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Potential antimicrobial activity of marine molluscs from tuticorin, southeast coast of India against 40 biofilm bacteria.

ABSTRACT Methanol: water (1:1), methanol: dichloromethane (1:1) and acetone extracts of molluscs comprising 77 whole body, four inks, four opercula, 10 egg masses, and 10 digestive glands were screened for antimicrobial activity on marine biofilm bacteria. The methanol: water (1:1) whole body extracts of Nerita albicilla and Nerita oryzarum showed broad spectral inhibitory activity against 93% and 95% of the 40 biofilm bacteria. The egg masses from 10 gastropods showed activity against more than a quarter of the biofilm bacteria. The methanolic extract of Chicoreus virgineus, Chicoreus ramosus egg masses, and acetone extract of the egg mass of Rapana rapiformis showed broad-spectrum antibacterial activity against all the 40 biofilm bacterial strains. The activity in gastropod egg masses was localized to their internal matrix. Overall screening showed activity in 38.1% of the methanolic extracts followed by 13.3% of methanol: water, 12.4% of methanol: dichloromethane, and 3.8% of acetone extracts. Gastropods showed good activity when compared with bivalves and cephalopods.

KEY WORDS: antifouling, bacteria, biofilm, egg mass, mollusc


The efforts to control marine biofouling are ongoing since humans started venturing into the sea. Among the antifoulants developed to date, organotin compounds are considered as the most effective and have been widely used since the 1970s. It is well established that TBT compounds cause imposex universally (Barroso et al. 2002) and has been documented in over 118 species in 63 genera (Bettin et al. 1996). Organotin pollution has also raised concerns over the accumulation of organotin compounds in the food chain and the associated risks related to the presence of contaminants (Belfroid et al. 2000). International Maritime Organization (IMO) has proposed a complete phasing out of TBT use by the year 2008. The alternative tin free coatings are not only costly, but also their metal constituents may pose environmental problems (Hellio et al. 2004). So, a great deal of research has been focused on finding an alternate nontoxic and eco-friendly antifoulant (De Nys & Steinburg 2002, Fusetani 2004). In this context, marine organisms, especially the sedentary forms have interested researchers very much because they exhibit characteristic chemical defense against the epizooic organisms. More than 100 species of marine organisms have been shown to exhibit antimicrobial activity, as well as the ability to prevent settlement of fouling organisms (Clare 1996, Koh 1997).

Most currently available antifouling compounds are active against barnacles and algae and not against microbes like biofilm bacteria, which cause initial fouling of a submerged surface (Tang & Cooney 1998). Bacteria are the first organisms to foul submerged surfaces in the marine environment. Their subsequent multiplication and production of exopolymers leads to the formation of bacterial biofilms, which attract a large number of secondary fouling organisms (Steinberg et al. 2002). The interaction between bacteria and macrofoulers are dynamic and varied. The resultant biofouling community leads to economic and ecologic implications on both human-made structures and living surfaces.

However, bacteria do not colonize all submerged surfaces in a uniform manner and are influenced by many physical and chemical factors (Fletcher & Marshall 1982). The colonization process of the bacteria in the living marine surfaces may also be influenced by the secondary metabolites produced by the epibiont host. There exists an inverse correlation between macrofouling and antimicrobial activity of the epibiont host (Koh 1997, Santhana Ramasamy & Murugan 2003). This has been proposed as evidence that secondary metabolites are involved in the control of epibiosis (Slattery et al. 1995, Newbold et al. 1999). So, the natural mechanism of upsetting the development of the microfouling bacterial community by nontoxic marine natural compounds produced by the foul free host organisms may be taken as the most desirable way for breaking the fouling chain to reduce macrofouling settlement.

The cnidarians, sponges, ascidians, bryozoans, marine plant, and the like have received much attention and widely studied for antifouling activity (Clare 1996, Fusetani 2004). The study on the bioactivity of marine mollusc metabolites is scanty except for the nudibranchs. Rinehart et al. (1981) opined that marine molluscs might provide potential for isolating compounds with specific activity against certain microorganisms or cell types. Molluscan forms do exhibit adaptations to avoid fouling of their surface. The rare incidence of fouling on the Helgoland snails of Littorina littorea is due to the density dependent mutual grazing (Wahl et al. 1998). The periostracum of the mussel Mytilus edulis has been shown to possess some repellents to avoid fouling of their surface (Wahl et al. 1998).

The gelatinous egg masses of marine molluscs also have no physical protection from predation. So, many marine molluscs might have evolved mechanisms to avoid predation and surface fouling on their egg masses. Lord (1986) has demonstrated that the content of the egg capsules in Nucella lapillus was free from bacterial contamination and the capsule wall was impermeable to bacteria. Benkendorff et al. (2001) indicated that antibacterial compounds are common in both tough egg capsules and gelatinous egg masses, suggesting that chemical defense is the primary mechanism used to maintain an axenic environment for embryonic development. The egg masses of some marine molluscs have been the source of several compounds with antibacterial activity (Kamiya et al. 1984, Matsunaga et al. 1986, Yamazaki 1993, Benkendorff et al. 2000a, Pawlik et al. 1988). But, study on the antimicrobial properties of the marine molluscs against marine bacteria is scanty except a few works (Kamiya et al. 1984, Benkendorff et al. 2000b, Mitta et al. 2000, Benkendorff et al. 2001) and no previous studies have tested for activity against biofilm bacteria. So, in this study antimicrobial activity of a broad range of marine molluscs was evaluated against marine biofilm bacteria.


