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Evidence for the antifouling potentials of marine macroalgae Sargassum wightii.


Biofouling can be defined as the undesirable phenomenon of adherence and accumulation of biotic deposits on a submerged artificial surface or in contact with seawater (Eguia and Trueba, 2007). Biofouling on ship hull is being a problem for decade and probably for as long as mankind has been sailing the oceans. In general, the process of biofouling on a surface consist of four essential stages which begins with (i) macromolecular adsorption; (ii) bacterial colonization; (iii) surface fouling by diatoms and protozoans and (iv) establishment of unicellular and multicellular epibionts such as invertebrate larvae and algal propagules (Qian et al., 2006). Formation of these complex layer of assemblages of marine organisms are not confined only to ship hulls and ship machineries, but also affects other shore line based installations such as coastal power stations, chemical and desalination plants which use seawater as cooling medium (Swami and Udhyakumar, 2010). It was estimated that marine biofouling on ship hulls augment fuel consumption, hull drag and consequently cleaning and painting, resulting in estimated costs of one billion dollars per year for the US Navy alone (Callow and Callow, 2002).

Among the different coatings tried throughout the maritime history, tributytlin self-polishing copolymer paints (TBT-SPC paints) have been the most successful in combating biofouling, however it was proved to have adverse effects on non-target marine organisms. Considering the threat posed by TBT, the International Maritime Organization (IMO) has proposed phasing out of TBT based antifouling paints on the surface of vessels from January 1, 2008 (Yebra et al., 2004). To assail this environmental impact, intensive research is being driven now throughout the world to develop environmentally friendly antifouling technologies as a compatible substitute to toxic biocides.

Marine natural products or extracts with antifouling activities are one of the most excellent alternative techniques to TBT, which have been isolated from wide number of marine organisms. The key chemical antifouling mechanism of marine organisms occurs via the production of secondary metabolites (also known as natural products) which deter foulers (Chambers et al., 2006). Seaweeds are one of the most prolific natural product synthesizers from which numerous bioactive metabolites exhibiting wide spectrum of bioactivity have been isolated. Preliminary studies of secondary metabolites from seaweeds have shown biological activity against bacteria (Sieburth and Conover, 1965), Mytilus edulis larvae (Katsuoka et al., 1990), bryozoan (Bugula neritina) larvae (Schmitt et al., 1995) and Balanus amphitrite larvae (Willemsen, 1994). These secondary metabolites are presumably potential antifoulants and numerous active antifouling compounds have been obtained so far from marine algae. Though antifouling compounds were reported from many seaweeds, the genus Sargassum (Phaeophyceae) is well known to produce compounds with antibacterial, antitumoral, antimalarial, antiherbivory and antifouling properties (Amsler and Fairhead, 2006; Afolayan et al., 2008; Plouguerne et al., 2010). For instance, most recently a lipidic metabolite isolated from S. muticum was reported to exhibit predominant antifouling activity (Plouguerne et al., 2010). Considering the above importance, the present study was undertaken to determine the substratum wise influence on settlement of biofilm bacteria of Chinnamuttom fisheries harbour, Southeast coast of Tamilnadu, India and also to assess the antifouling efficacy of secondary metabolites from seaweed Sargassum wightii.

Materials and methods

Description of the study:

Four different panels such as wood of Artocarpus hirsutus, fiber glass reinforced plastic (FRP), stainless steel (316L) and carbon steel which are widely used to construct ship hulls, boats and other structures were used for the present study. All the panels were of same size (15 cm length x 10 cm breadth x 2 mm width) and were exposed to a depth of 2 m at Chinnamuttom fisheries harbour (Lat. 8[degrees]05'44.50"N; Long. 77[degrees]33'44.20"E), South east coast of India. The panels were assessed every 24 h upto 72 h and the biofilms were swabbed using sterile cotton swabs (Wahl et al., 1994) and transported to the laboratory in aseptic condition. Simultaneously, water samples were also collected from the study area to determine the physicochemical properties of water. Water temperature was recorded using a thermometer (Hermes India; sensitivity [+ or -] 1[degrees]C). Salinity and pH of the water samples were recorded using a salinity refractometer (News-100, Tanaka Sanjiro Co., Ltd., Japan; Practical Salinity Unit (psu)) and a pH meter (Elico, India). Other chemical parameters, such as dissolved oxygen, silicate, nitrate, total phosphorous, phosphate and ammonia were analysed based on APHA (1985).

