Green Processing and Synthesis 2020; 9: 415–427 Research Article Raghad R. Alzahrani*, Manal M. Alkhulaifi*, and Nouf M. Al-Enazi In vitro biological activity of Hydroclathrus clathratus and its use as an extracellular bioreductant for silver nanoparticle formation https://doi.org/10.1515/gps-2020-0043 received April 22, 2020; accepted June 28, 2020 Abstract: The adaptive nature of algae results in producing unique chemical components that are gaining attention due to their efficiency in many fields and abundance. In this study, we screened the phytochemicals from the brown alga Hydroclathrus clathratus and tested its ability to produce silver nanoparticles (AgNPs) extracellularly for the first time. Lastly, we investigated its biological activity against a variety of bacteria. The biosynthesized nanoparticles were characterized by UV-visible spectroscopy, Fourier-transform infrared spectroscopy, dynamic light scattering, transmission electron microscopy, and energy-dispersive spectroscopy. The biological efficacy of AgNPs was tested against eighteen different bacteria, including seven multidrugresistant bacteria. Phytochemical screening of the alga revealed the presence of saturated and unsaturated fatty acids, sugars, carboxylic acid derivatives, triterpenoids, steroids, and other components. Formed AgNPs were stable and ranged in size between 7 and 83 nm and presented a variety of shapes. Acinetobacter baumannii, Staphylococcus aureus, Methicillin-resistant S. aureus (MRSA), and MDR A. baumannii were the most affected among the bacteria. The biofilm formation and development assay presented a noteworthy activity against MRSA, with an inhibition percentage of 99%. Acknowledging the future of nanoantibiotics encourages scientists to explore and enhance their potency, notably if they were obtained using green, rapid, and efficient methods.  * Corresponding author: Raghad R. Alzahrani, Department of Botany and Microbiology, College of Science, King Saud University, Riyadh 11451, Saudi Arabia, e-mail: raghad.r.alzahrani@gmail.com * Corresponding author: Manal M. Alkhulaifi, Department of Botany and Microbiology, College of Science, King Saud University, Riyadh 11451, Saudi Arabia, e-mail: manalk@ksu.edu.sa Nouf M. Al-Enazi: Department of Biology, College of Science and Humanities in Al-Kharj, Prince Sattam Bin Abdulaziz University, Al-Kharj 11942, Saudi Arabia, e-mail: n.alenazi@psau.edu.sa Open Access. © 2020 Raghad R. Alzahrani et al., published by De Gruyter. Public License. Keywords: brown algae, silver nanoparticles, biosynthesis, bio-nanoparticles, MDR bacteria Abbreviations AgNPs AgNPCB AgNPQB ATCC DLS DMSO EDS FTIR TEM silver nanoparticles silver nanoparticles synthesized by H. clathratus crude methanol extract silver nanoparticles synthesized by H. clathratus aqueous solution American Type Culture Collection dynamic light scattering dimethyl sulfoxide energy-dispersive spectroscopy Fourier-transform infrared spectroscopy transmission electron microscopy 1 Introduction Despite the progress of bacterial resistance to antibiotics, the development of a new, effective antibiotic against multidrug-resistant (MDR) bacteria has been progressing at a slow pace. Recent studies have suggested using nanotechnology as a promising strategy to challenge bacterial resistance to antibiotics using natural constituents as reducing agents to synthesize nanoparticles (NPs), although this approach could be limited due to the toxicity and unpredictability of NPs [1]. There were several cases that reported the existence of MDR bacteria in Saudi Arabia. For example, a few cases of MDR tuberculosis in different regions of Saudi Arabia were observed [2]. Besides, prevalence rates of extendedspectrum beta-lactamase (ESBL) producing isolates, such as Escherichia coli and Klebsiella pneumoniae, were 29% and 65%, respectively [3]. Bacterial biofilm formation is a significant virulence factor that aids in antibiotic resistance and bacterial survival, especially on This work is licensed under the Creative Commons Attribution 4.0 416  Raghad R. Alzahrani et al. medical devices [4]. Seaweeds are considered as promising sources of new antibacterial agents. More than one thousand bioactive components from a marine source, including algal secondary metabolites, have been globally defined as possible antibacterial, antiviral, anti-inflammatory, and anticancer agents [5]. Brown algae are known in the field of medicine and nutrition. Their medical history goes back to time when they were used for treating diarrhea, urinary disorders, and chronic bacterial infections in Europe [6]. Their cell walls comprise a distinct variety of bioactive polysaccharides, as well as fewer amounts of phenolic substances, proteins, and halide compounds such as iodide [7]. The thallus of Hydroclathrus clathratus appears perforated in a sponge resembling form; hence, its Latin name clathratus means latticed, indicating its morphology. The alga is distributed worldwide in warm seas and calm, shallow areas such as Europe, both coasts of Africa, the Pacific Islands, Asia, Australia, North America, and South America, from California through Chile and the Gulf of Mexico [8]. Algal phytochemicals contain functional groups, such as hydroxyl, carboxyl, and amino groups, which are beneficial for the reduction and capping process required to synthesize a stable and robust coating on metal NPs [9]. Scientists recommend algae in biological NP synthesis, due to their high metal uptake capacity and their minimal cost of production, which are golden advantages, compared to other bioreductants [10]. This study aims to screen the phytochemicals of the brown alga Hydroclathrus clathratus (methanol extract and raw powder). Then, we Figure 1: Hydroclathrus clathratus on the collection site. investigate its bioreducing ability of silver nitrate to silver nanoparticles (AgNPs) and explore its biological activity as an antibacterial and antibiofilm agent against a number of pathogenic bacteria, including MDR bacteria. 