The screened molluscs in the present study are constituted by 77% gastropods, 21% bivalves, and 2% cephalopods comprising the whole body, digestive gland, opercula, ink, and egg masses. It is to be noted that the ecologic role of the activity has not been addressed in this study. Because the study is on shelled molluscs and only the inanimate surfaces of the animals are exposed to the marine environment, the whole body was selected instead of surface layer. The digestive gland was also analyzed from a subset of species, because there have been reports of bioaccumulation of secondary metabolites by some molluscs from their food (Faulkner 1988, Cimino & Sodano 1993, Rogers et al. 2000). The foul free surface of the opercula is the reason for its inclusion in the study. The previous report of antimicrobial property of egg masses (Benkendorff et al. 2001) has formed the basis of their selection.

Extraction of Molluscs

Molluscs collected alive from the fish landings of the Tuticorin coast (Lat. 8[degrees]45'N; Long. 78[degrees]10'E) were transported to the laboratory and identified up to species level using standard keys (Satyamurti 1952a and b, Hylleberg & Kilburn 2002, Personal communications from Dr. Alan Kohn, University of Washington, USA and Mr. Peter Middelfart of Australian Museum, Australia; BIOSIS Indo-Pacific Mollusc database). The shells were cracked by a light tap with a hammer to remove the soft parts. Then the tissues were rinsed with sterile distilled water and the whole body, digestive gland, opercula, inks, and egg masses of the respective molluscs were dissected and extracted (10 mL solvent to 0.1 g of tissue) by using the solvents methanol: dichloromethane (1:1), methanol, methanol: water (1:1), and acetone separately. The solvents/combinations used for extraction were developed based on random experiments. Replicate tissue samples were used for extraction in different solvents. The molluscan whole body and the digestive glands were cut into small pieces and air-dried for 24 h at room temperature before extraction with solvents. The opercula were air dried, ground well, and extracted with solvents. Fresh encapsulated egg masses were crushed before extraction with solvents. The developmental stages were not taken into consideration in this study. The effect of epibiotic bacteria and the presence of activity in the internal matrix of the egg masses of Chicoreus virgineus, C. ramosus, and Rapana rapiformis were determined by screening the empty (spent) egg masses. Raw inks obtained from the respective glands were directly used for extraction. The extracts were cold steeped overnight at -18[degrees]C, filtered with Whatman No. 1 filter paper, evaporated, and concentrated (Becerro et al. 1994, Riguera 1997, Wright 1998, Murugan & Santhana Ramasamy 2003). These crude extracts were used for antibacterial assay against biofilm bacterial strains isolated from the panels.

Isolation of Biofilm Bacteria

The fouling bacteria used in the antibacterial assay were isolated from the biofilm formed over wood, stainless steel, and fiber panels by pour plate technique (Wahl 1995). The panels were deployed for about a month during February and June 2002 at a depth of 1 m in the vicinity of Tuticorin Harbour (Lat 8[degrees]45'N; Long 78[degrees]10'E). The panels were washed with sterile seawater before swabbing with sterile cotton swabs. The swabs were placed in a tube containing sterile seawater, serially diluted and inoculated in marine agar plates (Zobell Marine Agar, Himedia, Mumbai). The plates were incubated at room temperature for 7 days. The pure bacterial strains based on colony morphology, pigmentation, and appearance were isolated by repeated streaking and identified up to genus level (Holt et al. 1994). The isolated strains were stored in marine agar slants at 4[degrees]C.

Antibacterial Assay

Antibacterial assay was carried out by using the standard disc diffusion method (Becerro et al. 1994, Slattery et al. 1995, Murugan & Santhana Ramasamy 2003). The disc diffusion assay with the marine bacteria as the target organism has been widely used as a primary screening tool to assess the antifouling potential of a substance (Clare 1996). The advantage of such a preliminary screening is that it is simple, less time-consuming, and requires only a small quantity of the material, which ensures minimal removal of biodiversity from the environment (Devi et al. 1997). But, the results of the disc diffusion assay have to be interpreted carefully taking into account the fact that the lipophilic active compounds do not migrate across the watery agar matrix (Benkendorff et al. 2000b). The disc diffusion assay in the present study is not used for quantitative estimation of the activity or for comparison between different species.

The Whatman No. 1 sterilized discs of 9-mm diameter, impregnated with 50 [micro]g/disc of the crude extract, and air dried were placed on the marine agar plates (90 x 15 mm) seeded with individual biofilm bacterial strains. The plates were incubated for 24 h at 30[degrees]C. The 40 biofilm bacterial strains isolated from the fouling panels were used for the antibacterial assay. The biofilm bacterial strains were grown in Zobell marine broth for 24 h before seeding the marine agar plates. The assay was carried out in triplicate. Control plates with solvents were also maintained separately. The zone of inhibition was measured from the edge of the disc to the clear zone in millimeter. The positive results of the antibacterial assay were expressed as percent inhibition of extract against 40 biofilm bacteria.