Isolation of biofilm bacteria:

The collected swabs were transferred to test tubes individually containing 10 ml of sterile seawater. From each tube, 1ml of the sample was taken and serially diluted upto [10.sup.-5] dilution. 0.1 ml of each dilution was then plated on Petridish containing Zobell Marine agar (2216) and incubated at 37[degrees]C for 48 h. The mean total bacterial population was estimated and recorded in each plate. Pure bacterial colonies were isolated and identified upto species level by Bergey's Manual of Determinative Bacteriology (Holt et al., 1994). The individual biofilm colonies were restreaked in Zobell Marine Agar and kept as stock.

Collection of seaweed and extract preparation:

Seaweed S. wightii was collected freshly from Kanyakumari coast (Lat. 8[degrees]05'05.89"N; Long. 77[degrees]32'29.98"E), Kanyakumari District, Tamilnadu, India. The collected seaweed was washed thoroughly in running water to remove the soil and extraneous dirt and shade dried completely. Then the dried seaweed was powdered and subjected to percolation by soaking individually in solvents of varying polarity viz. methanol, chloroform and hexane for one week. Extracts were then filtered with Whatman No.1 filter paper, evaporated and concentrated under rotary vacuum evaporator. The crude extracts obtained were used for antifouling assays.

Antimicrofouling activity of seaweed extract:

The biofilm bacterial colonies were seeded individually in Muller Hinton agar plates. The antibacterial assay was determined using the standard disc diffusion method (Kirby Bauer method--Bauer et al., 1966). Sterile Whatman No.1 filter paper discs of 5mm diameter were impregnated with 0.5 mg/disc of the crude extracts individually, air dried and placed on Muller Hinton agar plates previously seeded with test bacterium and incubated for 24 h at 37[degrees]C. The assay was carried out in triplicate. The clear zone of inhibition was measured in mm.

Mussel Bioassay:

Based on the result of antibacterial assay, the most predominant result yielding methanolic extract of seaweed S. wightii was selected for mussel bioassay by following the method described by Wilsanand et al. (1999) and Murugan and Santhana Ramasamy (2003). To carryout this study, the brown mussel Perna indica were collected from Manavalakurichi coast (Lat. 8[degrees]08'49.72"N; Long. 77[degrees]18'08.23"E), Kanyakumari, Tamilnadu, India and transported to the laboratory and acclimatized in laboratory for 24 h. Uniform size group of (1.5 to 2 cm length) of mussels were used as test animal and their attachment with the help of byssus thread was used as toxic criterion. Aged, sterilized and filtered seawater was used for the experiment. The crude extract was weighed, dissolved in distilled water. Ten mussels were introduced into each beaker containing 100 ml of seawater with various concentrations of extracts ranged from 50 to 400 [micro]g [ml.sup.-1] at an interval of 50 [micro]g. Control group was also maintained separately without extract. In all the experimental setup, mild aeration was provided. The experiment was carried out in triplicate. The mussels were not fed during the experiment. After 24 h, the settlement of mussels at various concentrations was recorded and minimum concentration which prevented byssal production and attachment was denoted as [EC.sub.50] (effective concentration at which 50% of mussels showed inhibition of byssal attachment). Then the [LC.sub.50] value was estimated through probit analysis after exposing the same in test concentrations of the extract for 96 h (Wilsanand et al., 1999).

Anticrustacean assay:

Anticrustacean assay is one of the simple and inexpensive screening techniques to identify the cytotoxicity of the bioactive compounds. In the present study anticrustacean assay was carried out by the method described by Ortega-Morales et al. (2008) with Artemia salina as the model organism to investigate the cytotoxicity of the crude methanolic extract of S. wightii. For this cysts of brine shrimp (A. salina) were hatched in conical shaped vessel (1L), filled with filtered seawater under constant aeration for 24 h. After hatching, active nauplii, (I instar), were collected from brighter portion of the hatching chamber and used for the assay. In order to carry out the cytotoxicity assay ten nauplii were drawn through a thin capillary glass tube and placed in small test tube containing 10 ml of brine solution at various concentrations (30-240 [micro]g [ml.sup.-1]) of crude methanolic extract of S. wightii and maintained at room temperature for 24 h under the light. The assay was carried out in triplicate. After 24 h of exposure the number of larvae surviving in each test concentrations was counted. Lethal concentrations of 50% ([LC.sub.50]) values were determined by counting the dead nauplii after an incubation period of 24h.