2 Materials and methods 2.1 Algae collection and preparation H. clathratus (Figure 1) was collected from the northwest coast of Al-Haraa, Umluj City, Red Seashore, Kingdom of Saudi Arabia, in April 2017. The algae were kept in ice packs in plastic bags containing seawater for preservation. They were then washed and shade dried at room temperature and finely powdered using an electric coffee grinder. The algae were then preserved in tight dark containers in the freezer before use. H. clathratus was identified, according to [11,12]. 2.2 Crude methanol extraction H. clathratus (219 g) was saturated three times in 1 L of methanol for 72 h. The mixture was agitated (WNB shaker) to ensure constant agitation during the saturation procedure. Methanol extracts were collected, Bio-AgNPs and their biological activity filtered, and combined. The combined extract was concentrated by evaporating methanol on a rotavapor at ±50°C; after evaporating the solvent, the sample was preserved at room temperature. At the onset of each biological test, a stock solution was prepared at a concentration of 100 mg/mL in 100% DMSO. The extract was then kept in a sterile tube at 4°C until use.  417 however, due to intensity of the green chlorophyll pigment in the NPs formed by the methanol extract (AgNPCB), it was essential to use UV-vis spectroscopy to confirm their formation. 2.6 Characterization of biosynthesized silver NPs 2.3 Raw algal powder solution preparation To prepare the raw algal powder, 5 g of the powdered algae was added to 100 mL of distilled water for easier handling. The mixture was agitated on a magnetic stirrer for 5 h at room temperature and then filtered. The filtrate was then kept in a sterile dark lid flask at 4°C. 2.4 Qualitative phytochemical screening of algal powder and methanol extract We studied the chemical compositions of both the algal crude methanol extract and the raw powder by gaschromatography mass spectroscopy (GC-MS) on the Shimadzu model 2010 plus (Japan). The samples were prepared as follows: 247 mg of raw algal powder was dissolved in 20 mL of 3:1 dichloromethane to methanol; in contrast, 104 mg of algal methanol extract was dissolved in 20 mL of methanol. The samples were then analyzed by GC-MS, according to the following parameters. The MS model QP 2010 ultra and injector model AOC-20i were used and operated in total ion chromatogram scan mode and single ion monitoring ion mode to obtain the retention time of each unidentified compound in the mixture extract samples. After adjustment, we obtained sufficient and adequate separation. The obtained AgNPs were first characterized by UV-vis spectroscopy (Libra S60PC) in the range between 350 and 750 nm. The morphological assessment of AgNPs was conducted using both transmission electron microscopy (TEM) [JEM 1400] at 80 kV accelerating voltage and dynamic light scattering (DLS) (Nano ZS zetasizer system [Malvern Instruments]), which was also used to clarify the dispersity of formed NPs. Fourier-transform infrared spectroscopy (FTIR) [PerkinElmer FTIR system spectrum BX] was used to investigate the involvement of functional groups in the formation of AgNPs in the range between 4,000 and 400 cm−1. Lastly, evaluation of the elemental silver percentage was performed by energydispersive spectroscopy (EDS) [JSM-6380 LA]. 2.7 Antibacterial activity of biologically synthesized AgNPs Agar well-diffusion assay and minimum bactericidal concentration (MBC) on agar were performed following The National Committee for Clinical Laboratory (2006) standards. The minimum inhibitory concentration (MIC) assay was performed following [14], with a few modifications. The tested concentration range varied from 18 to 0.035 mg/mL. All experiments were repeated three times and evaluated in comparison to the positive and negative controls. 