The 105 crude extracts of various parts of marine molluscs tested against 40 biofilm bacteria showed varied antibacterial activity (Tables 1, 2, 3, 4, and 5). The overall screening showed activity in 38.1% of the methanolic extracts followed by 13.3% of methanol: water, 12.4% of methanol: dichloromethane and 3.8% of acetone extracts. No activity against any biofilm bacteria was observed for 32.4% of the extracts.

Characterization of Bacteria

The 40 biofilm bacterial strains isolated from the fouling panels included five strains of Vibrio sp., four strains each of Micrococcus sp., Pseudomonas sp., Flavobacterium sp., Aeromonas sp., Bacillus sp., Enterobacter sp., Alteromonas sp., Cornebacterium sp., and three species of Alcaligenes sp. (Table 6).

Antibacterial Assay of Whole Body Extracts

Nearly 71.4% of the 77 species tested using whole body molluscan extracts showed activity (Table 1). The 14.3% of the species showed activity against more than 50% of biofilm bacteria. Extracts from only two species (2.6%) showed high activity and inhibited more than 90% biofilm bacteria. The remaining 28.6% of the whole body extracts did not show any activity.

Among the whole body extracts of the gastropods screened, 70.7% showed activity against some biofilm bacteria and no activity was observed in 29.3% of the species. Among the gastropod extracts, which showed activity, 75.6% showed inhibitory activity against at least 50% of biofilm bacteria and 24.4% inhibited the growth of more than 50% biofilm bacteria. The methanol: water (1:1) extracts of Strombus marginatus (23), methanol: dichloromethane (1:1) extract of Cronia margariticola (28), methanol extracts of Bufonaria rana (47), Turbo bruneus (7), and acetone extract of Murex tribulus (30) exhibited activity against more than 60% of bacteria (Table 1). The whole body methanol: water (1:1) extracts of Nerita albicilla and Nerita oryzarum showed broad spectral activity against 95% and 93% of the biofilm bacteria, and the inhibition zone ranged from nil to 4 mm and nil to 8 mm respectively (Table 6). N. oryzarum methanol: water extracts showed high inhibitory activity against Aeromonas sp. 1 (Table 6).

With regard to the bivalves, 70.6% of the whole body extracts showed activity, almost equal to that of gastropod. But, no high activity (not more than 60% strains inhibited) was observed for bivalve extracts. Nearly 91.7% of the bivalve species showed activity against less than 50% biofilm bacteria and 29.4% showed no activity. Among the bivalves, the methanol: dichloromethane extract of Pinna bicolor (72) showed high inhibitory activity against 55% biofilm bacteria. The activity of two cephalopod whole body extracts was not prominent and they exhibited inhibition of less than 40% of biofilm bacteria (Table 1).

Antibacterial Assay of Digestive Gland, Ink, and Opercula Extracts

The digestive gland extracts of the 10 gastropods did not show much activity. Only the methanol extracts of Melo melo (81) and Haliotis varia (78) exhibited activity, with the remaining digestive gland extracts showing no activity (Table 2).

The ink extracts of two gastropods and two cephalopods, screened for antibacterial activity, did not show promising inhibitory activity either. Only the methanolic extracts of Aplysia extraordinaria (88), Dolabella auricularia (89) inks showed activity, but against less than 50% of biofilm bacteria (Table 3).

Among the five opercula screened, only that of Lambis lambis (93) and Chicoreus virgineus (95) showed activity (Table 4). The activity exhibited by opercula of Lambis lambis (93) was not prominent because it inhibited only 15% of biofilm bacteria.

Antibacterial Assay of Molluscan Egg Mass Extracts

Interestingly, all encapsulated egg masses of the 10 gastropods exhibited inhibitory activity (Table 5). Sixty percent of the egg masses showed activity against more than 50% of biofilm bacteria. The egg masses of the family Muricidae inhibited the highest number of biofilm bacteria, with, Chicoreus virgineus, C. ramosus, and Rapana rapiformis showing broad spectrum antibacterial activity against all the 40 biofilm bacterial strains (Table 6). The methanolic extract of Chicoreus virgineus egg mass showed an inhibition zone ranging from 1-10 mm against different bacteria. The acetone extract of the egg mass of Rapana rapiformis showed an inhibition zone of 1-7 mm. The methanolic extract of Chicoreus ramosus egg mass showed an inhibition zone of 1-5 mm. The extracts of these three egg masses showed high inhibitory activity against the biofilm bacteria Vibrio sp. 2 (Table 6). The extracts of empty egg masses did not show any activity.


Forty biofilm bacterial strains belonging to 10 genera were isolated and used in antibacterial assays. The strains are the common marine biofilm bacteria in India (Santhana Ramasamy & Murugan 2003). It is well known that many fouling invertebrates require the presence of a microbial surface film as a prerequisite for settlement (Scheltema 1974). Many marine organisms produce bioactive metabolites, which may play a direct role in preventing fouling (Pawlik 1993) and their production can be considered as a kind of autogenic protection (Wahl et al. 1998). In this study, some marine molluscs were shown to inhibit the growth of biofilm bacteria, although since the activity was not associated with surface tissues, the ecologic role is uncertain. Nevertheless, the extracts of marine organisms showing antimicrobial activity against biofilm bacteria could have potential applications as antifouling agents, irrespective of their ecologic function.