Phytochemical and FTIR analysis of active seaweed extract:

The prominent result rendering extract was screened for the presence of phytochemical constituents by following the method of Sofowora (1982) and Kepam (1986). To analyze the functional groups of the prominent result showing crude extract of seaweed, FTIR spectrum was accomplished in the frequency range of 4000 to 400 [cm.sup.-1] (Shimadzu FTIR--820 IPC).

Statistical analysis:

The results obtained in the present study were subjected to two way ANOVA test and Probit analysis through SPSS 13.0 software.


Physicochemical parameters of water sample:

During the three days of the study, the physicochemical parameters of seawater of Chinnamuttom fisheries harbour showed a meager fluctuation (Table 1). The salinity of the water fluctuated between 35 and 35.5. Similarly the levels of temperature, Dissolved oxygen (DO) and pH did not vary significantly (27-28[degrees]C, 5.2-5.4 [mg.sup.-1], DO and 8-8.2 pH). Other parameters such as silicate (0.149-0.155 [micro]/mol), nitrate (2.235-2.242 [micro]/mol), nitrite (0.319-0.332 [micro]/mol), total phosphorous (18.9-19.5 [micro]/mol), phosphate (15.90-16.26 [micro]/mol) and ammonia (0.022-0.026 [micro]/mol) showed relatively less variation.

Biofilm bacterial density on experimental panels:

The variation in Total Viable Count (TVC) of biofilm bacteria of the panels during 24, 48 and 72h of exposure in Chinnamuttom fisheries harbour water is given in Fig. 1. The TVC of biofilm was found to increase with increase in time intervals in all the tested panels. For instance, the TVC of biofilm samples of the wooden panel exposed to seawater during 24 h was 153.33 [+ or -] 6.94 x [10.sup.-5] CFU [ml.sup.-1]. This bacterial population gradually increased (235 [+ or -] 7.35 and 265 [+ or -] 6.16 x [10.sup.-5] CFU [ml.sup.-1]) during subsequent sampling intervals of 48 and 72 h, respectively.

The bacterial density on the stainless steel panel exposed to seawater during 24 h was 74.8 [+ or -] 4.59 x [10.sup.-5] CFU [ml.sup.-1]. This density increased to 118 [+ or -] 6.80 and 169 [+ or -] 4.90 x [10.sup.-5] CFU [ml.sup.-1] during 48 and 72 h, respectively. Similarly the bacterial population on FRP panel was 69.7 [+ or -] 4.92, 94 [+ or -] 3.27, 164 [+ or -] 6.16 x [10.sup.- 5] CFU [ml.sup.-1] during the time intervals of 24, 48 and 72 h, respectively. The bacterial density on the carbon steel panel also showed an increasing trend, but did not show much variation and it ranged between 32.6 [+ or -] 1.68 to 46 [+ or -] 2.72 x [10.sup.-5] CFU [ml.sup.-1] respectively. The bacterial density in the tested substrata was in the following order: wood> stainless steel>FRP>carbon steel. The statistical (Two-way ANOVA) analysis on bacterial density in the substrata revealed that the variation between the panels (F = 25.08; P<0.001) as well as between the time duration (F = 9.79; P<0.05) were statistically significant.


Biofilm bacterial diversity on tested panels:

From the tested substrata, ten bacterial strains were isolated and were subjected to gram's staining, motility test as well as series of biochemical and physiological tests for species level identification (Table 2). The identified strains were Pseudomonas aeruginosa, Halomonas aquamarina, Vibrio alginolyticus V. fischeri, V. parahaemolyticus, Enterobacter agglomerans, Serratia marcescens, S. liquefaciens, Shigella flexneri, and Aeromonas hydrophila. The diversity of bacterial strains during 24, 48 and 72 h were pooled together and produced as mean value. Among the identified biofilm bacterial strains P. aeruginosa was the predominant species recorded in all the tested panels which ranged from 34 [+ or -] 1.63 to 39 [+ or -] 2.45%, followed by H. aquamarina (20 [+ or -] 0.82 to 24 [+ or -] 2.16%), and V. alginolyticus (14 [+ or -] 0.82 to 21 [+ or -] 1.24%). The occurrence of E. agglomerans was maximum (9 [+ or -] 0.82%) in wooden panel but minimum (5 [+ or -] 0.82 and 5 [+ or -] 1.63%) in FRP and carbon steel panels. V. fischeri was more abundant in wooden and FRP panels (6 [+ or -] 1.63 and 6 [+ or -] 0.81%) than in other tested panels (3-4%). S. liquefaciencs was recorded with only 2 [+ or -] 0.82 and 3 [+ or -] 0.82% in FRP and Stainless steel panels, respectively, but it was not identified in wood and carbon steel panels. Likewise V. parahaemolyticus registered maximum of 3 [+ or -] 0.82% in FRP panel and 2 [+ or -] 0.82% in stainless steel and carbon steel panels, but not observed in wooden panel. S. flexneri and A. hydrophila were found in smaller proportions (1-3%) in all the tested panels.