2.5 Biological synthesis of AgNPs 2.8 Tested bacteria and media The biological synthesis of AgNPs was conducted after modifying the procedure used by [13]. Algal methanol extract stock (3 mL) and the aqueous solution (3 mL) were added dropwise into a flask containing 22 mL of 1 mM AgNO3 aqueous solution. The flasks were then exposed to heat (50°C) to reduce the reaction time. Color change to brown was the visual assessment of AgNPs’ formation for the algal aqueous solution (AgNPQB); The biologically obtained AgNPs were tested against eighteen bacteria, including seven MDR bacteria. Pure cultures of bacterial strains were obtained from the Microbiology laboratory, in Prince Sultan Military Medical City, Riyadh. The bacteria were first cultured on sheep blood agar (Oxoid) before each experiment. Mueller–Hinton agar was used in the agar well-diffusion 418  Raghad R. Alzahrani et al. assay, and 80 µL of tested AgNPs was loaded into 6 mm wells in the agar. Mueller–Hinton Broth was used in the MIC assay and SBA was used in the MBC assay. All tested plates were incubated at 37°C for 18–24 h. AgNPCB were tested for their antibiofilm ability in triplicate. Six biofilm-forming bacteria were tested (MDR A. baumannii, MDR P. aeruginosa, MRSA, A. baumannii, P. aeruginosa, and S. aureus). The protocol was adapted from [15] with modifications. The assay was performed in a 96-well flat-bottom microplate. The brain–heart infusion broth medium was used. For the biofilm quantitative analysis, we used crystal violet (CV) stain 1% (w/v) aqueous solution. Next, the microplates were destained and examined optically using an ELISA reader (EMax® Endpoint ELISA Microplate Reader) at OD450 nm. The results were averaged, prior to calculating the inhibition percentage using the formula [16]: Percentage of inhibition (%) Well 450 nm – AUntreated control 450 nm ) Component Hexadecanoic acid Oleic acid Octadecanoic acid Tetradecanoic acid Glyceryl-glycoside Others 2.9 Biofilm growth and development inhibition assay of biologically synthesized AgNPs = [1 – (ATreated Table 1: Major components present in H. clathratus methanol extract and powder, and their percent relative concentration (1) × 100] Percent relative concentration (%) H. clathratus MeOH H. clathratus powder 21.67 13.18 2.59 8.68 2.13 51.75 21.51 15.32 23.65 7.08 0 32.44 3.2 Characterization of biosynthesized AgNPs 3.2.1 UV-visible spectroscopy (UV-vis) The confirmation of AgNPs’ formation was assessed visually by a color change. However, due to the intensity of the green pigment in the algal methanol extract, a spectroscopic analysis was required. Biosynthesis of AgNPCB occurred shortly after adding the algal extract (within 1 h), and the highest absorption peak was evident at 411 nm (Figure 2a). In contrast, AgNPQB biosynthesis was monitored for a month; due to the slow formation of AgNPs, the formation of AgNPQB manifested at day 11 and was increased consistently. The highest peak was recorded on the 29th day at 451 nm (Figure 3a). Next, the sample was set for other characteristic techniques and biological tests. 2.10 Statistical analysis Each test was tested three times for repetitions; means and standard deviations were obtained using Microsoft Excel 16.19. 3 Results 3.1 Qualitative phytochemical screening of algal powder and methanol extract We were able to separate more than 60 components. The isolated phytochemicals included saturated and unsaturated fatty acids, sugars, carboxylic acid derivatives, triterpenoids, steroids, and other components such as alkenes and phytols (Table 1). 3.2.2 DLS Zeta size was used to investigate the particle size and distribution of AgNPs. Furthermore, it was used to determine the stability of obtained AgNPs in reference to the polydispersity index (PDI) value. AgNPCB showed an average NP size of 136.9 d.nm and PDI of 0.139 (Figure 2b), implicating the high stability and narrow size distribution of AgNPs. AgNPQB presented an average particle size of 83 d nm and PDI of 0.196 (Figure 3b), signifying its narrow size distribution and high stability. 3.2.3 TEM TEM was used to study the morphological features such as shape and size of biosynthesized AgNPs. The Bio-AgNPs and their biological activity  419 Figure 2: Characterization of biosynthesized AgNPCB by: (a) UV-vis spectroscopy, (b) DLS, (c) EDS of AgNPCB presenting 30% of Ag, (d) TEM imaging, and (e) FTIR of H. clathratus crude methanol extract before (black) and after AgNPCB synthesis (red). Figure 3: Characterization of biosynthesized AgNPQB using: (a) UV-vis spectroscopy, (b) DLS, (c) EDS of AgNPQB presenting 70% of Ag, (d) TEM imaging, and (e) FTIR of H. clathratus raw powder aqueous solution before (black) and after AgNPQB synthesis (red). 420  Raghad R. Alzahrani et al. Table 2: Antibacterial activity of biologically synthesized AgNPCB and AgNPQB Bacteria MDR Acinetobacter baumannii (MRSTAB) ATCC® BAA1790 ESBLs producing Enterobacter cloacae ATCC® BAA 2468 ESBLs producing Escherichia coli* Klebsiella pneumoniae carbapenemase (KPC) ATCC® BAA 2078 Methicillin-resistant S. aureus (MRSA) ATCC 43300 MDR Pseudomonas aeruginosa (MRSTPA) ATCC® BAA 2109 Vancomycin-resistant Enterococcus faecium (VRE) ATCC 700221 A. baumannii ATCC 19606 Salmonella Typhimurium ATCC 14028 E. coli ATCC 35218 K. pneumoniae ATCC® BAA 1706 Enterobacter cloacae ATCC 13047 P. aeruginosa ATCC 27853 Enterococcus faecalis ATCC 29212 S. aureus ATCC 25923 Proteus vulgaris ATCC 49132 Streptococcus pneumoniae ATCC 6305 S. pneumoniae* Inhibition zone (mm) AgNPCB AgNPQB 17.2 ± 1.3 09.3 ± 0.6 ND 07.3 ± 5.7 17.0 ± 0.0 11.8 ± 0.3 08.3 ± 0.6 16.7 ± 2.5 13.0 ± 0.9 12.0 ± 0.5 11.3 ± 0.6 8.0 ± 0.0 13.0 ± 0.0 00.0 ± 0.0 16.3 ± 0.6 9.3 ± 0.6 11.3 ± 0.6 ND 10.0 ± 1.0 ND 10.3 ± 0.6 00.0 ± 0.0 10.3 ± 0.6 08.8 ± 0.8 00.0 ± 0.0 14.3 ± 0.3 11.8 ± 0.8 12.2 ± 0.3 09.5 ± 0.5 11.3 ± 0.6 13.8 ± 1.4 00.0 ± 0.0 15.3 ± 0.6 05.3 ± 4.6 ND 00.0 ± 0.0 The diameter of the well (6 mm) was calculated within the zone of inhibition. The results shown are recorded as means ± standard deviation (SD). * – patient isolate. ND – no data. produced AgNPCB were spherical and polygonal and ranged in size from 7 to 31 nm (Figure 2d). On the contrary, AgNPQB showed a variation in shapes, including spherical, triangle, quadrangular, and rod shapes, and their sizes ranged from 11 to 49 nm (Figure 3d). 