbe UA. 1999. Studies on some spiders of the family Oxyopidae (Araneae: Arachnida) from India. Records of the Zoological Survey of India, 97(3): 31-79 Gajbe UA. 2005. Studies on some spiders of the family Gnaphosidae (Araneae: Arachnida) from Madhya Pradesh, India. Records of the Zoological Survey of India, 105(3-4): 111-140 Gajbe UA. 2007. Araneae: Arachnida. In: State Fauna Series, 15 Part 1, Fauna of Madhya Pradesh (including Chhattisgarh), (Ed. Director). 419-540, Zoological Survey of India, Kolkata, India Gajbe UA. 2008. Fauna of India and the adjacent countries Vol. 3. Spider (Arachnida: Araneae: Oxyopidae). Zoological Survey of India, Kolkata, India Gravely FH. 1921. Some Indian spiders of the subfamily Tetragnathinae. Records of the Indian Museum, Calcutta, 22: 423-459 Gravely FH. 1924. Some Indian spiders of the family Lycosidae. Records of the Indian Museum, Calcutta, 26: 587-613 Gravely FH. 1931. Some Indian spiders of the families Ctenidae, Sparassidae, Selenopidae and Clubionidae. Records of the Indian Museum, Calcutta, 33: 211-282 Grismado CJ, Deeleman-Reinhold CL, Piacentini LN, Izquierdo MA, Ramírez MJ. 2014. Taxonomic review of the goblin spiders of the genus Dysderoides Fage and their Himalayan relatives of the genera Trilacuna Tong and Li and Himalayana, new genus (Araneae, Oonopidae). Bulletin of the American Museum of Natural History, 387: 1-108 Gupta N, Siliwal M. 2012. A checklist of spiders (Arachnida: Araneae) of Wildlife Institute of India campus, Dehradun, Uttarakhand, India. Indian Journal of Arachnology, 1(2): 73-91 Halder J, Rai AB, Kodandaram MH, Shivalingaswamy TM, Dey D. 2012. Mud wasp, Sceliphron madraspatanum (Fabricius) (Hymenoptera: Sphecidae): A threat or nature’s regulation of spider fauna in the vegetable agroecosystem. Journal of Biological Control, 26: 373-375 Hore U, Uniyal VP. 2008a. Effect of prescribed fire on spider assemblage in Terai grasslands, India. Turkish Journal of Arachnology, 1(1): 15-36 Hore U, Uniyal VP. 2008b. Diversity and composition of spider assemblages in five vegetation types of the Terai Conservation Area, India. The Journal of Arachnology, 36(2): 251-258 Hore U, Uniyal VP. 2008c. Use of spiders (Araneae) as indicator for monitoring of habitat conditions in Tarai Conservation Area, India. Indian Forester, 134: 1371-1380 Huber BA. 2011. Revision and cladistic analysis of Pholcus and closely related taxa (Araneae, Pholcidae). Bonner Zoologische Monographien, 58: 1-509 Jäger P. 2001. Diversität der Riesenkrabbenspinnen im Himalaya — die Radiation zweier Gattungen in den Schneetropen (Araneae, Sparassidae, Heteropodinae). Courier Forschungsinstitut Senckenberg, 232: 1-136 Jäger P. 2012. Asian species of the genera Anahita Karsch 1879, Ctenus Walckenaer 1805 and Amauropelma Raven, Stumkat & Gray 2001 (Arachnida: Araneae: Ctenidae). Zootaxa, 3429: 1-63 IAEES www.iaees.org Arthropods, 2022, 11(1): 18-55 51 Jithin V, Devarajan A, Dharmaraj J. 2021. Predation on Asian common toad, Duttaphrynus melanostictus (Lütken, 1864) (Anura: Bufonidae), tadpoles by a fish-eating spider, Nilus sp. (Araneae: Pisauridae). Reptiles & Amphibians, 28(1): 56-58 Khan AA, Misra DS. 2003. Studies on qualitative and quantitative composition of spider fauna in rice ecosystem of eastern Uttar Pradesh. Plant Protection Bulletin, 55(1-2): 35-41 Kumar, A, Kanaujia A, Kumar A, Kumar V, Mishra H. 2017a. Diversity of spiders in Kukrail Reserve Forest, Lucknow, Uttar Pradesh, India. Journal of Environmental Science and Technology, 4(5): 42-45 Kumar A, Kanaujia A, Kumar A, Kumar V, Mishra H. 2017b. Araneofauna of Nawabganj bird sanctuary, Unnao, Uttar Pradesh, India. Journal of Entomology and Zoology Studies, 5(4): 1952-1955 Lawania KK, Mathur P. 2014a. Baseline studies on the spider fauna (Araneae) of Braj region (Braj-Bhoomi), India. International Journal of Basic and Applies Biology, 2(1): 137-141 Lawania KK, Mathur P. 2014b. Diversity and distribution of spider fauna in and around the Taj Mahal and Taj Protected Forest, Agra (U.P.), India. International Journal of Basic and Applied Biology, 2(2): 111-114 Lawania KK, Mathur P. 2014c. On the diversity of spiders in and around Sur-Sarovar Bird Sanctuary, Agra (U.P.), India. International Journal of Basic and Applied Biology, 2(3): 189-194 Lawania KK, Mathur P. 2014d. Diversity and distribution of spiders in and around Vrindavan, Mathura, (UP), India. International Journal of Basic and Applied Biology, 2(2): 115-119 Leardi in Airaghi Z. 1901. Aracnidi d'Almora. Atti della Societa Italiana di Scienze Naturali e del Museo Civico di Storia Naturale di Milano, 40: 85-94 Levi, HW. 1982. The spider genera Psechrus and Fecenia (Araneae: Psechridae). Pacific Insects, 24: 114-138 Lin YC, Ballarin F, Li SQ. 2016. A survey of the spider family Nesticidae (Arachnida, Araneae) in Asia and Madagascar, with the description of forty-three new species. ZooKeys, 627: 1-168 Lin YJ, Marusik YM, Gao CX, Xu H, Zhang XQ, Wang ZY, Zhu WH, Li SQ. 2021. Twenty-three new spider species (Arachnida: Araneae) from Asia. Zoological Systematics, 46(2): 91-152 Logunov DV, Hereward J. 2006. New species and synonymies in the genus Synagelides Strand in Bösenberg & Strand, 1906 (Araneae: Salticidae). Bulletin of the British Arachnological Society, 13: 281-292 Logunov DV. 2019. Taxonomic notes on the Harmochirina Simon, 1903 from South and South-East Asia (Aranei: Salticidae). Arthropoda Selecta, 28(1): 99-112 Logunov DV. 2021. New species and records of the jumping spiders from India and Nepal (Aranei: Salticidae). Arthropoda Selecta, 30(3): 351-361 Majumder SC, Tikader BK. 1991. Studies on some spiders of the family Clubionidae from India. Records of the Zoological Survey of India, Occasional Paper, 102: 1-175 Marusik YM, Zheng G, Li SQ. 2008. A review of the genus Paratus Simon (Araneae, Dionycha). Zootaxa, 1965: 50-60 Marusik YM, Ballarin F, Omelko MM. 2012. On the spider genus Amaurobius (Araneae, Amaurobiidae) in India and Nepal. ZooKeys, 168: 55-64 Mishra A, Rastogi N. 2020. Unraveling the roles of solitary and social web-making spiders in perennial ecosystems: influence on pests and beneficials. Proceedings of the National Academy of Sciences, India, Section B: Biological Sciences, 90: 567-576 Mishra A, Kumar B, Rastogi N. 2021. Predation potential of hunting and web-building spiders on rice pests of Indian subcontinent. International Journal of Tropical Insect Science, 41: 1027-1036 Mishra R, Ahmad G, Chaubey SN. 2012a. Study on the morphology, feeding capcityand prey preference of Neoscona crucifera and N. adianta (orb-weaving spiders). Indian Journal of Life Sciences, l(2): 29-34 IAEES www.iaees.org 52 Arthropods, 2022, 11(1): 18-55 Mishra R, Chaubey SN, Ahmad G. 2012b. Study on the morphology, feeding capacity and prey preference of orb-weaving spider Neoscona nautica (L. Koch, 1875). Journal of Experimental Zoology, India, 15(2): 467-472 Nigam S, Patel SK. 1996. A new species of spider of the genus Eilica (Gnaphosidae), a predator from Uttar Pradesh, India. Geobios new Reports, 15: 2-4 Patel SK, Nigam S. 1994. New spider species (Araneae: Araneidae) from Uttar Pradesh, India. Journal of Ecobiology, 6: 201-205 Platnick NI, Dupérré N, Ott R, Baehr BC, Kranz-Baltensperger Y. 2012. The goblin spider genus Pelicinus (Araneae, Oonopidae), Part 1. American Museum Novitates, 3741: 1-43 Pocock RI. 1899. Diagnoses of some new Indian Arachnida. Journal of the Bombay Natural History Society, 12: 