In general, the whole body extracts of the molluscs with very few exceptions showed antibacterial activity against a small proportion of biofilm bacteria. Among the inhibitory activity observed in the whole body extracts, only Nerita albicilla and Nerita oryzarum showed broad spectral activity (Table 6). The observed inhibition range of [less than or equal to]8 mm was also higher when compared with the whole body extracts of Cerithidea cingulata and Hemifusus pugilinus, which showed a range of up to 1.5 mm against nine pathogenic bacteria (Rajaganapathi 1996). The broad spectral activity observed for Nerita albicilla could be due to the previously reported occurrence of antibacterial secondary metabolites in Nerita albicilla (Sanduja et al. 1985).

The bivalve whole body extracts in this study have not shown promising activity except the moderate activity observed for Pinna bicolor (72) and Pteria inquinata (74). The observation of lack of activity for Circe scripta (65) in the present study is in contrast to the observation of Jayaseeli et al. (2001) who have reported 20mm zone of inhibition for the ethanolic extract of Circe scripta against Bacillus subtilis. Though the previous reports indicated broad spectrum activity against the human pathogenic bacteria, the extracts of bivalves in the present study seem to be ineffective or inhibit fewer biofilm bacteria.

No promising antibacterial activity has been recorded for the whole body extracts and inks of cephalopods against the biofilm bacterial strains. But, broad-spectrum antibacterial activity for aqueous ink extract of the cephalopods Loligo duvaucelli and Sepia pharaonis against nine human pathogens has been reported (Patterson Edward & Murugan 2000). This suggests that marine biofilm bacteria are more resistant to the active compounds found in cephalopod ink.

The observation of antibacterial activity in the egg masses of all the 10 gastropods in this study (Table 5) substantiates previous reports that a number of marine molluscs protect their eggs with natural products (see Kamiya et al. 1984, Matsunaga et al. 1986, Yamazaki 1993, Benkendorff et al. 2000a, Pawlik et al. 1988). The observation on Rapana rapiformis egg mass coincides with the observation of Rajaganapathi (1996) and Prem Anand et al. (1997) who have observed broad spectrum activity of the egg mass against nine pathogenic bacteria. The gelatinous egg masses of molluscs in the Aplysiidae exhibit broad spectrum of biologic activity, and several glycoproteins from the egg masses of Aplysia kurodai and A. juliana were found to have antimicrobial activity (Kamiya et al. 1984, Yamazaki et al. 1984, Yamazaki et al. 1985, Kisugi et al. 1987, Yamazaki 1993).

The high antimicrobial activity of the egg masses of Chicoreus ramosus, C. virgineus, and Rapana rapiformis indicated the localization of the inhibitory substance in the egg masses. Three antimicrobial compounds namely Tyrindoleninone, Tyriverdin, and 6-bromoisatin from the egg masses of muricids have been attributed to the antimicrobial properties observed in these egg capsules (Benkendorff et al. 2001). Benkendorff et al. (2000a) observed a form of chemical ripening in the egg masses of Dicathais orbita where the toxic compound tyrindoleninone is converted into tyriverdin, a bacteristatic compound that is then converted into Tyrian purple at the time of hatching and has no antibiotic property. It is likely that the egg masses of Chicoreus ramosus, C. virgineus, and Rapana rapiformis possess the same types of bioactive compounds, because these also turn purple on exposure to sunlight.

The origin of these antibacterial compounds in the egg masses has been traced to hypobranchial glands. This could be substantiated from the observation of broad-spectrum inhibitory activity of the hypobranchial gland of Chicoreus virgineus against pathogenic bacteria (Rajaganapathi 1996, Prem Anand et al. 1997). Murugan et al. (1991) reported the antibacterial property of the hypobranchial gland extract of Rapana rapiformis against eight human pathogens. It is possible that the antimicrobial substances in the adult molluscs may be held in a nontoxic state and then enzymatically activated in the reproductive tract or egg masses as opined by Benkendorff (1999). This statement is substantiated by the observation of low antibacterial activity for the whole body extract of these three muricids in the present study.

The activity against fewer biofilm bacteria observed in Cronia margariticola and in other gastropod egg masses of Cassidae, Conidae, Cypraeidae, Alplysidae, and Buccinidae in the present study could be influenced by their stage of development. Benkendorff et al. (2001) demonstrated that antimicrobial metabolites are likely to be more concentrated in freshly laid egg masses than in well developed or hatching egg masses. This has been demonstrated in the gelatinous egg masses of sea hare Aplysia juliana wherein the loss of antimicrobial activity was observed with embryonic development (Kamiya et al. 1988). In this study, only the egg masses with developing embryos were selected and the developmental stages have not been taken into account.

Earlier studies indicate that a range of micro and macro algae and protozoans foul the egg capsules, which may have harmful or beneficial effects (Biermann et al. 1992, Benkendorff 1999, Przeslawski & Benkendorff 2005). However, the proteinaceous matrix of the leathery egg capsules appears to provide less suitable surface for fouling organisms than gelatinous egg masses (Przeslawski & Benkendorff 2005). The screening of empty egg cases of Chicoreus ramosus, C. virgineus, and Rapana rapiformis did not show any inhibitory activity, indicating the presence of active substances in only the intracapsular fluid and/or embryos of the egg masses. The wide spectrum antibacterial activity exhibited by the encapsulated egg masses of the three muricid gastropods indicated the presence of potential antifouling natural products that are worthy of further isolation and characterization.