3.2.4 EDS EDS was used to conduct chemical composition analysis using a scanning electron microscope and to further confirm the presence of elemental Ag for verifying the reduction reaction. The analysis of synthesized AgNPs within H. clathratus showed the presence of elemental silver at 2.983 keV (Figures 2 and 3c). C═C groups. A band disappeared at 1245.86 cm−1 after AgNPCB production, and a new band emerged in a lower frequency at 1023.61 cm−1 with a higher intensity; this may be a result of Ag bonding with O2 in the C–O group (Figure 2e). In contrast, AgNPQB showed strongly stretched bands at 3427.22 cm−1, suggesting the hydroxyl group’s involvement in the biosynthesis. The peak at 2928.14 cm−1 is attributed to the presence of aliphatic hydrocarbons. The reduction in the band 1635.12 cm−1 may indicate the involvement of benzene in the reduction of AgNPs. A bending of the peak at 1430.42 cm−1 was also observed with a lower intensity. Furthermore, medium stretching of bands was noted between 1262.32 and 1033.63 cm−1 with an evidence of lower intensity. The reduction in the intensity of the peak at 1033.63 cm−1 is attributed to the C–O group (Figure 3e). 3.2.5 FTIR FTIR of AgNPCB unveiled a strongly stretched band at a lower frequency (3400.38 cm−1) than the extract before AgNPCB formation (3413.56 cm−1). Sharp bands were apparent at 2924.12 and 2854.96 cm−1, all of which reduced in intensity after AgNPCB formation. A band at 1714.17 cm−1 shifted to a higher frequency (1,725 cm−1) and reduced in intensity. The peaks from 1419.49 to 1646.16 cm−1 might be assigned to the involvement of 3.3 Antibacterial activity of biologically synthesized AgNPs 3.3.1 Agar well-diffusion assay To assess the biological activity of synthesized AgNPs, we performed an agar well-diffusion assay. Results are Bio-AgNPs and their biological activity shown in (Table 2) that presents the diverse responses of bacteria to AgNPs.  421 Table 4: Antibiofilm growth and development activity of AgNPCB Tested bacteria Biofilm inhibition percentage (%) AgNPs synthesized using H. clathratus 3.3.2 Microtiter MIC and MBC The microtiter MIC and MBC assays showed various values depending on the tested bacteria with different responses to each of the biosynthesized AgNPs (Table 3). Some bacteria exhibited the same MIC and MBC values, indicating that the same concentration was both bacteriostatic and bactericidal. 3.3.3 Antibiofilm activity of AgNPs synthesized using H. clathratus The CV staining assay of produced biofilm investigated the antibiofilm growth and development activity of AgNPs. As shown in (Table 4), all bacterial biofilms were resistant; however, MRSA was highly susceptible to AgNPCB. 4 Discussion In the present study, phytochemicals from the alga H. clathratus were screened and used for the first time as a bioreductant to biosynthesize AgNPs. Producing inorganic AgNPs using “green” techniques is known to be ecofriendly, fast, and efficient. Other merits include their Conc. (100%) MDR Acinetobacter baumannii MRSA MDR Pseudomonas aeruginosa Acinetobacter baumannii Staphylococcus aureus Pseudomonas aeruginosa Conc. (50%) 1.0 1.5 98.9 4.3 99.1 4.6 2.4 0 1.0 3.0 0.5 0.9 Conc. – concentration. All data were averaged before calculating the inhibition percentage. stability, endurance to high temperature, and low toxicity to humans, which can benefit medical applications [9]. The phytochemical screening using GC-MS identified saturated and unsaturated fatty acids, mostly, carboxylic acids, sugars, steroids, phytols, and other products. These components attributed to having an antibacterial activity or suggested as a reducing agent in the green synthesis of NPs [9]. Also, the detected fatty acids in algal extracts such as: palmitic, palmitoleic, stearic, oleic, and linoleic acids have a role in destroying the bacterial cell wall structure and function. They act as anionic surfactants, thus exhibiting their antibacterial and antioxidant activity [16]. Table 3: MIC and MBC of biologically synthesized AgNPCB and AgNPQB Bacteria MDR A. baumannii ESBLs producing E. cloacae ESBLs producing E. coli* K. pneumoniae carbapenemase MRSA MDR P. aeruginosa A. baumannii Salmonella Typhimurium E. coli K. pneumoniae E. cloacae P. aeruginosa Staphylococcus aureus Proteus vulgaris AgNPCB (mg/mL) AgNPQB (mg/mL) MIC MBC MIC MBC 1.125 ± 0.0 2.250 ± 0.0 ND 4.500 ± 0.0 2.250 ± 0.0 0.938 ± 0.3 0.938 ± 0.3 1.875 ± 0.6 0.938 ± 0.3 4.500 ± 0.0 3.750 ± 1.3 0.938 ± 0.3 1.125 ± 0.0 1.875 ± 0.6 1.50 ± 0.6 6.00 ± 2.6 ND >18 ± 0.0 9.00 ± 7.8 2.25 ± 0.0 1.500 ± 0.6 2.250 ± 0.0 1.125 ± 0.0 9.000 ± 0.0 4.500 ± 0.0 1.125 ± 0.0 2.250 ± 0.0 ND 4.50 ± 0.0 ND 9.00 ± 0.0 >18 ± 0.0 9.00 ± 0.0 4.50 ± 0.0 0.938 ± 0.32 2.250 ± 0.00 1.125 ± 0.00 7.500 ± 2.60 0.938 ± 0.32 3.750 ± 1.30 1.875 ± 0.65 4.500 ± 0.00 4.50 ± 0.0 ND 9.00 ± 0.0 >18 ± 0.0 18.0 ± 0.0 9.00 ± 0.0 1.125 ± 0.00 