744-753 Pocock, RI. 1900. The Fauna of British India including Ceylon and Burma Arachnida. Taylor and Francis London, UK Pocock RI. 1901. Descriptions of some new species of spiders from British India. Journal of the Bombay Natural History Society, 13: 478-498 Pooja A, Anilkumar, Quasin S, Lekshmi S, Uniyal VP. 2019. Spider Fauna of Navdanya Biodiversity Farm, Uttarakhand, India. Indian Forester, 145(4): 392-397 Prasad P, Tyagi K, Caleb JTD, Kumar V. 2019. A new species of the cob web spider genus Theridion from India (Araneae: Theridiidae). Ecologica Montenegrina, 26: 108-117 Prószyński J. 1992. Salticidae (Araneae) of the Old World and Pacific Islands in several US collections. Annales Zoologici, Warszawa, 44: 87-163 Quasin S, Uniyal VP. 2010. Preliminary Investigation of spider diversity in Kedarnath Wildlife sanctuary, Uttarakhand, India. Indian Forester, 136(10): 1340-1345 Quasin S, Uniyal VP. 2011a. Spider diversity along altitudinal gradient in Milam Valley, Nanda Devi Biosphere Reserve, Western Himalaya. Indian Forester, 137(10): 1207-1211 Quasin S, Uniyal VP. 2011b. First record of the genus Phylloneta from India with description of P. impressa L. Koch, 1881 (Araneae: Theridiidae). Biosystematica, 5: 59-61 Quasin S, Uniyal VP. 2013. Spiders (Araneae) in Bhyundar valley of Nanda Devi Biosphere Reserve. In: Biodiversity Communities and Climate Change (Kala CP, C.S. Silori CS, eds). 13-24, The Energy and Resource Institute, TERI, New Delhi, India Quasin S, Uniyal VP, Jose K.S. 2011. First report of Episinus affinis (Araneae: Theridiidae) from India. Records of the Zoological Survey of India, 111(4): 97-98 Quasin S, Siliwal M, Uniyal VP. 2015. New species of Himalmartensus Wang & Zhu, 2008 (Araneae: Amaurobiidae) with the first description of a male from the Nanda Devi Biosphere Reserve, Western Himalaya, India. Journal of Asia-Pacific Biodiversity, 8: 247-250 Quasin S, Siliwal M, Uniyal VP. 2017a. First report of the genus Draconarius Ovtchinnikov, 1999 (Araneae: Agelenidae: Coelotinae) with description of a new species from India. European Journal of Zoological Research, 5(1): 19-22 Quasin S, Siliwal M, Patil V, Uniyal VP. 2017b. First record of Ruborridion musivum Simon, 1873 (Araneae: Theridiidae) from India. Munis Entomology and Zoology, 12(1): 27-30 Quasin S, Siliwal M, Uniyal VP. 2018. New species of Trachelas (Araneae: Trachelidae) from Nanda Devi Biosphere Reserve-Western Himalaya, India. Journal of Asia-Pacific Biodiversity, 11(1): 158-160 Quasin S, Siliwal M, Uniyal VP. 2019. First report of Steatoda cingulata (Thorell, 1890) (Araneae: Theridiidae) from Nanda Devi Biosphere Reserve, Western Himalaya. Munis Entomology and IAEES www.iaees.org Arthropods, 2022, 11(1): 18-55 53 Zoology, 14(2): 638-642 Rajpoot A, Kumar VP, Bahuguna A, Maity P, Kumar D. 2018. Araniella cucurbitina: the first molecular evidence of a Palearctic species of genus Araniella inhabiting India. Mitochondrial DNA Part A, 29(6): 831-839 Sankaran PM, Sebastian PA. 2016. A checklist of Indian armored spiders (Araneae, Tetrablemmidae) with the description of a new species from the Western Ghats. Zootaxa, 4084(3): 443-450 Sankaran PM, Caleb JTD, Sebastian PA. 2020a. Taxonomic notes on the genus Makdiops Crews & Harvey, 2011 in India (Araneae: Selenopidae. Zootaxa, 4896(4): 595-600 Sankaran PM, Caleb JTD, Joseph MM, Sebastian PA. 2020b. On a new synonymy in the spider genus Nephila Leach, 1815 (Araneidae, Nephilinae) from India with supplementary notes on colour polymorphism in the genus. Zootaxa, 4786(4): 592-596 Sethi VD, Tikader BK. 1988. Studies on some giant crab spiders of the family Heteropodidae from India. Records of Zoological Survey of India, Miscellaneous Publications, 93: 1-9 Sharma A, Singh R. 2018a. Biodiversity and guild structure of spiders in northeastern Uttar Pradesh. Journal of Life Sciences, Bioinformatics, Pharmaceuticals & Chemical Sciences, 4(4): 525-541 Sharma A, Singh R. 2018b. Species diversity and guild structure of spiders from Siddharthnagar, Uttar Pradesh, India. Research Journal of Life Sciences, Bioinformatics, Pharmaceuticals and Chemical Sciences, 4(4): 383-390 Sharma A, Singh G, Singh R. 2020. Faunal diversity of Liocranidae, Mimetidae, Miturgidae, Nesticidae and Oecobiidae (Arachnida: Araneae) of India. Serket, 17(3): 270-283 Sherriffs WR. 1931. South Indian Arachnology. Part V. Annals and Magazine of Natural History, 10: 537-546 Sherriffs, WR. 1951. Some oriental spiders of the genus Oxyopes. Proceedings of the Zoological Society of London, 120: 651-677 Siddhu J, Lohani HP, Pathak G, Kaushal BR. 2020. Spider diversity in rice and mix vegetable agro bhabar region of Nainital district, Uttarakhand. Bulletin of Environment, Pharmacology and Life Sciences, 9(2): 101-105 Siliwal M, Molur S, Raven R. 2011. Mygalomorphs of India: An overview. ENVIS Bulletin: Arthropods and their Conservation in India, 14(1): 175-188 Siliwal, M, Kumar RS, Raven R. 2014. A new species of Atypus Latreille, 1804 (Araneae: Atypidae) from Northern India. Arthropoda Selecta, 23(2): 221-224 Siliwal M, Hippargi R, Yadav A, Kumar D. 2020. Five new species of trap-door spiders (Araneae: Mygalomorphae: Idiopidae) from India. Journal of Threatened Taxa, 12(13): 16775-16794 Simon E. 1889. Arachnides de l'Himalaya, recueillis par MM. Oldham et Wood-Mason, et faisant partie des collections de l'Indian Museum. Première partie. Journal of the Asiatic Society of Bengal, part II (Natural Science), 58: 334-344 Simon E. 1897. Matériaux pour servir à la faune arachnologique de l'Asie méridionale. V. Arachnides recueillis à Dehra-Dun (N.W. Prov.) et dans le Dekkan par M. A. Smythies. Mémoires de la Société Zoologique de France, 10: 252-262 Simon E. 1900. Etudes arachnologiques. 30e Mémoire. XLVII. Descriptions d'espèces nouvelles de la famille des Attidae. Annales de la Société Entomologique de France, 69: 27-61 Simon E. 1906. Etude sur les araignées de la section des cribellates. Annales de la Société Entomologique de Belgique, 50: 284-308 IAEES www.iaees.org 54 Arthropods, 2022, 11(1): 18-55 Singh A, Devi J, Tripathi S, Singh S. 2021. Diversity assessment of spiders in Parvati Aranga Bird Sanctury district Gonda, Uttar Pradesh, India. International Research Journal of Modernization in Engineering Technology and Science, 3(10): 508-515 Singh BB, Singh R. 2014. Incidence and biodiversity of riceland spiders (Arthropoda: Arachnida) in northeastern Uttar Pradesh, India. Indo-American Journal of Life Sciences & Biotechnology, 2(1): 64-89 Singh R, Singh G. 2021a. Faunistic diversity of orb-weaver spiders (Araneidae: Araneomorphae: Araneae: Arachnida) in India. International Journal of Biological and Environmental Investigations, 1(2): 62-133 Singh R, Singh G. 2021b. An updated checklist of spiders (Arachnida: Araneae) in Northeast India. Serket, 18(1): 91-144 Singh R, Singh G. 2021c. Faunal diversity of spiders (Chelicerata: Araneae) in Bihar and Jharkhand, India. International Journal of Biological Innovations, 3(2): 382-391 Singh R, Singh G. 2021d. Updated checklist of spider (Arachnida: Araneae) diversity in Haryana, Himachal Pradesh, Punjab, Chandigarh and Delhi (India). Serket, 18(2) Singh RS, Parasnath & Kumar A. 2013. Effect of sustainable pest management approaches on the population dynamics of spider fauna inhabiting pigeon pea agro-ecosystem. Agricultural Science Digest, 33(1): 63-67 Strand E. 1907. Vorläufige Diagnosen süd- und ostasiatischer