Antimicrofouling activity of S. wightii extracts:

In the present study, the antibacterial activity of different solvent based crude extracts of seaweed was tested against all the isolated biofilm bacterial strains. Among the crude extracts tested, methanolic extract showed 100% growth inhibitory activity against all the tested biofilm bacterial strains with maximum zone of inhibition of 13mm against two biofilm bacterial strains such as P. aeruginosa and S. liquefaciens and minimum of 9 [+ or -] 0.23 mm against H. aquamarina. Chloroform extract exerted better level of 90% bioactivity against biofilm bacterial strains with maximum zone of inhibition against S. liquefaciens (12 [+ or -] 0.40 mm) and minimum level of zone of inhibition against V. fischeri (6 [+ or -] 0.11 mm). Hexane extract displayed lesser level of growth inhibitory activity of 6.5 [+ or -] 0.44 to 8.5 [+ or -] 0.44 mm against all the tested biofilm bacterial strains except against H. aquamarina and A. hydrophila (Table 3).

Antimacrofouling activity:

The effective concentration ([EC.sub.50]) of methanolic extract for 50% inhibition of byssal production and attachment of brown mussel P. indica after 24 h of exposure was recorded as 205 [+ or -] 14.7 [micro]g [ml.sup.-1]. The same mussels subjected to toxic criterion study inferred that after 96 h of exposure, 50% lethal concentration ([LC.sub.50]) value estimated and recorded through probit analysis was 306 [+ or -] 19.6 [micro]g [ml.sup.-1]. Whereas in the control group, normal byssus production and attachment was noticed (Table 4).

Anticrustacean assay:

Anticrustacean assay is one of the most reliable assay methodology to test the toxic criterion of a chemical/extract. In the present study Artemia salina (nauplii) was used as model organism to test the toxicity of methanolic extract of S. wightii. From the result, the [LC.sub.50] value for crude methanolic extract of S. wightii was recorded as 161 [+ or -] 4.64 [micro]g [ml.sup.-1] (Table 5).

Phytochemical and FTIR analysis of methanolic extract of S. wightii:

Analysis of crude methanolic extract of S. wightii displayed the presence of phytochemicals such as phenols, coumarins, quinones, saponins, alkaloids and flavonoids (Table 6). Also FTIR analysis revealed the presence of functional groups such as Iodine-halogen by recording its peak signal at 370.31, 397.31 and 426.24 [cm.sup.-1], bromine (659.61[cm.sup.-1]), fluorine (1012.56 and 1026.06 [cm.sup.-1]), and carbon-nitrogen covalent bonding (1112.85 [cm.sup.-1]), carbon-carbon covalent bonding (1407.94 and 2040.55 [cm.sup.-1]), nitro group (1452.30, 1652.88 and 2148.55 [cm.sup.-1]), hydroxyl group (2520.79 and 2948.96 [cm.sup.-1]) based and carbon-hydrogen covalent bonding (2837.09 and 3303.83 [cm.sup.-1]) on the vibration stretches (Fig. 2).


The competition for living space is more intense in marine environment; hence all submerged surfaces in the marine environment are rapidly colonized by bacteria and they form the important component in the development of a fouling community (Mitchell and Kirchman, 1984). In the present study, four different panels (substrata) were exposed to seawater so as to assess the influence of substrata on biofilm bacteria recruitment. During the experimental period, the microbial load (TVC) increased gradually i.e. from 153.3 to 264; 75.6 to 169; 69.33 to 163 and 32.67 to 46 x [10.sup.5] CFU [ml.sup.-1] from 24 to 72 h intervals in the wooden, stainless steel, FRP and carbon steel panels, respectively. The observed increase in the microfouling biomass was due to enhanced settlement and growth of already colonized microbes on the panel surface (Immanuel et al., 2005).