4.500 ± 0.00 1.125 ± 0.00 >18 ± 0.0 1.688 ± 0.97 3.750 ± 1.30 3.750 ± 1.30 4.500 ± 0.00 Results are shown as means ± SD. ND – no data. 422  Raghad R. Alzahrani et al. The obtained AgNPs were characterized, and their biological activity was investigated against a variety of bacteria, including some MDR bacteria. We confirmed the formation of AgNPs, first using UV-vis spectroscopy showing peaks between 400 and 450 nm, specifically at 411 and 451 nm for AgNPCB and AgNPQB, respectively. Thus, this signifies the surface plasmon resonance of silver, which is similar to [17] and other studies that used algae and plants as reducing agents [18,19]. DLS and TEM analyses were performed to evaluate the morphological properties and stability of AgNPs. AgNPCB and AgNPQB varied in size and shape (spherical, quadrangular, triangular, polygonal, and rods). Remarkably, biosynthesis using the algal powder produced smaller particles relative to those formed using the extract, albeit slowly. There was a noticeable inconsistency in the size of AgNPs in both DLS and TEM. To illustrate, DLS measures the hydrodynamic size of the particles involving its capping phytochemicals from the algal extract, while TEM measures the exact geometric size of NPs [20,21]. Despite these inconsistencies, our findings are mostly consistent with those of other researchers and are deemed acceptable, based on the assumption that the discrepancies are due to technical variability associated with the different equipment used rather than measurement errors. EDS verified the presence of elemental silver in all nanosolutions. The optical absorbance at 3 keV is attributed to plasmon resonance of the metallic silver nanocrystals and is known as the Ag region [22]. This result is consistent with that of Shaik et al. [23], which used Salvadora persica L. root extract (Miswak) in the green synthesis of AgNPs. FTIR was used to confirm the involvement of the algal functional groups in biomolecules during the reduction reaction [24]. The extracts were tested before and after the formation of AgNPs to compare the occurring shift in the resulting peaks. Use of metal salts to synthesize NPs requires a stabilizer against the van der Waals forces of attraction to avoid coagulation [25]. FTIR results accentuate the contribution of the hydroxyl group (O–H bond) at 3400.38 and 3427.22 cm−1 after AgNPCB and AgNPQB synthesis, respectively. The hydroxyl group particularly has been proven to reduce the metal ions of silver to its atom “nano” form and conduct stability for formed NPs, hence its ability to increase oxygen bonding. These findings are compatible with those observed in earlier studies [26,27]. Hydroclathrus clathratus has been used as an antitumor, antioxidant, and antibacterial agent [28–30]; yet, to the best of our knowledge, there are no published studies on the biosynthesis of NPs from this algal species. Algae are considered as effective bionanofactories for synthesizing metallic NPs; hence, they are abundantly available, and both dead and live biomass can be successfully used in the production of metallic NPs [31]. There are two acceptable antibacterial actions of AgNPs: contact killing by infiltrating bacterial cells and Ag+ ion-mediated killing by generating reactive oxygen species (ROS) [32]. Oves et al. [33] used the fluorescent probe 2,7-dichlorofluorescein diacetate dyes to detect ROS production by AgNPs inside the bacterial cells. Results showed that free radicals’ production in the media was associated with increased concentrations of AgNPs and incubation time. Many studies have demonstrated that bio-AgNP activity is concentrationdependent. More recent evidence proposes that bacterial aggregation and physiology are essential determinants that define the predominance of one or several of the proposed mechanisms for the AgNPs’ antibacterial activity [34]. Moreover, studies have demonstrated that the relatively large surface area of smaller AgNPs facilitates the release of more silver ions, which penetrate the bacterial cell membrane leading to its death [35,36]. The in vitro biological tests against bacteria exhibited varied responses to the AgNPs, which were expressed by the diameter of the inhibition zone. AgNPs synthesized by algal methanol extracts were more effective, compared to the raw powder aqueous solution against bacteria. Remarkably, A. baumannii and S. aureus, both the sensitive and resistant strains, were the most affected by the algal methanol extracts, compared to other bacteria. Alavi et al. [37] studied the antibacterial activities of Ag, Cu, TiO2, ZnO, and Fe3O4 NPs biologically synthesized using Protoparmeliopsis muralis lichen aqueous extract against MRSA, E. coli, and P. aeruginosa. The highest antibacterial activity was noticed with 0.1 M concentration of AgNPs against P. aeruginosa, MRSA, and E. coli, respectively. In contrast, AgNPQB was more effective against nonresistant S. aureus, A. baumannii, and P. aeruginosa. Studies have shown that Gram-positive bacteria are more susceptible to AgNPs, compared to Gramnegative bacteria due to their structural differences. Contradicting earlier findings [38,39], we found no biased antibacterial action against either Gram-positive or Gramnegative bacteria, which could