Clubioniden, Ageleniden, Pisauriden, Lycosiden, Oxyopiden und Salticiden. Zoologischer Anzeiger, 31: 558-570 Strand E. 1909. Süd- und ostasiatische Spinnen. II. Fam. Clubionidae. Fam. Salticidae. Abhandlungen der Naturforschenden Gesellschaft Görlitz, 26: 1-128 Sujayanand GK, Chandra A, Pandey S, Bhatt S. 2021. Seasonal abundance of spotted pod borer, Maruca vitrata Fabricius in early pigeonpea [Cajanus cajan (L.) Millsp.] and its management through farmscaping in Uttar Pradesh. Legume Research, 44(2): 233-239 Tanasevitch AV. 2011. Linyphiid spiders (Araneae, Linyphiidae) from Pakistan and India. Revue Suisse de Zoologie, 118: 561-598 Tanasevitch AV. 2017. New species and new records of linyphiid spiders from the Indo-Malayan Region (Araneae, Linyphiidae). Zootaxa, 4227 (3): 325-346 Tandon PL, Lal B. 1983. Predatory spiders associated with insect pests of mango in India. Bulletin of Entomology, 24: 144-147 Tikader BK. 1965. On some new species of spiders of the family Thomisidae from India. Proceedings of the Indian Academy of Science, 61(B): 277-289. Tikader BK. 1966a. On a collection of spiders (Araneae) from the desert areas of Rajasthan (India). Records of the Indian Museum, Calcutta, 59: 435-443 Tikader BK. 1966b. On some new species of spiders of the genus Philodromus Walck. (family: Thomisidae) from India. Proceedings of the Linnean Society of London, 177(1): 35-44 Tikader BK. 1969a. Studies on some spiders of the family Oxyopidae from India. Oriental Insects, 3: 33-36. Tikader BK. 1969b. Studies of some rare spiders of the families Selenopidae and Platoridae from India. Proceedings of the Indian Academy of Science, 69(B): 252-255 Tikader BK. 1971. Revision of Indian crab spiders (Araneae: Thomisidae). Memoirs of the Zoological Survey of India, 15(8): 1-90 Tikader BK. 1980. Araneae: Thomisidae (Crab-spiders). In: Fauna of India Vol. 1, Part 1 (Director, ed). Zoological Survey of India, Kolkata, India Tikader BK. 1982. Spiders: Araneae. In: The Fauna of India, Vol. 2, Part 1 Family Araneidae (=Argiopidae) Typical Orb-Weavers (Director ed). Zoological Survey of India, Kolkata, India IAEES www.iaees.org Arthropods, 2022, 11(1): 18-55 55 Tikader BK, Gajbe, UA. 1975. New species of Drassodes spiders (Araneae: Gnaphosidae) from India. Oriental Insects, 9: 273-281 Tikader BK, Gajbe, UA. 1976a. Studies on some spiders of the genus Zelotes Gistel from India (family: Gnaphosidae). Proceedings of the Indian Academy of Science, 83(B): 109-122 Tikader BK, Gajbe UA. 1976b. A new species of spider of the genus Plator Simon (family: Platoridae) from India. Journal of the Bombay Natural History Society, 72: 797-799 Tikader BK, Gajbe UA. 1977a. Studies on some spiders of the genera Gnaphosa Latreille and Callilepis Westring (family: Gnaphosidae) from India. Records of the Zoological Survey of India, 73: 43-52 Tikader BK Gajbe UA. 1977b. Taxonomic studies on some spiders of the genera Drassodes Westring, Haplodrassus Chamberlin, Geodrassus Chamberlin and Nodocion Chamberlin (family: Gnaphosidae) from India. Records of the Zoological Survey of India, 73: 63-76 Tikader BK, Gajbe UA. 1977c. Studies on some spiders of the genera Scopodes Chamberlin, Megamyrmecion Reuss, Scotophaeus Simon and Liodrassus Chamberlin (family: Gnaphosidae) from India. Records of the Zoological Survey of India, 73: 13-22 Tikader BK, Malhotra MS. 1976. Studies on some spiders of the genus Pardosa Koch from India (family: Lycosidae). Proceedings of the Indian Academy of Science, 83(3): 123-131 Tikader BK, Malhotra MS. 1980. Araneae: Lycosidae (Wolf-spiders). In: The Fauna of India, Volume 1, Part 2 (Director, ed). Zoological Survey of India, Kolkata, 248-447 Tikader BK, Malhotra MS. 1981. Revision of spiders of the genus Ctenus Walckenaer from India (Araneae: Ctenidae). Records of the Zoological Survey of India, 79: 105-124 Tikader BK, Sethi VD. 1990. Studies of some giant crab spiders of the family Heteropodidae from India. Part II. Records of the Zoological Survey of India, 87: 165-186 Uniyal VP, Hore U. 2006. Studies on the spider fauna in mixed sal forest area of Chandrabani, Dehradun. Indian Forester, 132(12a): 83-88 Uniyal VP, Hore U. 2009. Effect of Management Practices on Spider Diversity in Terai Conservation Area (TCA). Final Project Report, Wild Life Institute of India, 222. Uniyal VP, Sivakumar K, Quasin S. 2011. Diversity of Spiders in Nanda Devi Biosphere Reserve. Wildlife Institute of India, Dehradun. DST Project Completion Report, 199 Wanless FR. 1978. A revision of the spider genus Portia (Araneae: Salticidae). Bulletin of the British Museum of Natural History (Zoology), 34: 83-124 WSC. 2021. World Spider Catalog. Version 22.5. Natural History Museum Bern. http://wsc.nmbe.ch. Accessed on November 27, 2021 Yadav A, Chaubey SN, Beg MA. 2012a. Hippasa holmerae Thorell (garden wolf spider) as biocontrol agent for insect pests of crop fields collected from Azamgarh and Mau districts (U.P.) India. Journal of Experimental Zoology, India, 15(2): 495-498 Yadav A, Chaubey SN, Beg MA. 2012b. Morphology, prey preference and feeding capacity of decorative spider, Leucauge decorata (Blackwall) from Azamgarh, India. Journal of Applied Bioscience, 38(1): 63-67 Yadav H, Prakash S. 2021. Diversity of spiders (Araneae) in the flood plains of the Taj trapezium zone of Agra. Applied Ecology and Environmental Sciences, 9(2): 149-155 Yadav RS. 2018. First report of Nephila clavata L. Koch, 1878 (Araneae: Araneidae) from Bihar and Uttar Pradesh, India. Journal of Entomology and Zoology Studies, 6(1): 754-756 Zamani A, Marusik YM. 2020. Two new species of Araniella (Aranei: Araneidae) from Western Himalaya, with notes on species reported from India. Arthropoda Selecta, 29(3): 361-366 IAEES www.iaees.org Arthropods, 2022, 11(1): 56-64 Article Dragonflies and damselflies (Odonata: Insecta) of the Seloo city, Wardha, Maharashtra, Central India Ashish Tiple, Rahul Bhende, Parag Dandge PG Department of Zoology, Vidyabharti College, Seloo, Wardha 442 104, India E-mail: ashishdtiple@gmail.com Received 14 October 2021; Accepted 20 November 2021; Published 1 March 2022 Abstract Dragonflies and damselflies (Odonata) species diversity was studied in the Seloo city from 2011 to 2021. Its geographical location is 20083’73’’N; 78070’70’’E; 265 m. A total of 62 species of odonates belonging to 2 Suborders and 8 families were recorded. The highest number of odonates belong to the family Libellulidae (30 species) followed by Coenagrionidae (13 species), Aeshnidae (5 species), Gomphidae (4 species), Platycenemididae (3 species) and Lestidae (4 species), Macromiidae (2 species) and Chlorocyphidae (1 species). Of the total, 30 species were abundant or very common, 16 were common, 6 were not rare, 7 rare and 3 very rare. Among all, 3 species were Data Deficient, Indothemis carnatica (Fabricius, 1798) are listed as Near Threatened and 57 were least concern in IUCN red-list of threatened species. The observations support the value of the Seloo city area in providing valuable resources for Odonata. Keywords Odonata; diversity; Seloo city; Wardha,;Mahrashtra; India. Arthropods ISSN 2224­4255 URL: http://www.iaees.org/publications/journals/arthropods/online­version.asp RSS: http://www.iaees.org/publications/journals/arthropods/rss.xml E­mail: arthropods@iaees.org Editor­in­Chief: WenJun Zhang Publisher: International Academy of Ecology and Environmental Sciences 1 Introduction Odonata (damselflies and dragonflies) are very interesting