In this study, the frequency of activity was high in methanolic extracts (Table 1-6) and acetone extract of Rapana rapiformis egg mass. The observation of lack of activity for 32.4% of the molluscan extracts may be due to limitations with the disc diffusion assay, because only hydrophilic compounds tend to diffuse across the watery agar surface (Benkendorff et al. 2000b).
Proportion of biofilm bacteria inhibited by the whole body extracts of
molluscs (% inhibition against 40 biofilm bacteria).

S. No Family Molluscan Species

 Gastropoda: Eogastropoda:
 1. Patellidae Cellana radiata (Born, 1778)
 2. Patellidae Cellana rota (Gmelin, 1791)
 Gastropoda: Orthogastropoda:
 3. Haliotidae Haliotis varia (Linne, 1758)
 4. Trochidae Euchelus asper (Gmelin, 1791)
 5. Trochidae Trochus radiatus (Gmelin, 1791)
 6. Trochidae Umbonium vestiarium (Linne, 1758)
 7. Turbinidae Turbo bruneus (Roding, 1798)
 Gastropoda: Orthogastropoda:
 8. Neritidae Nerita oryzarum (Recluz, 1841)
 9. Neritidae Nerita albicilla (Linne, 1758)
 Gastropoda: Orthogastropoda:
 10. Cypraeidae Cypraea tigiris (Linne, 1758)
 11. Cypraeidae Cypraea errones (Linne, 1758)
 12. Cypraeidae Cypraea onyx (Linne, 1758)
 13. Cypraeidae Cypraea arabica (Linne, 1758)
 14. Cypraeidae Cypraea vitellus (Linne, 1758)
 15. Cypraeidae Cypraea lentiginosa (Gray, 1825)
 16. Cypraeidae Cypraea caurica (Linne, 1758)
 17. Cypraeidae Cypraea moneta (Linne, 1758)
 18. Cypraeidae Cypraea gracilis (Gaskoin, 1848)
 19. Cypraeidae Cypraea caputserpentis (Linne, 1758)
 20. Cypraeidae Cypraea talpa (Linne, 1758)
 21. Strombidae Tibia delicatula (Nevill, 1881)
 22. Strombidae Tibia curta (Sowerby, 1842)
 23. Strombidae Strombus marginatus (Linne, 1767)
 24. Strombidae Lambis lambis (Linne, 1758)
 25. Olividae Agaronia gibbosa (Born, 1778)
 26. Muricidae Chicoreus virgineus (Roding, 1798)
 27. Muricidae Chicoreus ramosus (Linne, 1758)
 28. Muricidae Cronia margariticola (Broderip, 1832)
 29. Muricidae Rapana rapiformis (Born, 1778)
 30. Muricidae Murex tribulus (Linne, 1758)
 31. Muricidae Thais biserialis. (Blainville, 1832)
 32. Muricidae Thais bufo (Lamarck, 1822)
 33. Muricidae Thais tissoti (Petit, 1852)
 34. Volutidae Harpulina Iapponica (Linne, 1767)
 35. Buccinidae Cantharus wrightae (Cernohorsky, 1974)
 36. Conidae Conus araneosus (Lightfoot, 1786)
 37. Conidae Conus figulinus (Linne, 1758)
 38. Conidae Conus madagascariensis (Sowerby, 1858)
 39. Conidae Conus textile (Linne, 1758)
 40. Potamididae Cerithidea cingulata (Gmelin, 1791)
 41. Littorinidae Littorarina undulata (Gray, 1839)
 42. Littorinidae Littorarina scabra (Linne, 1758)
 43. Fasciolariidae Fusinus forceps (Perry, 1811)
 44. Cassidae Phalium glauccum (Linne, 1758)
 45. Cassidae Casmaria erinacea (Linne, 1758)
 46. Harpidae Harpa davidis (Roding, 1798)
 47. Bursidae Bufonaria rana (Linne, 1758)
 48. Naticidae Natica vitellus (Linne, 1758)
 49. Naticidae Tanea tigrina (Roding, 1798)
 50. Naticidae Polinices didyma (Roding, 1798)
 51. Naticidae Polinices albumen (Linne, 1758)
 52. Turritellidae Turitella duplicata (Linne, 1758)
 53. Ranellidae Cymatium perryi (Emerson & Old, 1963)
 Gastropoda: Orthogastropoda:
 54. Ficidae Ficus ficus (Linne, 1758)
 55. Bullidae Bulla ampulla (Linne, 1758)
 56. Hydatinidae Hydatina velum (Gmelin, 1791)
 57. Aplysiidae Aplysia extraordinaria (Allan, 1932)
 58. Aplysiidae Dolabella auricularia (Lightfoot, 1786)
 Cephalopoda: Teuthoidea
 59. Loliginidae Sepioteuthis lessoniana (Lesson, 1830)
 Cephalopoda: Sepioidea
 60. Sepiidae Sepia pharaonis (Ehrenberg, 1831)
 Bivalvia: Veneroida
 61. Donacidae Donax faba (Gmelin, 1791)
 62. Donacidae Donax cuneatus (Linne, 1758)
 63. Veneridae Gafrarium tumidum (Roding, 1798)
 64. Veneridae Gafrarium pectinatum (Linne, 1758)
 65. Veneridae Circe scripta (Linne, 1758)
 66. Veneridae Dosinia modesta (Sowerby)
 67. Veneridae Marcia opima (Gmelin, 1791)
 68. Veneridae Meretrix casta (Gmelin, 1791)
 69. Mesodesmatidae Atactodea striata (Gmelin, 1791)
 70. Cardiidae Vasticardium pectiniforme (Born, 1778)
 71. Carditidae Cardita bicolor (Linne, 1758)
 Bivalvia: Pterioida
 72. Pinnidae Pinna bicolor (Gmelin, 1791)
 73. Pteriidae Pteria castanea (Reeve, 1858)
 74. Pteriidae Pteria inquinata (Reeve, 1857)
 Bivalvia: Mytiloida
 75. Mytilidae Modiolus metcalfei (Hanley, 1843)
 Bivalvia: Ostreoida
 76. Spondylidae Spondylus layardi (Reeve, 1856)
 77. Placunidae Placuna placenta (Lamarck, 1819)