be attributed to the charge difference between the AgNPs and bacterial cells [32]. Previous research work revealed the difficulty in biofilm growth and development inhibition, compared to cell attachment inhibition [15]. In this study, biofilm growth and development inhibition assay were used to evaluate the antibiofilm activity of AgNPs. We tested the antibiofilm Bio-AgNPs and their biological activity activity of AgNPCB against selected biofilm-forming bacteria in two concentrations. There were no notable differences between tested concentrations, and they profoundly inhibited the biofilm of MRSA with 99% inhibition. A study investigated the antibiofilm effect of bio-AgNPs fabricated using the Artemisia scoparia plant as a bioreductant and compared it to that of commercial AgNPs against 50 strains of S. aureus. They assessed this effect on bacterial biofilm at a molecular level, specifically on icaABCD genes, which are essential for biofilm formation. The results registered a more notable reduction and induction in icaA and icaR gene expression with the sub-MIC doses of biosynthetic AgNP contrasted to commercial AgNP [40]. Rolim et al. [41] synthesized AgNPs using the fungus Stereum hirsutum and two plant extracts (green tea and dill) and studied their antibiofilm activity against several bacterial strains of medical interest. MDR P. aeruginosa was highly susceptible to the AgNPs synthesized using S. hirsutum with 97% inhibition. Our findings, while dissatisfactory, further implicate the cost of bacterial antibiotics’ resistance in its fitness and virulence in MRSA specifically. A study suggested using nanotechnology to coat surgical devices and medical implants to control biofilm formation because NPs are capable of breaching the extracellular polymeric substance layer and the bacterial membrane of both Gram-positive and Gram-negative bacteria [42]. Even though we did not find this association with all tested bacteria, there are still controversies on whether the antibiotics’ resistance to bacteria contributes positively or negatively to biofilm formation [43]. We recommend testing the effect of AgNPCB against MDR bacteria and on different stages of biofilm formation to comprehend their antibiofilm activity fully in the future. 5 Conclusions This study has explored the capacity of the brown alga H. clathratus to produce NPs extracellularly following a rapid and biological approach. Different techniques were used to characterize the produced AgNPs and obtain a thorough background for the future application of formed AgNPs. Our research work has further proven the activity of biosynthesized AgNPs as antibacterial and antibiofilm agents with room for improvement. We also introduced many questions in need of further investigation regarding the consequences of drug resistance on bacterial fitness and bacterial biofilm formation, which may help in developing antibacterial/antibiofilm drugs and improving the fields of biotechnology and health altogether.  423 Acknowledgments: The authors would like to thank the Deanship of Scientific Research in King Saud University for funding and supporting this research work through the initiative of DSR Graduate Students Research Support (GSR). The authors would also like to acknowledge the help and advice provided by Prof. Nawal M. Almusayeib and Prof. Musarat Amina, from King Saud University, and Prof. Manal Awad, from King Abdullah Institute for Nanotechnology. Also, the authors thank Dr. Aref Elmubarak from the Plant Protection Department at King Saud University for performing the phytochemical screening of the algal samples. Author contributions: M. M. A. and N. M. A. prepared the research design and supervision; R. R. A. performed experiments and wrote the manuscript. References Baptista PV, McCusker MP, Carvalho A, Ferreira DA, Mohan NM, Daniela AF, et al. Nano-strategies to fight multidrug resistant bacteria—“a battle of the titans”. Front Microbiol. 2018;9:1441. [2] Al-Hajoj S, Varghese B, Shoukri MM, Al-Omari R, AlHerbwai M, AlRabiah F, et al. Epidemiology of antituberculosis drug resistance in Saudi Arabia: findings of the First National Survey. Antimicrob Agents Chemother. 2013;57(5):2161–66. [3] Zowawi HM, Balkhy HH, Walsh TR, Paterson DL. β-Lactamase production in key Gram-negative pathogen isolates from the Arabian Peninsula. Clin Microbiol Rev. 2013;26(3):361–80. [4] Hoiby N, Bjarnsholt T, Givskov M, Molin S, Ciofu O. Antibiotic resistance of bacterial biofilms. Int J Antimicrob Agents. 2010;35(4):322–32. [5] Shannon E, Abu-Ghannam N. Antibacterial derivatives of marine algae: an overview of pharmacological mechanisms and applications. Mar Drugs. 2016;14(4):81. [6] Khan AR, Qari R. Antibacterial activities of brown seaweed Sargassum boveanum (J. Ag.) against diarrhea along the Coast of Karachi, Pakistan. J Environ Res Dev. 2012;6(3A):753–7. [7] Torode TA, Marcus SE, Jam M, Tonon T, Blackburn RS, Herve C, et al. Monoclonal antibodies directed to fucoidan preparations from brown algae. PLoS One. 2015;10(2):e0118366. [8] Pereira L. Seaweed flora of the European North Atlantic and Mediterranean. In: Kim SK, editor. Springer Handbook of Marine Biotechnology. Berlin: Springer Berlin Heidelberg; 2015. [9] El-Sheekh MM, El-Kassas HY. Algal production of nano-silver and gold: Their antimicrobial and cytotoxic activities: a review. J Genet Eng Biotechnol. 