and diverse insects. Odonata are paleopterous, exopterygote aquatic insects, probably more closely related to the Ephemeroptera (mayflies) than any other living insect group. They instantly attract attention with their amazing flight skills and beautiful colours. Odonate is prominent freshwater insects and plays an important role in wetland and terrestrial food chains as predators. The adults are generally predacious insects, while the larvae are carnivores and voracious feeders. They are also actively used in controlling causative agent of malaria and filaria throughout the world (Tiple et al., 2008). Even though species are usually highly specific to a habitat, some have adapted to urbanization and use man-made water bodies. They probably mark the first time that evolution experimented with the ability to hover in air over an object of interest. Being primarily aquatic, their life history is closely linked to specific aquatic habitats. Naturally, these insects become a marker, an indicator of wetland health (Andrew et al., 2008). Dragonflies mostly occur in the vicinity of different fresh water habitats like rivers, streams, marshes, lakes IAEES www.iaees.org Arthropods, 2022, 11(1): 56-64 57 and even small pools and rice fields. Odonates are also good indicators of environmental changes as they are sensitive and are directly affected by changes in the habitats, atmospheric temperature and the weather conditions (Dijkstra and Lewington, 2006). They are also Bio-control Agents, many species of odonates inhabiting in agro ecosystems play a crucial role controlling pest populations (Tiple et al., 2013). Globally 6335 species in 693 genera of odonates have been reported (Schorr and Paulson, 2021), of which 498 species, 27 Subspecies in 154 genera and 18 families are known from India (Subramanian and Babu, 2020; Joshi and Sawant, 2020; Bedjanič et al., 2020; Payra et al., 2020; Payra et al., 2021; Dawn, 2021). After Fraser’s seminal work on Odonata of India (Fraser, 1933, 1934, 1936), there was a gap of almost 50 years in Odonata studies across the country. After establishing the Zoological Survey of India (ZSI) in 1916, trained taxonomists started collecting data and publishing lists of Odonata of localities or regions. Researchers from the ZSI and various odonatologists from the academic institutes of India have often surveyed various parts of Maharashtra region. The odonata fauna of the State of Maharashtra is well-documented with 134 species (reviewed in Tiple and Koparde, 2015), but few spatial gaps still remain. The eastern part of the State of Maharashtra (Vidarbha) is home to 85 odonates (Tiple et al., 2013; Tiple et al., 2014; Talmale and Tiple, 2013; Tiple, 2015, 2020). The present paper reports detailed survey on the dragonfly and damselfly (Odonata) diversity of Seloo city. No published checklist of Odonata species of Seloo city region is known, hence, the present work was initiated. 2 Materials and Methods 2.1 Study area The Seloo city (20083’73’’N; 78070’70’’E; 265 m) is situated nearer to Bor Wildlife Sanctuary at the bank of river Bor along with the dense shrub, natural vegetation and tree vegetation which are the major attraction to the Odonata. It has tropical wet and dry climate with dry conditions, an annual rainfall of about 1,205 mm (June to September); temperature raises up to 48.9°C during summer (March-June) and falls up to 10°C to 6.9°C in winter (November-January). Annual relative humidity varies in between 22% to 80% (Tiple, 2018). 2.2 Identification Odonates were photographed and identified in different regions of the Seloo city 2011 to 2021. Most of the sampling was done between 10 AM to 2 PM, when odonates are most active (Subramanian, 2009; Payra and Tiple, 2019). Odonates were surveyed in lakes, rivers, pond, temporary and permanent flowing or still water bodies and surrounding area, during the monsoon and post monsoon period. A weekly survey was undertaken during the monsoon (July–August) and post monsoon period at all sites. The adult odonates were identified with the help of identification keys provided by Fraser, 1933-1936 and Mitra, 1986. All nomenclature follows Subramanian and Babu (2017). The species were categorized on the basis of their abundance in Seloo city VC Very common (> 100 sightings), C Common (50-100 sightings), NR Not rare (15-50 sightings), R Rare (2-15 sightings), VR Very rare (< 2 sightings) (Tiple et al., 2008). 3 Results and Discussion A total of 62 species of odonates belonging to 2 Suborders and 08 families were recorded. The highest number of odonates belong to the family Libellulidae (30 species), followed by Coenagrionidae (13 species), Aeshnidae (5 species), Gomphidae (4 species), Platycenemididae (3 species) and Lestidae (4 species), Macromiidae (2 species) and (1 species) Chlorocyphidae (Fig. 1, 2). Of the total, 30 species were abundant or very common, 16 were common, 6 were not rare, 7 rare and 3 very rare. Among the 62 odonates recorded from Seloo city, 61 species come under the International Union of Conservation for Nature (IUCN) red-list of threatened species. Among them Indothemis carnatica come under IAEES www.iaees.org 58 Arthropods, 2022, 11(1): 56-64 Near Threatened. 57 species recorded which come under Least Concern category, the species recorded which come under Data Deficient category (i.e. Microgomphus torquatus, Lestes umbrinus, Elattoneura nigerrima) and 1 was not listed in IUCN red-list of threatened species. The family Gomphidae is also represented by the highest number of Data Deficient species as well as species for which information is not available in the IUCN red list of threatened species (Table 1). The members of this family are fast moving insects and may have crepuscular habits. These insects are difficult to observe or collect. Many Gomphidae are already rare. Therefore, there are high chances of not detecting them during surveys (Tiple and Khoparde, 2015). The list of odonates along with their scientific names and their occurrence status and Threat status is provided in Table 1. Fig. 1 The number of Odonates species encountered in different families in the Seloo city, Wardha, Maharashtra. IAEES www.iaees.org Arthropods, 2022, 11(1): 56-64 IAEES 59 www.iaees.org Arthropods, 2022, 11(1): 56-64 60 Fig. 2 Some recorded species of Odonata from Seloo city. Table 1 Checklist of Odonata of Seloo city. OS: Occurrence status; TS: Threat status as assigned from IUCN. NA: Not available; LC: Least concern; DD: Data deficient; VU: Vulnerable; NT: Near threatened. No. Scientific name OS TS Suborder: Anisoptera (Dragonflies) Family: Aeshnidae IAEES (05) 1. Anax guttatus (Burmeister, 1839) NR LC 2. Anax immaculifrons (Rambur, 1842) C LC 3. Anax indicus Lieftinck, 1942 VC LC www.iaees.org Arthropods, 2022, 11(1): 56-64 61 4. Gynacantha bayadera Selys,1891 C LC 5. Anax ephippiger (Burmeister, 1839) NR LC VR LC Family: Gomphidae (04) 6. Gomphidia 7. Ictinogomphus rapax (Rambur, 1842) VC LC 8. Microgomphus torquatus Selys, 1854 R DD 9. Paragomphus lineatus (Selys,1850) VC LC t-nigrum Selys, 1854 Family:Libellulidae (30) 10. Acisoma panorpoides Rambur, 1842 C LC 11. Brachydiplax sobrina (Rambur, 1842) NR LC 12. Brachythemis VC LC 13. Bradinopyga geminata (Rambur, 1842) VC LC 14. Crocothemis servilia (Drury, 1770) VC LC 15. Diplacodes lefebvrii (Rambur,1842) R LC 16. Diplacodes nebulosa (Fabricius, 1793) R LC 17. Diplacodes trivialis (Rambur,1842) VC LC 18. Indothemis