 Type of the Extract


 1. -- -- 20 --
 2. -- -- 23 --
 3. -- -- -- --
 4. -- 30 -- --
 5. -- -- -- --
 6. -- -- -- --
 7. 73 -- -- --
 8. -- 95 -- --
 9. -- 93 -- --
 10. -- -- 48 --
 11. -- -- 40 --
 12. -- -- 28 --
 13. -- -- 25 --
 14. -- -- 30 --
 15. -- -- -- --
 16. -- -- -- --
 17. -- -- -- --
 18. -- -- 45 --
 19. -- 38 -- --
 20. -- -- -- --
 21. -- 40 -- --
 22. -- 55 -- --
 23. -- 70 -- --
 24. -- -- -- 55
 25. -- 40 -- --
 26. 50 -- -- --
 27. 58 -- -- --
 28. -- -- 63 --
 29. -- -- -- 28
 30. -- -- -- 60
 31. 35 -- -- --
 32. 25 -- -- --
 33. 18 -- -- --
 34. 13 -- -- --
 35. -- -- -- --
 36. 33 -- -- --
 37. 30 -- -- --
 38. 15 -- -- --
 39. 33 -- -- --
 40. -- -- -- --
 41. -- -- 28 --
 42. -- -- 25 --
 43. -- -- 30 --
 44. -- -- -- --
 45. 20 -- -- --
 46. -- -- -- --
 47. 63 -- -- --
 48. -- -- -- --
 49. -- -- -- --
 50. -- -- -- --
 51. -- -- -- --
 52. 33 -- -- --
 53. -- -- -- --
 54. 45 -- -- --
 55. 40 -- -- --
 56. -- -- -- --
 57. 28 -- -- --
 58. 38 -- -- --
 59. 40 -- -- --
 60. 30 -- -- --
 61. -- 25 -- --
 62. -- 28 -- --
 63. 33 -- -- --
 64. 20 -- -- --
 65. -- -- -- --
 66. -- 25 -- --
 67. -- 38 -- --
 68. -- 30 -- --
 69. -- -- -- --
 70. 8 -- -- --
 71. -- -- -- --
 72. -- -- 55 --
 73. 30 -- -- --
 74. 45 -- -- --
 75. -- -- -- --
 76. -- -- -- --
 77. -- 10 -- --

M, Methanol; MW, Methanol: Water; MD, Methanol: Dichloromethane; A,

Proportion of biofilm bacteria inhibited by the molluscan
digestive gland extracts (% inhibition against 40 biofilm bacteria).

S. No Family Molluscan Species

 Gastropoda: Orthogastropoda:
 78. Haliotidae Haliotis varia (Linne, 1758)
 Gastropoda: Orthogastropoda:
 79. Fasciolariidae Pleuroploca trapezium (Linne, 1758)
 80. Tonnidae Tonna dolium (Linne, 1758)
 81. Volutidae Melo melo (Lightfoot, 1786)
 82. Melongenidae Hemifusus pugilinus (Born, 1778)
 83. Buccinidae Babylonia spirata (Linne, 1758)
 84. Buccinidae Babylonia zeylanica (Bruguiere, 1789)
 85. Muricidae Chicoreus virgineus (Roding, 1798)
 86. Muricidae Chicoreus ramosus (Linne, 1758)
 87. Strombidae Lambis lambis (Linne, 1758)

 Type of the Extract


 78. 23 -- -- --
 79. -- -- -- --
 80. -- -- -- --
 81. 28 -- -- --
 82. -- -- -- --
 83. -- -- -- --
 84. -- -- -- --
 85. -- -- -- --
 86. -- -- -- --
 87. -- -- -- --

M, Methanol; MW, Methanol: Water; MD, Methanol: Dichloromethane; A,

Proportion of biofilm bacteria inhibited by the molluscan ink extracts
(% inhibition against 40 biofilm bacteria).