Surfaces in contact with seawater medium are rapidly colonized by bacteria due to ease access to nutrients, protection against antibiotics, maintenance of extracellular enzyme activities and shelter for predation (Dang and Lovell, 2000). In the present study, out of four substrata used, wooden panel showed the highest bacterial load, whereas the lowest bacterial population was observed on the carbon steel panel. The highest population on the wooden panel may be due to its surface nature. It has been reported that rough surfaces are more favourable for the settlement of microfoulers than the smooth surfaces (Anderson and Underwood, 1994). The results of the present study thus suggested that rough surface of wooden panel (substratum) had greatly influenced the colonization of microbiota. Next to wooden panel, stainless steel panel displayed higher rate of succession of bacterial population than that of FRP panel with respect to time interval, however the difference was not too high. The observed variation in the succession primarily reflects the influence of substrata. In agreement with the present result, Fletcher and Marshall (1982) stated that the pattern of colonization of the submerged surfaces by bacteria is influenced by the physical and chemical characteristics of the surface. Therefore in the present study, the differences in the succession of microbiota in stainless steel and FRP panels may be due to differences in materials and nature of the surfaces used. Similarly assessment of bacterial population in the carbon steel inferred slow rate of succession of bacterial load with respect to time interval. This may be due to corrosive nature and changes in ionic charges of the substratum. Marszalek et al. (1979) also reported that loss of surface physical stability or the release of corrosion products which may be toxic at increased concentrations makes it biologically not suitable for fouling. Little and Wagner (1997) have reported that the rate of initial bacterial colonization is substratum-dependent, i.e. not all surfaces are colonized at the same rate or to be same the extent, hence the observed differences in biofilm bacterial load with different substrata is common.


In the marine environment, 90% of bacteria are Gram-negative with different characteristics and the Gram-negative cell wall is better adapted for survival in the marine environment (Das et al., 2006). Accordingly, here the data on diversity of biofilm bacterial strains in the experimental panels revealed that most of the strains identified were gram negative in nature. Among the isolated biofilm bacterial strains, the most predominant bacterium recorded in all the experimental panels were P. aeruginosa (34-39%), followed by H. aquamarina (20-24%) and V. alginolyticus (14-21%) respectively. The other biofilm bacterial strains recorded were in smaller proportion (1-8%). In consonance to the present study, occurrence of bacterial strains like V. alginolyticus (Sonak and Bhosle, 1995), E. coli, P. aeruginosa, S. oneidensis (Lee and Newman, 2003), B. subtilis (Omoike and Chorover, 2004), Micrococcus sp., Staphylococcus sp., Pseudomonas sp., Salmonella sp., Vibrio sp. and Proteus sp. (Hari et al., 2009) have been found in the biofouling process. Swami and Udhayakumar (2010) reported that the potential quality and quantity of fouling assemblages at a given site or environment depend on its biotic and hydrographical condition. Hence, the observed variation in occurrence and composition of bacterial population may be attributed to the biotic and hydrographical condition prevailed in the study area.

Marine natural products or extracts with antifouling activities have been isolated from a wide number of seaweeds (Chambers et al., 2006). In view of the above, in the present study antifouling activity of brown seaweed S. wightii was evaluated against fouling organisms through relevant antifouling assays. Antimicrofouling assay through disc diffusion is a rapid method particularly useful for initial screening of antimicrobial activity (Jenkins et al., 1998). Generally increasing polarity of solvents tends to extract the range of active metabolites from the plant materials. Accordingly, the antibacterial activity of different solvent extracts of seaweed S. wightii was tested against biofilm bacterial strains and the results inferred that methanolic extract had 100% growth inhibitory activity with the zone of inhibition ranged between 9-13 mm; whereas, the antagonistic activity of chloroform and hexane extracts of S. wightii was less. In agreement to this, Ara (2001) pointed out that methanol soluble fraction of brown seaweed such as S. binderii, S. swatzii, S. tenerrimum, S. variegatum and S. wightii had better zone of inhibition between 7-15 mm against bacterial strains such S. aureus, B. subtilus, S. typhimurium, E. coli and P. aeruginosa than chloroform and hexane soluble fractions. In general, ability of an extract to represent bioactivity depends on the chemodiversity within the extract. Like wise in the present study, differences in the growth inhibitory activity exerted by different solvent extracts may be due to unequal distribution of antimicrobial substances. Similarly, Bhadury and Wright (2004) have clearly pointed out that brown macroalgae are the major producers of antifouling defenses. Rizwi and Shameel (2003) inferred that methanolic extract of brown algae had better antibacterial activity against both gram positive and gram negative bacteria than green and red algae. Sastry and Rao (1995) stated that the antibacterial activity of S. wightii against S. aureus, P. vulgaris, E. coli, S. typhi, S. paratyphi A, S. typhiridium and P. aeruginosa was mainly due to the phycoconstitutent dioctyl phthalate.