carnatica (Fabricius, 1798) R NT 19. Lathrecista asiatica (Fabricius, 1798) C LC 20. Neurothemis intermedia (Rambur, 1842) VC LC 21. Neurothemis tullia (Drury, 1773) C LC 22. Orthetrum sabina (Drury, 1773) VC LC 23. Orthetrum glaucum (Brauer, 1865) C LC 24. Orthetrum luzonicum (Brauer, 1868) VC LC 25. Orthetrum pruinosum (Burmeister, 1839) VC LC 26. Orthetrum taeniolatum (Schneider,1845) C LC 27. Pantala flavescens (Fabricius, 1798) VC LC 28. Potamarcha congener (Rambur, 1842) VC LC 29. Rhodothemis rufa (Rambur, 1842) VR LC 30. Rhyothemis variegata (Linnaeus, 1763) VC LC 31. Tholymis tillarga(Fabricius, 1798) C LC 32. Tramea basilaris (Palisot de Beauvois, 1807) C LC 33. Tramea limbata (Desjardins, 1832) C LC 34. Trithemis aurora (Burmeister, 1839) VC LC 35. Trithemis festiva (Rambur, 1842) VC LC 36. Trithemis kirbyi Selys, 1891 NR LC 37. Trithemis pallidinervis (Kirby, 1889) VC LC 38. Urothemis signata Rambur, 1842 R LC 39. Zyxomma petiolatum Rambur, 1842 C LC contaminata (Fabricius,1793) Family: Macromiidae (02) 40. Epophthalmia vittata Burmeister,1839 C LC 41. Macromia cingulata Rambur, 1842* C LC Suborder: Zygoptera (Damselflies) Family: Chlorocyphidae (01) IAEES www.iaees.org Arthropods, 2022, 11(1): 56-64 62 42. Libellago lineata (Burmeister, 1839) VC LC Family: Coenagrionidae (13) 43. Agriocnemis pygmaea (Rambur, 1842) VC LC 44. Paracercion calamorum (Ris, 1916) C NA 45. Paracercion malayanum (Selys, 1876) C LC 46. Ceriagrion coromandelianum(Fabricius, 1798) VC LC 47. Enallagma parvum (Selys,1876) VC LC 48. Ischnura aurora (Brauer, 1865) VC LC 49. Ischnura senegalensis(Rambur, 1842) VC LC 50. Pseudagrion spencei Fraser, 1922 NR LC 51. Pseudagrion decorum(Rambur, 1842) VC LC 52. Pseudagrion hypermelas (Selys,1876) R LC 53. Pseudagrion microcephalum (Rambur, 1842) C LC 54. Pseudagrion rubriceps (Selys, 1876b) VC LC 55. Ischnura nursei (Morton,1907) VC LC Family: Lestidae (04) 56. Lestes umbrinus Selys,1891 VC DD 57. Lestes thoracicus Laidlaw, 1920 R LC 58. Lestes viridulus VC LC 59. Lestes nodalis Selys, 1891 VR LC Rambur, 1842 Family: Platycnemididae (03) 60. Copera marginipes (Rambur, 1842) VC LC 61. Disparoneura quadrimaculata (Rambur,1842) VC LC 62. Elattoneura nigerrima (Laidlaw, 1917) NR DD Odonates are predatory in nature. They are also a good source of energy to different animals, especially for birds and other insects such as spiders. Being a predator both at larval and adult stage, their role as an important component in wetland. In addition to their significant role in ecosystem function, their value as indicators' of quality of biotope is now being increasingly recognized (Subramanian and Sivaramakrishnan, 2002). The Families Libellulidae (31) and Coenagrionidae (15) are dominant in Seloo city. Earlier studies on the Maharashtra odonates from other region also have reported Libellulidae family in Dragonfly and in Damselfly family Coenagrionidae as a dominant (Tiple et al., 2008; Tiple, 2012; Andrew, 2013; Kulkarni and Subramanian, 2013; Koparde et al., 2014 ). The area of Seloo city is highly disturbed. The Brachythemis contaminate, Orthetrum sabina, Pantala flavescens, Bradinopyga geminate, Ceriagrion coromandelianum, Agriocnemis pygmaea was commonly sighted in human settlement areas and its presents clearly indicates the polluted water quality of that area. Human development activity is expected to have a deleterious impact on Odonata populations; it only because of construction of buildings and concretes replaces or reduces the area of natural and semi-natural habitats. The quality of residual habitats may also be adversely affected by various forms of pollutants (Tiple and Chandra, 2013). Tiple and Koparde (2015) reported 134 species of Odonata from Maharashtra and Tiple et al. (2012) reported 82 species of Odonates from Vidarbha Region of Maharashtra state. The present study on the Odonata of Seloo city revealed the presence of 62 species which account 76% of total species reported in Vidarbha IAEES www.iaees.org Arthropods, 2022, 11(1): 56-64 63 Region and 46% of species reported in Maharashtra State. The Seloo city seems to be having rich Odonate diversity of 62 varieties of species. Probably due to the presence of rivers, lakes and temporary and permanent flowing or still water bodies with dense shrub and tree vegetation a major attraction to the Odonata species. The observations recorded in the present study may prove valuable as a reference for assessing the changes due to the environmental conditions in the locality, in future. Acknowledgements The authors are thankful to the Principal, Vidyabharti College Seloo, Wardha for providing facilities and kind encouragement. References Andrew RJ. 2013. Odonates of Zilpi Lake of Nagpur (India) with a note on the emergence of the libellulid dragonfly, Trithemis pallidinervis. Journal on New Biological Reports, 2(2): 177-187 Andrew RJ, Subramaniam KA, Tiple AD. 2008. A Handbook on Common Odonates of Central India. South Asian Council of Odonatology, Nagpur, India Bedjanič M, Kalkman V, Subramanian K. 2020. A new species of Orthetrum Newman, 1833 (Odonata: Libellulidae) from the Andaman Islands, India. Zootaxa, 4779(1): 91-100 Dawn, P. 2021. A new species of Cephalaeschna Selys, 1883 (Odonata: Anisoptera: Aeshnidae) from Neora Valley National Park, West Bengal, India, with notes on C. acanthifrons Joshi & Kunte, 2017 and C. viridifrons (Fraser, 1922). Zootaxa, 4949(2): 371-380 Dijkstra KDB, Lewington R. 2006. Field Guide to the Dragonflies of Britain and Europe. British Wildlife Publishing, UK Fraser FC. 1933. The Fauna of British India including Ceylon and Burma. Odonata Vol. I. Taylor and Francis Ltd. London, UK Fraser FC. 1934. The Fauna of British India including Ceylon and Burma. Odonata Vol. II. Taylor and Francis Ltd. London, UK Fraser FC. 1936. The Fauna of British India including Ceylon and Burma. Odonata Vol. III. Taylor and Francis Ltd., London, UK IUCN. 2020. International Union of Conservation Network red-list of threatened species. Available at: http://www.iucnredlist.org/. Joshi S, Sawant D. 2020. Description of Bradinopyga konkanensis sp. nov. (Odonata: Anisoptera: Libellulidae) from the coastal region of Maharashtra, India. Zootaxa, 4779(1): 65-78. Koparde P, Mhaske P, Patwardhan A. 2014. New records of dragonflies and damselflies (Insecta: Odonata) from the Western Ghats of Maharashtra, India Journal of Threatened Taxa, 6(5): 5744-5754 Kulkarni AS, Subramanian KA. 2013. Habitat and seasonal distribution of Odonata (Insecta) of Mula and Mutha river basins, Maharashtra, India. Journal of Threatened Taxa, 5(7): 4084-4095 Mitra TR. 1986. Note on the Odonata fauna of Central India. Zoological Survey of India, 83: 69-81 Payra A, Tiple AD. 2019. Odonata fauna in adjoining coastal areas of Purba Medinipur District, West Bengal, India. Munis Entomology and Zoology, 14(2): 358-367 Payra A, Subramanian KA, Chandra K, Tripathy B. 2020. A first record of Camacinia harterti Karsch, 1890 (Odonata: Libellulidae) from Arunachal Pradesh, India. Journal of Threatened Taxa, 12(8): 15922-15926 Schorr M, Paulson D. 2021. World Odonata List. https://www.pugetsound.edu/academics/academicresources/slater-museum/. Accessed 1 October 2021 IAEES www.iaees.org 64 Arthropods, 2022, 11(1): 56-64 Subramanian KA, Babu R. 2017. Checklist of Odonata (Insecta) of India, Version 3.0. www. zsi.gov.in Subramanian KA, Babu R. 2020. Dragonflies and damselflies (Insecta: Odonata) of India. In: Indian Insects Diversity and Science (Ramani S, Prrashanth M, Yeshwanath HM, eds). 