S. No Family Molluscan Species

 Gastropoda: Orthogastropoda:
 88. Aplysiidae Aplysia extraordinaria (Allan, 1932)
 89. Aplysiidae Dolabella auricularia (Lightfoot, 1786)
 Cephalopoda: Teuthoidea
 90. Loliginidae Sepioteuthius lessoniana (Lesson, 1830)
 Cephalopoda: Sepioidea
 91. Sepiidae Sepia pharaonis (Ehrenberg, 1831)

 Type of the Extract


 88. 30 -- -- --
 89. 43 -- -- --
 90. -- -- -- --
 91. -- -- -- --

M, Methanol; MW, Methanol: Water; MD, Methanol: Dichloromethane; A,

Proportion of biofilm bacteria inhibited by the molluscan opercula
extracts (% inhibition against 40 biofilm bacteria).

S. No Family Molluscan Species

 Gastropoda: Orthogastropoda: Neritimorpha
 92. Neritidae Nerita oryzarum (Recluz, 1841)
 Gastropoda: Orthogastropoda: Caenogastropoda
 93. Strombidae Lambis lambis (Linne, 1758)
 94. Buccinidae Babylonia spirata (Linne, 1758)
 95. Muricidae Chicoreus virgineus (Roding, 1798)

 Type of the Extract


 92. -- -- -- --
 93. 15 -- -- --
 94. -- -- -- --
 95. 35 -- -- --

M, Methanol; MW, Methanol: Water; MD, Methanol: Dichloromethane; A,

Proportion of biofilm bacteria inhibited by the encapsulated molluscan
egg mass extracts (% inhibition against 40 biofilm bacteria).

S. No Family Molluscan Species

 Gastropoda: Orthogastropoda:
 96. Muricidae Chicoreus virgineus (Roding, 1798)
 97. Muricidae Chicoreus ramosus (Linne, 1758)
 98. Muricidae Cronia margariticola (Broderip, 1832)
 99. Muricidae Rapana rapiformis (Born, 1778)
 100. Cassidae Casmaria erinacea (Linne, 1758)
 101. Conidae Conus madagascariensis (Sowerby, 1858)
 102. Cypraeidae Cypraea errones (Linne, 1758)
 103. Buccinidae Babylonia spirata (Linne, 1758)
 Gastropoda: Orthogastropoda:
 104. Aplysiidae Aplysia extraordinaria (Allan, 1932)
 105. Aplysiidae Dolabella auricularia (Lightfoot, 1786)

 Type of the Extract


 96. 100 -- -- --
 97. 100 -- -- --
 98. 70 -- -- --
 99. -- -- -- 100
 100. 55 -- -- --
 101. 53 -- -- --
 102. 35 -- -- --
 103. 35 -- -- --
 104. 40 -- -- --
 105. 28 -- -- --

M, Michael; MW, Methanol; Water; MD, Methanol; Dichloromethane; A,

Zone of inhibition (mm) of five molluscan crude extracts against 40
biofilm bacteria.

 Inhibition Zone (mm)

 Chicoreus Rapana ramosus
 virgineus rapiformis (Egg Mass)
Biofilm Bacteria (Egg Mass)(M) (Egg Mass)(A) (M)

Micrococcus sp. 1 7 5 3
Micrococcus sp. 2 5 4 3
Micrococcus sp. 3 4 2 1
Micrococcus sp. 4 3 2 1
Vibrio sp. 1 8 6 4
Vibrio sp. 2 10 7 5
Vibrio sp. 3 5 4 2
Vibrio sp. 4 3 2 1
Vibrio sp. 5 2 2 1
Pseudomonas sp. 1 6 4 3
Pseudomonas sp. 2 5 4 3
Pseudomonas sp. 3 6 3 3
Pseudomonas sp. 4 2 1 2
Alcaligenes sp. 1 3 1 1
Alcaligenes sp. 2 2 2 2
Alcaligenes sp. 3 6 5 3
Flavobacterium sp. 1 4 3 3
Flavobacterium sp. 2 3 1 1
Flavobacterium sp. 3 6 5 1
Flavobacterium sp. 4 7 4 1
Aeromonas sp. 1 4 3 2
Aeromonas sp. 2 8 3 2
Aeromonas sp. 3 7 5 2
Aeromonas sp. 4 3 3 3
Bacillus sp. 1 6 1 1
Bacillus sp. 2 3 2 3
Bacillus sp. 3 2 1 1
Bacillus sp. 4 4 1 1
Enterobacter sp. 1 4 3 4
Enterobacter sp. 2 5 1 3
Enterobacter sp. 3 8 4 5
Enterobacter sp. 4 2 2 1
Alteromonas sp. 1 5 2 3
Alteromonas sp. 2 7 5 4
Alteromonas sp. 3 4 3 3
Alteromonas sp. 4 6 4 4
Cornebacterium sp. 1 2 2 1
Cornebacterium sp. 2 1 1 1
Cornebacterium sp. 3 2 1 1
Cornebacterium sp. 4 4 2 1

 Inhibition Zone (mm)

 Nerita oryzarum
 albicilla (Whole Body)
Biorilm Bacteria (Whole Body)(MW) (MW)