Among the invertebrate species that cause biofouling problems, mussels are one of the major fouling organisms settling on man made or natural surfaces. Generally mussels are used as one of the bioindicators to study the antifouling potency against the macroorganisms. Assessment of toxicity of marine compounds is vital for induction of compounds in antifouling paints. In the present study, based on the results of antibacterial activity, the prominent result rendered methanolic extract of S. wightii was subjected for mussel bioassay. The results inferred the significant antifouling chemical defense strategy of crude methanolic extract of S. wightii and documented complete inhibition of byssal thread production and attachment of brown mussel P. indica at lower concentration ([EC.sub.50]) of 205 [+ or -] 14.7 [micro]g [ml.sup.-1]. The [EC.sub.50] value recorded by methanolic extract of S. wightii relatively less when compared to the results of Wilsanand et al. (1999), where they recorded [EC.sub.50] value of crude extracts of sponges, gorgonians, soft corals and antiptharians against green mussel P. virdis between 164 [+ or -] 12 to 898 [+ or -] 11 [micro]g [ml.sup.-1]. The findings of the present result thus fall in line with the observations of Rittschoff et al. (1992) and clearly manifested that the composition of secondary metabolites localized in the crude extract of S. wightii has actively participated in byssal thread inhibition and settlement of mussel P. indica. It also indicated that the [EC.sub.50] value obtained is lesser than the [LC.sub.50] value and thus indicate the non toxic nature of the extract. A similar phenomenon of inhibition of fouling organisms such as bacteria, fungi and mussels without any toxicity by extracts of brown algae S. muticum and red algae P. lanosa was also recorded by Hellio et al. (2000a, b). Accordingly, it was reported that often mussels close their shells and secrete fewer byssal thread with increasing concentration of the active extract and this may be an important criterion in improving the survivability and loss of attachment of mussels to the substrata during the course of experiment (Wilsanand et al., 1999).

The marine crustacean Artemia is ideally suited as a bioassay organism for detecting toxicity in plant extracts (Lewis, 1995). Toxicity against Artemia may be an indication of potency against marine fouling organisms, more specifically; crustacean foulers like barnacles can also be used to determine the toxicity of antifouling paints (Persoone and Castritsi-Catharios, 1989). In this context, in the present study different concentrations of methanolic extract of S. wightii was subjected to determine the cytotoxicity ([LC.sub.50]) effect against Artemia nauplii and was recorded as 161 [+ or -] 4.64 [micro]g [ml.sup.-1]. Results of the present study inferred increase in mortality rate with respect to increase in concentration of the extract. In accordance to the present study, Ayesha et al. (2010) examined ethanolic extracts of nine seaweeds for their cytotoxicity using brine shrimp Artemia salina and inferred that Dictyota indica, Iyengaria stellata and Melanothamnus afahusainii were found to be effective and showed considerable activity with the [LC.sub.50] value of 141, 186, 190 [micro]g [ml.sup.-1]. Also they pointed out that Phaeophyta group (brown seaweeds) possesses more cytotoxic property than others. In general, lethality of an extract mainly depends on the potent cytotoxic property of the extract. Also it was reported that degree of lethality is directly proportional to the concentrations of the extract.

Generally, presence of phytochemicals varied with respect to plant and solvent used for extraction. In the present study, based on the key result of antifouling activity, preliminary phytochemical constituents of in methanolic extract of S. wightii was identified. It inferred that the presence of phytochemicals viz. phenols, coumarins, quinones, saponins, alkaloids and flavanoids. In comparable with the present result, Mansuya et al. (2010) revealed the presence of possible phytochemicals such as alkaloids, steroids, tannin, phenols, saponins, proteins, flavonoids and volatile oils in aqueous and methanolic extract of S. wightii. The presence of one or more phytochemical (Secondary metabolites) present in the extract might be responsible for antibacterial and antimacrofouling activity. Several reports were well documented that phytochemicals such as phenols, flavanoids and terpenoids in the brown seaweeds were found to display antibacterial, antiviral and antitumor activity (Chuyen et al., 1982; Parameswaran et al., 1996 and Meenakshi et al., 2009). In general, presence of functional groups in the extract exerts significant biological activity. In the present study, FT-IR analysis of methanolic extract of S. wightii inferred the presence of possible functional groups such as halogen, carbon-nitrogen covalent bonding, carbon-carbon covalent bonding, nitro group, hydroxyl group and carbon-hydrogen covalent bonding. Presence of functional groups in the crude extracts may also be an important factor in contributing antifouling activity against bacteria and mussel P. indica. Thus the overall results of the present study clearly revealed the antifouling property of S. wightii and further research on identification is being directed for the development of green antifouling coatings.