29-45, CRC Press, Taylor & Francis, USA Subramanian KA, Sivaramakrishnan KG. 2002. Conservation of Odonate fauna in Western Ghats. Vistas of Entomological Research For The New Millennium. India Subramanian KA. 2009. Dragonflies and Damselflies of Peninsular India - A Field Guide. Vigyan Prasar, Noida, India Talmale SS, Tiple AD. 2013. New records of damselfly Lestes thoracicus Laidlaw, 1920 (Odonata: Zygoptera: Lestidae) from Maharashtra and Madhya Pradesh states, central India. Journal of Threatened Taxa, 5(1): 3552-3555 Tiple AD. 2015. New Record of Damselfly Lestes nodalis Selys (Odonata: Lestidae) from Central India. ENVIS (SACON) Newsletter, 11(1): 6-7 Tiple AD. 2018. Butterflies (Lepidoptera: Rhopalocera) of the Bor Wildlife Sanctuary, Wardha, Maharashtra, Central India. Biodiversity Journal, 9(3): 171-180 Tiple AD. 2018. Butterfly diversity in relation to a relative abundance and status in Seloo city, Wardha Maharashtra, Central India. International Journal of Research In Biosciences, Agriculture and Technology, 1:1-5. Tiple AD. 2020. Dragonflies and Damselflies (Odonata: Insecta) of the Bor Wildlife Sanctuary,Wardha, Maharashtra, Central India. Travaux du Muséum National d’Histoire Naturelle “Grigore Antipa”, 63(2): 131-140 Tiple AD, Andrew RJ, Subramanian KA, Talmale SS. 2013. Odonata of Vidarbha region, Maharashtra state, Central India. Odonatologica, 42(3): 237-245 Tiple AD, Chandra K. 2013. Dragonflies and Damselflies (Insecta, Odonata) of Madhya Pradesh and Chhattisgarh States, India. Care 4Nature, 1(1): 2-11 Tiple AD, Gathalkar GB, Talmale SS. 2014. New record of dragongfly Ictinogomphus angulosus (Selys, 1854) from State Maharashtra, India. Ambient Science, 1: 56-58 Tiple AD, Khurad AM, Andrew RJ. 2008. Species diversity of Odonata in and around Nagpur City, Central India. Fraseria, 7: 41-45 Tiple AD, Koparde P. 2015. Dragonflies and Damselflies (Insecta, Odonata) of Maharashtra States, India. Journal of Insect Science, 15(1): 1-10 Tiple AD, Paunikar S, Talmale SS. 2012. Dragonflies and Damselflies (Odonata: Insecta) of Tropical Forest Research Institute, Jabalpur, Madhya Pradesh, central India. Journal of Threatened Taxa, 4(4): 2529-2533 IAEES www.iaees.org Arthropods, 2022, 11(1): 65-71 Article Determination of fenpropathrin residue by QuEChERS method and GC/MS Bahareh Rafiei1, Seyed Reza Bastan2 1 Plant Protection Research Department, Guilan Agricultural and Natural Resources Research and Education Center, Agricultural Research, Education and Extension Organization (AREEO), Rasht, Iran 2 Department of Agronomy, Rasht Branch, Islamic Azad University, Rasht, Iran E-mail: B.Rafiei@areeo.ac.ir Received 15 November 2021; Accepted 25 December 2021; Published 1 March 2022 Abstract Chemical pesticides are used worldwide to control pests. This study investigated the residues of the pesticide fenpropathrin in greenhouse tomatoes (Vendor variety). Sample preparation was performed by QuEChERS method, and solid-phase extraction (SPE) cartridges were used for purification. Residual evaluation of this pesticide was carried out using doses (1, 2, and 4 g/lit) in greenhouse tomatoes. Samples were collected at intervals of one, three, five, seven, and ten days after spraying and analyzed by chromatographic gas spectroscopy. The results were compared with the maximum residue level (MRL = 0.5 mg/kg) established by Codex Alimentarius. The recovery of fenpropathrin was estimated to be 98.68% at a level of 0.5 ppm. In addition, the preharvest period for fenpropathrin 2 g/lit was determined in greenhouse tomatoes 3 days after spraying. The results also illustrated that increasing the dose of pesticide enhanced the remaining amount. Keywords fenpropathrin; residue; QuEChERS; SPE; GC-MS. Arthropods ISSN 2224­4255 URL: http://www.iaees.org/publications/journals/arthropods/online­version.asp RSS: http://www.iaees.org/publications/journals/arthropods/rss.xml E­mail: arthropods@iaees.org Editor­in­Chief: WenJun Zhang Publisher: International Academy of Ecology and Environmental Sciences 1 Introduction One of the most important agricultural problems in the world is the occurrence of pests, as about one-third of the world's agricultural product is destroyed by pests at various stages of production. The protection of plant crops is a necessary part of agricultural production, which increases crop yield in terms of both quantity and quality. Therefore, the importance of pesticides in agriculture as a control strategy is clear. Although organic farming is becoming increasingly important, pesticides are still used for pest control in many countries around the world, and the residues of these compounds have many negative effects on human health and the environment (Zhang, 2018; Zhang et al., 2019; Bastan et al., 2021; Romero-González, 2021). Fenpropathrin is a contact-gastrointestinal insecticide and acaricide. This pesticide can control a variety of IAEES www.iaees.org 66 Arthropods, 2022, 11(1): 65-71 mites as well as insects such as whiteflies, butterfly larvae, minnows, leaf-feeding insects, aphids, psyllids, and stem-feeding pests. This pesticide is used in greenhouses (cucumbers, tomatoes, ornamental plants, so on.) but is dangerous to humans and animals (LD50 = 70-164 mg/kg) (Rafiei et al., 2010). Fenpropathrin causes degeneration of dopaminergic neurons and parkinsonism (Jiao et al., 2020). The residues of fenpropathrin have been studied in products such as cucumber, tomato puree, orange nectar, orange juice and canned onion, papaya and various vegetables in different countries (Lopez- Lopez et al., 2001; Sannino et al., 2002; Parrilla Vazquez et al., 2008; Ramadan et al., 2020; Xiao et al., 2021). Different methods have been applied to extract pesticide residues from foods, and today QuEChERS (Quick, Easy, Cheap, Effective, Rugged, and Safe) followed by clean-up steps involving dispersive solid phase extraction (dSPE) is the most commonly used procedure (Reis et al., 2020). The current experiment aimed to investigate the residual levels of different doses of fenpropathrin in greenhouse tomatoes at intervals after pesticide application using the QuEChERS method and SPE clean-up. 2 Materials and Methods 2.1 Standards and materials All chemicals and solvents were of analytical quality grade. Analytical grade of fenpropathrin applied for GCMS (Gas chromatograph-masses) analysis was purchased from Sigma-Aldrich (USA). HPLC grade acetonitrile (MeCN), n-hexane (Hex), and methanol (MeOH) were purchased from Merck (Darmstadt, Germany). Syringe filters (color coded, Chromafil MV, 25 mm, 0.45 µm) used in the sample extraction step was acquired from MACHEREY-NAGEL (Dueren, Germany). Chromabond SPE Cartridge (Chromabond (C18) 45 µm, 3 mL/200 mg) used in sample purification step was purchased from MACHEREY-NAGEL (Dueren, Germany). 2.2 Design of experiment The experiments were conducted in a greenhouse in Markazi province, Iran (35° 2' 51.211'' N, 48° 40' 0.156'' E) and were carried out on a Vendor table tomato cultivar. The average greenhouse temperature was recorded as 21-24°C at night and 28-31°C during the day with 73% humidity. A randomized complete block design was used with three replicates. The experimental treatments included: fenpropathrin (recommended half dose: 1g/lit), fenpropathrin (recommended doses: 2 g/lit), fenpropathrin (double recommended dose: 1g/lit), and water (control). Tomato plants were sprayed twice with EC 10% formulation of fenpropathrin at concentrations of 1, 2, and 3 g/lit at one-week intervals. In the control plants, only water was sprayed instead of the pesticide. 