2016;14(2):299–310. [10] Davis TA, Volesky B, Mucci A. A review of the biochemistry of heavy metal biosorption by brown algae. Water Res. 2003;37(18):4311–30. [11] Aleem AA. The marine algae of Alexandria. Alexandria: Egypt. Univ. Alexandria; 1993. [1] 424  Raghad R. Alzahrani et al. [12] Coppejans E, Leliaert F, Dargent O, Gunasekara R, De Clerck O. Sri Lankan Seaweeds – Methodologies and field guide to the dominant species. Brussels: Belgian Development Cooperation; 2009. [13] Rajeshkumar S, Kannan C, Annadurai G. Green synthesis of silver nanoparticles using marine brown algae Turbinaria conoides and its antibacterial activity. Int J Pharm Bio Sci. 2012;3(4):502–10. [14] Xie JL, Singh-Babak SD, Cowen LE. Minimum inhibitory concentration (MIC) assay for antifungal drugs. Bio Protoc. 2012;2(20):e252. [15] Bazargani MM, Rohloff J. Antibiofilm activity of essential oils and plant extracts against Staphylococcus aureus and Escherichia coli biofilms. Food Control. 2016;61:156–64. [16] Karimi E, Jaafar HZ, Ghasemzadeh A, Ebrahimi M. Fatty acid composition, antioxidant and antibacterial properties of the microwave aqueous extract of three varieties of Labisia pumila Benth. Biol Res. 2015;48(1):9. [17] Rajoriya P. Green synthesis of silver nanoparticles, their characterization and antimicrobial potential [PhD thesis]. Sam Higginbottom University of Agriculture Technology and Sciences, India, 2017. [18] Patra JK, Baek KH. Antibacterial activity and synergistic antibacterial potential of biosynthesized silver nanoparticles against foodborne pathogenic bacteria along with its anticandidal and antioxidant effects. Front Microbiol. 2017;8:167. [19] Khalifa KS, Ragaa AH, Hanafy AH. Antitumor activity of silver nanoparticles biosynthesized by micro algae. J Chem Pharm Res. 2016;8(3):1–6. [20] Tomaszewska E, Soliwoda K, Kadziola K, Tkacz-Szczesna B, Celichowski G, Cichomski M, et al. Detection limits of DLS and UV-Vis spectroscopy in characterization of polydisperse nanoparticles colloids. J Nanomater. 2013;2013:10. [21] Chartarrayawadee W, Charoensin P, Saenma J, Rin T, Khamai P, Nasomjai P, et al. Green synthesis and stabilization of silver nanoparticles using Lysimachia foenum-graecum Hance extract and their antibacterial activity. Green Proc Synth. 2020;9(1):107–18. [22] Fayaz AM, Balaji K, Girilal M, Yadav R, Kalaichelvan PT, Venketesan R. Biogenic synthesis of silver nanoparticles and their synergistic effect with antibiotics: a study against Grampositive and Gram-negative bacteria. Nanomed Nanotechnol Biol Med. 2010;6(1):103–9. [23] Shaik M, Albalawi GH, Khan ST, Khan M, Adil SF, Kuniyil M, et al. “Miswak” based green synthesis of silver nanoparticles: evaluation and comparison of their microbicidal activities with the chemical synthesis. Molecules. 2016;21(11):1478. [24] Faghihzadeh F, Anaya NM, Schifman LA, Oyanedel-Craver V. Fourier transform infrared spectroscopy to assess molecularlevel changes in microorganisms exposed to nanoparticles. Nanotechnol Environ Eng. 2016;1(1):1. [25] Sivagnanam SP, Getachew AT, Choi JH, Park YB, Woo HC, Chun BS. Green synthesis of silver nanoparticles from deoiled brown algal extract via Box-Behnken based design and their antimicrobial and sensing properties. Green Proc Synth. 2017;6(2):147–60. [26] Wang Z, Xu C, Zhao M, Zhao C. One-pot synthesis of narrowly distributed silver nanoparticles using phenolic-hydroxyl [27] [28] [29] [30] [31] [32] [33] [34] [35] [36] [37] [38] [39] [40] modified chitosan and their antimicrobial activity. RSC Adv. 2014;4(87):47021–30. Guerrini L, Alvarez-Puebla RA, Pazos-Perez N. Surface modifications of nanoparticles for stability in biological fluids. Materials. 2018;11(7):1154. Vimala T, Poonghuzhali T. In vitro antimicrobial activity of solvent extracts of marine brown alga, Hydroclathrus clathratus (C. Agardh) M. Howe from Gulf of Mannar. J Appl Pharm Sci. 2017;7(04):157–62. Kelman D, Posner EK, McDermid KJ, Tabandera NK, Wright PR, Wright AD. Antioxidant activity of Hawaiian marine algae. Mar Drugs. 2012;10(2):403–16. Wang H, Chiu LC, Ooi VE, Ang PO. A potent antitumor polysaccharide from the edible brown seaweed Hydroclathrus clathratus. Bot Mar. 2010;53(3):265–74. LewisOscar F, Vismaya S, Arunkumar M, Thajuddin N, Dhanasekaran D, Nithya C. Algal nanoparticles: synthesis and biotechnological potentials. Algae-organisms for imminent. Biotechnology. 2016;7:157–82. Yun’an Qing LC, Li R, Liu G, Zhang Y, Tang X, Wang J, et al. Potential antibacterial mechanism of silver nanoparticles and the optimization of orthopedic implants by advanced modification technologies. Int J Nanomed. 2018;13:3311. Oves M, Rauf MA, Hussain A, Qari HA, Khan AP, Muhammad P, et al. Antibacterial silver nanomaterial synthesis from Mesoflavibacter zeaxanthinifaciens and targeting biofilm formation. Front Pharmacol. 