The authors gratefully acknowledge NMRL, DRDO, Govt. of India (No: NMRL/mmt/0401/04/MBT/LP/ 128/2008) for providing financial assistance to carry out this work through the Project.


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(1) Palanisamy Iyapparaj, (1) Ramasamy Ramasubburayan, (1) Thulasi Raman, (2) Narayana Das, (2) Pradeep Kumar, (1) Arunachalam Palavesam and (1) Grasian Immanuel

(1) Centre for Marine Science and Technology, Manonmaniam Sundaranar University, Rajakkamangalam 629 502, Tamilnadu, India

(2) Naval Materials Research Laboraotory, DRDO, Ambernath, Maharastra, India

Corresponding Author: Grasian Immanuel, Centre for Marine Science and Technology, Manonmaniam Sundaranar University, Rajakkamangalam 629 502, Tamilnadu, India

E-mail: Telefax: +914652 253078
Table 1: Daily variations in physicochemical parameters
of seawater of the study area.

Physicochemical parameters            Time (h)


Salinity                         35.50[+ or -]0.00
Temperature ([degrees]C)         28.00[+ or -]0.00
DO (mg/l)                         5.20[+ or -]0.12
pH                                8.00[+ or -]0.00
Silicate ([micro]/mol)           0.149[+ or -]0.002
Nitrate ([micro]/mol)            2.238[+ or -]0.020
Nitrite (n/mol)                  0.319[+ or -]0.010
Total phosphorous ([micro]/mol)  19.5[+ or -] 1.20
Phosphate ([micro]/mol)          16.10[+ or -]1.50
Ammonia ([micro]/mol)            0.023[+ or -]0.002

Physicochemical parameters            Time (h)


Salinity                         35.00[+ or -]0.00
Temperature ([degrees]C)         28.00[+ or -] 0.00
DO (mg/l)                         5.40[+ or -]0.10
pH                                8.20[+ or -]0.00
Silicate ([micro]/mol)           0.152[+ or -]0.003
Nitrate ([micro]/mol)            2.242[+ or -]0.012
Nitrite (n/mol)                  0.327[+ or -]0.002
Total phosphorous ([micro]/mol)  19.20[+ or -]1.50
Phosphate ([micro]/mol)          16.26[+ or -]1.10
Ammonia ([micro]/mol)            0.026[+ or -]0.004

Physicochemical parameters            Time (h)


Salinity                         35.50[+ or -]0.00
Temperature ([degrees]C)         27.00[+ or -]0.00
DO (mg/l)                         5.30[+ or -]0.16
pH                                8.10[+ or -]0.00
Silicate ([micro]/mol)           0.155[+ or -]0.002
Nitrate ([micro]/mol)            2.235[+ or -]0.008
Nitrite (n/mol)                  0.332[+ or -]0.011
Total phosphorous ([micro]/mol)  18.90[+ or -]1.30
Phosphate ([micro]/mol)          15.90[+ or -]1.30
Ammonia ([micro]/mol)            0.022[+ or -]0.003

Each value is a Mean [+ or -] SD of three replicates

Table 2: Percentage diversity of biofilm bacteria in different

Biofilm Bacterial        % Diversity in tested panels

                            Wood                FRP

P. aeruginosa         34.0[+ or -]1.63    36.0[+ or -]2.45
H. aquamarina         21.0[+ or -]0.47    20.0[+ or -]0.82
V. alginolyticus      14.0[+ or -]0.82    18.0[+ or -]2.45
E. agglomerans         9.0[+ or -]0.82    7.0[+ or -]0.47
S. marcescens          8.0[+ or -]1.63    5.0[+ or -]0.82
S. liquefaciens        0.0[+ or -]0.0     2.0[+ or -]0.82
V. fischeri            6.0[+ or -]1.63    6.0[+ or -]0.81
V. parahaemolyticus    0.0[+ or -]0.0     3.0[+ or -]0.82
S. flexneri            3.0[+ or -]0.82    1.0[+ or -]0.82
A. hydrophila          2.0[+ or -]0.82    1.0[+ or -]0.82