2.3 Sampling Sprayed tomato fruits were collected one hr after the second spraying and then 1, 3, 7, and 10 days after treatment. At each sampling, 1 kg of tomato was harvested from each replication. Samples were packed in polythene bags and transported to the laboratory after labeling with the relevant information while maintaining the cold chain. 2.4 Extraction procedure Sample preparation for pesticide extraction was conducted according to the QuECHERS method (Paya et al., 2007). First, the samples were divided into two-cm pieces and 200 grams were homogenized. Then, 10 ml acetonitrile, 10 ml methanol, and 10 ml distilled water were added to 20 g of the homogenized sample and stirred for 30 minutes using a shaking machine. Then, the mixture was centrifuged at 2500 rpm for 10 minutes. The supernatant, the upper part of the sample which is a clear liquid, was passed through a syringe filter. At this stage, pH was measured, and pH requirements were adjusted if necessary. 2.5 Purification IAEES www.iaees.org Arthropods, 2022, 11(1): 65-71 67 Solid phase cartridges (C18) (200 mg, 3 mL, 45 µm from Machery-Nagel) were used as sorbent materials for purification. First, column preparations were performed, and the cartridges were washed with 10 cc of normal hexane. Next, 5 cc of distilled di-ionized water was used followed by 5 cc of acetonitrile. In the next step, the extract was passed through the column. Finally, the cartridge was washed with 5 cc of ethyl acetate. The extracts were collected in glass vials. Then 10 cc of normal hexane was passed through the column, and the extracts were collected as in the previous step. The extractions were concentrated with a slow nitrogen flow, and the total volume of the extracts reached 200 ml. 2.6 Pesticide analysis Pesticide residues were measured by chromatography-mass spectrometry. The injection temperature was set at 200°C, the mass detector temperature was set at 160°C, and the capillary column (HP5) had a length of 30 m, an inner diameter of 0.53 mm, and an adsorbent thickness of 25 m. The capillary column (HP5) was used to measure pesticide residues. 2.7 Method of validation Standard and working solutions were prepared for the construction (0.01 - 0.5 mg/kg) of calibration curves and recovery tests and were stored in the dark at 4°C. Limit of detection (LOD) and limit of quantification (LOQ) were according to the European Union SANCO/12495/2011 guidelines. 2.8 Statistical analysis Statistical analyses were performed using SAS V.8.0 software. One-way analysis of variance (ANOVA) was used for statistical analysis. The results were reported significant at the 5% level (p < 0.05). 3 Results The retention time was determined from the peaks of the standard pesticide. The chromatograms of different days after spraying were compared with the standard peaks, and qualitative identification was performed. Then the residual pesticide content was quantitatively evaluated by comparing the curved surface area of each sample. The average recovery of fenpropathrin at 0.5 ppm was reported to be 98.68%. The average pesticide residue in tomatoes sprayed with fenpropathrin (1 g/lit) was 2.984 mg/kg one hour after spraying and 1.537 mg/kg one day after spraying. The residue of this toxin on the third day after spraying was below the MRL (0.5 mg/kg) established by Codex Alimentarius (WHO/FAO, 2005) (Table 1, Fig. 1). Fig. 1 The mean of fenpropathrin (1 g/lit) residues on different days in greenhouse tomatoes. IAEES www.iaees.org Arthropods, 2022, 11(1): 65-71 68 Table 1 Mean Comparison of Fenpropathrin (1 g/lit) residues with MRL (mg/kg). Times after application of p t Mean ± SE 0.008 4.28 2.984± 0.020 1 hour 0.007 6.101 1.537 ± 0.020 1 day 0.026 0.412 0.351 ± 0.020 3 day 0.032 0.321 0.295 ± 0.020 5 day 0.041 0.085 0.079 ± 0.020 7 day 0.015 0.072 0.053 ± 0.020 10 day pesticide The average residue of fenpropathrin (2 g/lit) one hour after spraying was 4.85 mg/kg and one day after spraying was 3.037 mg/kg. The comparison of the mean residue of the pesticide (2 g/lit) with (MRL=0.5 mg/kg) illustrated that there was no significant difference with the MRL on the third day (Table 2, Fig. 2). Fig. 2 The mean of fenpropathrin (2 g/lit) residues on different days in greenhouse tomatoes Table 2 Mean Comparison of Fenpropathrin (2 g/lit) residues with MRL (mg/kg). IAEES Times after application of p t Mean ± SE 0.011 9.06 4.85 ± 0.031 1 hour 0.010 3.63 3.037 ± 0.031 1 day 0.015 2.32 0.508 ± 0.031 3 day 0.032 0.78 0.435 ± 0.031 5 day 0043 0.04 0.289 ± 0.031 7 day 0.029 0.003 0.199 ± 0.031 10 day pesticide www.iaees.org Arthropods, 2022, 11(1): 65-71 69 The residue of fenpropathrin (4 g/lit) was calculated to be 6.546 mg/kg one hour after spraying and 5.843 mg/kg one day after spraying. Comparison of the average residue of the pesticide (4 g/lit) and (MRL) revealed that there was no significant difference with the MRL on the tenth day (Table 3, Fig. 3). The LODs of fenpropathrin were 0.108 μg/kg, and the LOQs were 0.452 μg/kg in the original samples. Moreover, the calibration curve was linear with an R2 of 0.99. Fig. 3 The mean of fenpropathrin (4 g/lit) residues on different days in greenhouse tomatoes. Table 3 Mean Comparison of fenpropathrin (4 g/lit) residues with MRL (mg/kg). Times after application of p t Mean ± SE 0.008 9.96 6.546 ± 0.041 1 hour 0.012 3.48 5.843 ± 0.041 1 day 0.106 2.67 3.646 ± 0.041 3 day 0.181 0.83 0.842 ± 0.041 5 day 1.167 0.057 0.698 ± 0.041 7 day 2.22 0.001 0.439 ± 0.041 10 day pesticide 4 Discussion Researchers have obtained remarkable results regarding the side effects of pesticides and their fate in the environment, and for each pesticide, conditions such as application methods (formulation and recommended concentrations), application time, and exposure duration have been considered. The results of the present study have shown that the residual amount of fenpropathrin pesticide depends on the consumed dose. The higher the consumed dose of pesticide is, the longer it takes to fall below the MRL that is safe for humans. When the dose was 2 g/lit, it was below the MRL limit (0.5 mg/kg) on the third day after spraying; by the tenth day after spraying, the remaining pesticides were no longer measurable. On the IAEES www.iaees.org 70 Arthropods, 2022, 11(1): 65-71 other hand, when twice the recommended dose of fenpropathrin was used, the pre-harvest time increased significantly, and on the seventh day the residual MRL was elevated. According to studies in Spain, the residual amount of fenpropathrin pesticide in cucumber using SPE and HPLC three days after harvest was 0.39 ppm, and the pesticide recovery was between 96% and 116%. In another study, the residual amount of pesticide in tomato puree and orange nectar was evaluated using gas chromatography-mass spectrometry. The pesticide recovery rate was between 70.2% and 96% (Sannino et al., 2002). In a third study, the recovery rate of fenpropathrin residues in cucumber was estimated at 63% to 108% (Parrilla Vazquez et al., 2008). In a study of pesticide residues on various vegetables in Saudi Arabia, fenpropathrin was one of the pesticides in these products. Xiao et al. (2021) also investigated fenpropathrin residues in Chaenomeles