2019;10:801. Rodriguez-Serrano C, Guzman-Moreno J, Angeles-Chavez C, Rodriguez-Gonzalez V, Ortega-Sigala JJ, Ramirez-Santoyo RM, et al. Biosynthesis of silver nanoparticles by Fusarium scirpi and its potential as antimicrobial agent against uropathogenic Escherichia coli biofilms. PLoS One. 2020;15(3):e0230275. Chowdhury S, Basu A, Kundu S. Green synthesis of protein capped silver nanoparticles from phytopathogenic fungus Macrophomina phaseolina (Tassi) Goid with antimicrobial properties against multidrug-resistant bacteria. Nanoscale Res Lett. 2014;9(1):1–11. Raza MA, Kanwal Z, Rauf A, Sabri AN, Riaz S, Naseem S. Sizeand shape-dependent antibacterial studies of silver nanoparticles synthesized by wet chemical routes. Nanomaterials. 2016;6(4):74. Alavi M, Karimi N, Valadbeigi T. Antibacterial, antibiofilm, antiquorum sensing, antimotility, and antioxidant activities of green fabricated Ag, Cu, TiO2, ZnO, and Fe3O4 NPs via protoparmeliopsis muralis lichen aqueous extract against multi-drug-resistant bacteria. ACS Biomater Sci Eng. 2019;5(9):4228–43. Chatterjee T, Chatterjee BK, Majumdar D, Chakrabarti P. Antibacterial effect of silver nanoparticles and the modeling of bacterial growth kinetics using a modified Gompertz model. BBA-Gen Subjects. 2015;1850(2):299–306. Al-Sharqi A, Apun K, Vincent M, Kanakaraju D, Bilung LM. Enhancement of the antibacterial efficiency of silver nanoparticles against Gram-positive and Gram-negative bacteria using blue laser light. Int J Photoenergy. 2019;2019:12. Moulavi P, Noorbazargan H, Dolatabadi A, Foroohimanjili F, Tavakoli Z, Mirzazadeh S, et al. Antibiofilm effect of green Bio-AgNPs and their biological activity engineered silver nanoparticles fabricated from Artemisia scoporia extract on the expression of icaA and icaR genes against multidrug-resistant Staphylococcus aureus. J Basic Microbiol. 2019;59(7):701–12. [41] Rolim WR, Lamilla C, Pieretti JC, Díaz M, Tortella GR, Diez MC, et al. Comparison of antibacterial and antibiofilm activities of biologically synthesized silver nanoparticles against several bacterial strains of medical interest. Energy Ecol Environ. 2019;4(4):143–59.  425 [42] LewisOscar F, MubarakAli D, Nithya C, Priyanka R, Gopinath V, Alharbi NS, et al. One pot synthesis and anti-biofilm potential of copper nanoparticles (CuNPs) against clinical strains of Pseudomonas aeruginosa. Biofouling. 2015;31(4): 379–91. [43] Cepas V, López Y, Munoz E, Rolo D, Ardanuy C, Martí S, et al. Relationship between biofilm formation and antimicrobial resistance in Gram-negative bacteria. Microb Drug Resist. 2019;25(1):72–9. 426  Raghad R. Alzahrani et al. Appendix Table A1: Phytochemicals identified in the MeOH extract and powder of H. clathratus RT Compound name Phytocomponents in the MeOH extract Saturated fatty acids 24.135 n-Tridecanoic acid 25.885 Tetradecanoic acid 27.115/29.185 Hexadecanoic acid 27.48 n-Pentadecanoic acid 28.64/28.905 Palmitelaidic acid 28.74 cis-9-Hexadecenoic acid 31.98 Octadecanoic acid 34.68 Eicosanoic acid Unsaturated fatty acids 29.905 9-Octadecenoic acid 30.17 cis-10-Heptadecenoic acid 31.555 9,12-Octadecadienoic acid (Z,Z)31.695 Oleic acid 33.78 Arachidonic acid 33.88 cis-5,8,11,14,17-Eicosapentaenoic acid 34.17 9-Decenoic acid Sugars 27.055 Galactopyranose, 1,2,3,4,6-pentakis-O-(trimethylsilyl)-,.beta.-d27.655 1,5-Anhydro-D-sorbitol 33.54 Glyceryl-glycoside 38.615 2-Monostearin Carboxylic acids (derivatives) 6.38 Formamide, N,N-diethylSteroids 14.715 Glycerol Others 25.015 Borneol 25.655/26.38 3,7,11,15-Tetramethyl-2-hexadecen-1-ol 36.695 Hexadecanoic acid, 2,3-bis[(trimethylsilyl)oxy]propyl ester 39.02 Octadecanoic acid, 2,3-bis[(trimethylsilyl)oxy]propyl ester RT Compound name Phytocomponents in the powder of H. clathratus Saturated fatty acids 27.47 n-Pentadecanoic acid 27.71 Trimethylsilyl ether of glucitol 29.05 Hexadecanoic acid 31.965 Octadecanoic acid 34.67 Eicosanoic acid Unsaturated fatty acids 28.625 n-Tridecanoic acid 28.725 cis-9-Hexadecenoic acid 28.885 Palmitelaidic acid 30.16 cis-10-Heptadecenoic acid 31.54 9,12-Octadecadienoic acid (Z,Z)31.61/31.705 Oleic acid 33.765 Arachidonic acid 33.87 cis-5,8,11,14,17-Eicosapentaenoic acid 35.4 Oleanitrile acid; oleic acid with ammonia Synonym Tridecanoic acid Myristic acid Palmitic acid NS 9-Hexadecenoic acid Palmitoleic acid Stearic acid Arachidic acid Elaidic acid 10Z-Heptadecenoic acid Linoleic acid Omega 9 cis-5,8,11,14-Eicosatetraenoic acid Eicosapentaenoic acid Caproleic acid Galactopyranose Glucitol; (sugar alcohol) NS 2-Stearoylglycerol Formamide (Glycerin) (−)-Borneol (bicyclic monoterpenoids) Phytol NS NS Synonym NS Glucitol; sorbitol (sugar alcohol) Palmitic acid Stearic acid Arachidonic acid Tridecanoic acid Palmitoleic acid 9-Hexadecenoic acid 10Z-Heptadecenoic acid Linoleic acid Omega 9 cis-5,8,11,14-Eicosatetraenoic acid Eicosapentaenoic acid NS Bio-AgNPs and their biological activity Table A1: Continued RT Sugars 38.605 39.38 Carboxylic acid 39.525 Steroids 45.855 Others 7.125 25.645/26.37 36.69 38.705 Compound name Phytocomponents in the powder of H. clathratus Synonym 2-Monostearin 2-Stearoylglycerol N-Acetyl-D-glucosamine, tetrakis(trimethylsilyl) ether, benzyloxime (isomer 1) NS Tetracosanoic acid Lignoceric acid 24-Nor-22,23-methylenecholest-5-en-3.beta.-ol (Sterol) 2,3,3-Trimethyl-, 1-butene 3,7,11,15-Tetramethyl-2-hexadecen-1-ol Hexadecanoic acid, 2,3-bis[(trimethylsilyl)oxy]propyl ester Eicosanoic acid, 2,3-bis[(trimethylsilyl)oxy]propyl ester Alkene Phytol NS NS RT – retention time. NS – no synonyms. Synonyms were obtained via NIST, PubChem, ChEBI, and HMDB.  427