Biofilm Bacterial        % Diversity in tested panels

                      Stainlesss steel     Carbon steel

P. aeruginosa         39.0[+ or -]2.45   35.0[+ or -]0.82
H. aquamarina         22.0[+ or -]1.63   24.0[+ or -]2.16
V. alginolyticus      18.0[+ or -]1.63   21.0[+ or -]2.45
E. agglomerans        5.0[+ or -]0.82    5.0[+ or -]1.63
S. marcescens         6.0[+ or -]1.63    6.0[+ or -]2.45
S. liquefaciens       3.0[+ or -]0.82     0.0[+ or -]0.0
V. fischeri           3.0[+ or -]0.94    4.0[+ or -]1.24
V. parahaemolyticus   2.0[+ or -]0.82    2.0[+ or -]0.82
S. flexneri           1.0[+ or -]0.82    2.0[+ or -]0.82
A. hydrophila         1.0[+ or -]0.82    1.0[+ or -]0.82

Each value is a Mean [+ or -] SD of three replicates

Table 3: Antimicrofouling activity (Zone of inhibition--mm)
of solvent based extracts of seaweed S. wightii.

Bacterial strains             Solvent extracts

                          Methanol          Chloroform

P. aeruginosa         13.0[+ or -]0.40   7.5[+ or -]0.44
H. aquamarina         9.0[+ or -]0.23    6.5[+ or -]0.44
V. alginolyticus      12.0[+ or -]0.62   8.0[+ or -]0.62
E. agglomerans        12.5[+ or -]0.44   11.0[+ or -]0.23
S. marcescens         10.5[+ or -]0.44    0.0[+ or -]00
S. liquifaecians      13.0[+ or -]0.40   12.0[+ or -]0.40
V. fischeri           11.0[+ or -]0.62   6.0[+ or -]0.11
V. parahaemolyticus   10.5[+ or -]0.44   11.0[+ or -]0.23
S. flexneri           12.5[+ or -]0.44   6.0[+ or -]0.62
A. hydrophila         10.0[+ or -]0.11   8.0[+ or -]0.40

Bacterial strains     Solvent extracts


P. aeruginosa         8.5[+ or -]0.44
H. aquamarina          0.0[+ or -]00
V. alginolyticus      8.0[+ or -]0.62
E. agglomerans        8.0[+ or -]0.23
S. marcescens         7.0[+ or -]0.23
S. liquifaecians      8.0[+ or -]0.40
V. fischeri           7.5[+ or -]0.44
V. parahaemolyticus   8.0[+ or -]0.11
S. flexneri           6.5[+ or -]0.44
A. hydrophila          0.0[+ or -]00

Each value is a Mean [+ or -] SD of three replicates

Table 4: [EC.sub.50] and [LC.sub.50] values of crude methanolic
extract of S. wightii used in Mussel Bioassay

Effective concentration ([EC.sub.50])/    Concentration of
Lethal Concentration ([LC.sub.50])        the extract
                                          ([micro]g [ml.sup.-1])

[EC.sub.50]                               205[+ or -]14.7
[LC.sub.50]                               306[+ or -]19.6

Each value is a Mean [+ or -] SD of three replicates

Table 5: Anticrustacean assay of crude methanolic extract of
S. wightii.

Concentration of the   No. of Artemia       No.         [LC.sub.50]
extract ([micro]g      nauplii exposed   Responding      ([micro]g
[ml.sup.-1])                                           [ml.sup.-1])

Control                      10              0        161[+ or -]4.64
30                           10              0
60                           10              0
90                           10              0
120                          10              1
150                          10              3
180                          10              5
210                          10              9
240                          10              10

Each value is a Mean [+ or -] SD of three replicates

Table 6: Qualitative phytochemical constituents
of methanolic extract of S. wightii.

Sl.No    Phytochemical     Present/Absent

1          Alkaloids             +
2         Glycosides             -
3          Saponins              +
4           Phenols              +
5         Falvonoids             +
6          Steroids              -
7           Tannins              -
8       Carboxylic acid          -
9          Quinones              +
10          Resins               -
11         Coumarins             +

+ Present; - Absent
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Title Annotation:Original Article
Author:Iyapparaj, Palanisamy; Ramasubburayan, Ramasamy; Raman, Thulasi; Das, Narayana; Kumar, Pradeep; Pala
Publication:Advances in Natural and Applied Sciences
Article Type:Report
Geographic Code:9INDI
Date:Mar 1, 2012
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