speciosa was safe for humans when the pesticide was applied at twice the recommended dose (GAP) compared to the EU maximum residue levels (EU, 0.01 mg/kg) 14 days after the last application. It is also important to note that the MRL for each pesticide compound varies from country to country and from product to product. The MRL for each product is set based on its toxicity, its production method (greenhouses or farms), its per capita consumption, and its application method in each country, among other factors. The pre-harvest time of a pesticide depends on various factors such as the amount of pesticide used, climatic conditions, irrigation cycle, species and variety planted, planting date, and type of pesticide formulation. In the current study, two sprays were made in the greenhouse, but because greenhouse pests are sporadic and some have multiple generations, sprays are often repeated during a growing season. Moreover, producers sometimes apply more than is recommended. Therefore, it is possible that the frequency of spraying leads to an increase in the amount of residue in the product, even beyond the determined values. References Bastan SR, Rafiei B. 2021. Evaluation of Permethrin residue in greenhouse tomatoes. Genetic Engineering and Biosafety Journal, 9(1): 19-27 Fevery D, Houbraken M, Spanoghe P. 2016. Pressure of non-professional use of pesticides on operators, aquatic organisms and bees in Belgium. Science of the Total Environment, 550: 514e521 Jiao Z, Wu Y, Qu S. 2020. Fenpropathrin induces degeneration of dopaminergic neurons via disruption of the mitochondrial quality control system. Cell Death Discovery, 6: 78 Lopez-Lopez T, Gil-Garciaa MD, Martinez-Vidal JL, Martinez Galeraa M. 2001. Determination of pyrethroids in vegetables by HPLC using continuous on-line post-elution photoirradiation with fluorescence detection. Analytica Chimica Acta, 447(1-2): 101-111 Parrilla Vazquez P, Mughari R, Martinez MG. 2008. Solid-phase microextraction (SPME) for the determination of pyrethroids in cucumber and watermelon using liquid chromatography combined with post-column photochemically induced fluorimetry derivatization and fluorescence detection. Analytica Chimica Acta, 706: 74-82 Paya P, Anastassiades M, Mack D, Sigalova I, Tasdelen B, Oliva J, Barba A. 2007. Analysis of pesticide residues using the Quick Easy Cheap Effective Rugged and Safe (QuEChERS) pesticide multiresidue method in combination with gas and liquid chromatography and tandem mass spectrometric detection, Analytical Bioanal Chemistry, 389(6): 1697-1714 Rafiei B, Imani S, Bastan R. 2016. Determination of residue of Deltamethrin on greenhouse cucumber. Journal of Entomological Research, 7(4): 307-316 IAEES www.iaees.org Arthropods, 2022, 11(1): 65-71 71 Rafiei B, Imani S, Alimoradi M, Shafiee H, Khaghani S, Bastan SR. 2010. Survey on residuals of Fenpropathrin in greenhouse cucumber. Journal of Entomological Research, 2(3): 193-201 Ramadan MFA, Abdel-Hamid MMA, Altorgoman MMF, AlGaramah HA, Alawi MA, Shati AA, Shweeta HA, Awwad NS. 2020. Evaluation of pesticide residues in vegetables from the Asir Region, Saudi Arabia. Molecules, 25: 205-225 Romero-González R. 2021. Detection of residual pesticides in foods. Foods, 10: 1113 Sannino A, Bandini M, Bolzoni L. 2003. Determination of pyrethroid pesticide residues in processed fruits and vegetables by gas chromatography with electron capture and mass spectrometric detection. Journal of AOAC International, 86(1): 101-108 Xiao J, Wang F, Ma JJ, Xu X, Lia M, Fang QK, Cao HQ. 2021. Acceptable risk of fenpropathrin and emamectin benzoate in the minor crop Mugua (Chaenomeles speciosa) after postharvest processing. Environmental Pollution, 276: 116716 Zhang WJ. 2018. Global pesticide use: Profile, trend, cost / benefit and more. Proceedings of the International Academy of Ecology and Environmental Sciences, 8(1): 1-27 Zhang XQ, Hao XX, Huo SS, LinWZ, Xia XX, Liu K, Duan BH. 2019. Isolation and identification of the Raoultella ornithinolytica-ZK4 degrading pyrethroid pesticides within soil sediment from an abandoned pesticide plant. Archives of Microbiology, 201(9): 1207e1217 IAEES www.iaees.org Arthropods Arthropods play the role of both pests and beneficial organisms. Some arthropods are important crop pests but others are natural enemies. Some arthropods are important health pests but many crustaceans are important food sources of humankinds. Arthropods govern the structures and functions of natural ecosystems, but are always ignored by researchers. On the global scale, the surveys of mammals, birds and vascular plants were relatively perfect because they were economically important and easily surveyed. However, arthropods, despite their ecological and economical importance, have not yet been fully surveyed and recorded due to their difficulties to be sampled. The research on arthropods must be further promoted. The journal, Arthropods, aims to provide a public and appropriate platform for the publication of studies and reports on arthropods. Arthropods (ISSN 2224-4255) is an international open access (BOAI definition), open peer reviewed online journal (users are free to read, download, copy, distribute, print, search, or link to the full texts of the articles) devoted to the publication of articles on various aspects of arthropods, e.g., ecology, biogeography, systematics, biodiversity (species diversity, genetic diversity, et al.), conservation, molecular biology, biochemistry, physiology, control, etc. The journal provides a forum for examining the importance of arthropods in biosphere (both terrestrial and marine ecosystems) and human life in such fields as agriculture, forestry, fishery, environmental management and human health. The scope of Arthropods is wide and includes all arthropods-insects, arachnids, crustaceans, centipedes, millipedes, and other arthropods. Articles/short communications on new taxa (species, genus, families, orders, etc.) of arthropods are particularly welcome. Authors can submit their works to the email box of this journal, arthropods@iaees.org. All manuscripts submitted to Arthropods must be previously unpublished and may not be considered for publication elsewhere at any time during review period of this journal. In addition to free submissions from authors around the world, special issues are also accepted. The organizer of a special issue can collect submissions (yielded from a research project, a research group, etc.) on a specific topic, or submissions of a conference for publication of special issue. Editorial Office: arthropods@iaees.org Publisher: International Academy of Ecology and Environmental Sciences Address: Unit 3, 6/F., Kam Hon Industrial Building, 8 Wang Kwun Road, Kowloon Bay, Hong Kong E-mail: office@iaees.org Arthropods ISSN 2224-4255 Volume 11, Number 1, 1 March 2022 Articles Effect of continuous rearing generations on some biological parameters of Habrobracon hebetor (Hymenoptera: Braconidae) under insectarium conditions Ghadir Momenian, Mohammad Hasan Sarayloo, Ali Afshari 1-17 Diversity of spiders (Chelicerata: Araneae) in Uttar Pradesh and Uttarakhand, India Rajendra Singh, Garima Singh 18-55 Dragonflies and damselflies (Odonata: Insecta) of the Seloo city, Wardha, Maharashtra, Central India Ashish Tiple, Rahul Bhende, Parag Dandge 56-64 Determination of fenpropathrin residue by QuEChERS method and GC/MS Bahareh Rafiei, Seyed Reza Bastan 65-71 IAEES http://www.iaees.org/