Micrococcus sp. 1 1 2
Micrococcus sp. 2 2 4
Micrococcus sp. 3 4 1
Micrococcus sp. 4 1 1
Vibrio sp. 1 1 Trace
Vibrio sp. 2 1 1
Vibrio sp. 3 2 2
Vibrio sp. 4 2 2
Vibrio sp. 5 2 1
Pseudomonas sp. 1 1 1
Pseudomonas sp. 2 1 3
Pseudomonas sp. 3 Nil 1
Pseudomonas sp. 4 Trace Nil
Alcaligenes sp. 1 Trace 3
Alcaligenes sp. 2 2 3
Alcaligenes sp. 3 1 2
Flavobacterium sp. 1 Trace 1
Flavobacterium sp. 2 3 Trace
Flavobacterium sp. 3 3 4
Flavobacterium sp. 4 Trace Trace
Aeromonas sp. 1 1 8
Aeromonas sp. 2 4 3
Aeromonas sp. 3 4 3
Aeromonas sp. 4 2 2
Bacillus sp. 1 Trace Trace
Bacillus sp. 2 Nil Trace
Bacillus sp. 3 Nil Trace
Bacillus sp. 4 Trace 1
Enterobacter sp. 1 2 1
Enterobacter sp. 2 2 1
Enterobacter sp. 3 1 1
Enterobacter sp. 4 1 1
Alteromonas sp. 1 3 2
Alteromonas sp. 2 1 Nil
Alteromonas sp. 3 1 Trace
Alteromonas sp. 4 2 2
Cornebacterium sp. 1 2 3
Cornebacterium sp. 2 1 1
Cornebacterium sp. 3 3 1
Cornebacterium sp. 4 1 1

M, Methanol extracts; A, Acetone extracts; MW, Methanol: Water


The first author thanks DANIDA and Prof. Jorgen Hylleberg of Aarhus University, Denmark, for the fellowship through Tropical Marine Mollusc Programme (TMMP). The authors thank Dr. Alan Kohn, University of Washington, USA and Mr. Peter Middelfart of Australian Museum, Australia, for help in identification of specimens. The authors also gratefully acknowledge the constructive review of the manuscript by Dr. Kirsten Benkendorff of School of Biological Sciences, Flinders University, Australia.


Barroso, C. M., M. A. Reis-Henriques, M. S. Ferreira & M. H. Moreira. 2002. The effectiveness of some compounds derived from antifouling paints in promoting imposex in Nassarius reticulates. J. Mar. Biol. Assoc. UK. 82:249-255.

Biermann, C., G. Schinner & R. Strathmann. 1992. Influence of solar radiation, microalgal fouling, and current on deposition site and survival of embryos of a dorid nudibranch gastropod. Mar. Ecol. Progr. Ser. 86:205-215.

Becerro, M. A., N. I. Lopez, X. Turon & M. J. Uniz. 1994. Antimicrobial activity and surface bacterial film in marine sponges. J. Exp. Mar. Biol. Ecol. 179:195-205.

Belfroid, A. C., M. Purperhart & F. Ariese. 2000. Organotin levels in seafood. Mar. Pollut. Bull. 40(3):226-232.

Benkendorff, K. 1999. Bioactive molluscan resources and their conservation: Ph.D. Thesis, University of Wollongong, Australia. Available at: 154039/index.html.

Benkendorff, K., J. B. Bremner & A. R. Davis. 2000a. Tyrian purple precursors in the egg masses of the Australian muricid, Dicathais orbita: a possible defensive role. J. Chem. Ecol. 26:1037-1050.

Benkendorff, K., A. R. Davis & J. B. Bremner. 2000b. Rapid screening methods for detecting antimicrobial activity in the egg masses of marine molluscs. J. Med. & Appl. Malacol. 10:211-223.

Benkendorff, K., A. Davis & J. B. Bremner. 2001. Chemical defense in the egg masses of benthic invertebrates: An assessment of antibacterial activity in 39 Molluscs and 4 Polychaetes. J. Invertebr. Pathol. 78: 109-118.

Bettin, C., J. Oehlmann & E. Stroben. 1996. TBT-induced imposex in marine neogastropods is mediated by an increasing androgen level. Helgolander Meeresunters 50:299-317.

Cimino, G. & G. Sodano. 1993. Biosynthesis of secondary metabolites in marine molluscs. Top. Curr. Chem. 167:77-115.

Clare, A. 1996. Marine natural product antifoulants: status and potential. Biofouling 9:211-229.

De Nys, R. & P. D. Steinburg. 2002. Linking marine biology and biotechnology. Current Opinion in Biotechnology 13:244-248.

Devi, P., W. Solimabi, L. D'Souza, S. Sonal, S. Y. Kamar & S. Y. S. Singbal. 1997. Screening of some marine plants for activity against marine fouling bacteria. Bot. Mar. 40:87-91.

Faulkner, D. J. 1988. Feeding deterrent in Molluscs. in: D. G. Fautin, editor. Biomedical importance of marine organisms. California Academy of Sciences. pp. 29-36.

Fletcher, M. & K. C. Marshall. 1982. Are solid surfaces of ecological significance to aquatic bacteria? Adv. Microb. Ecol. 6:199-236.

Fusetani, N. 2004. Biofouling and antifouling. Nat. Prod. Rep. 21:94-104.

Holt, J. G., N. R. Krieg, P. H. Sneath, J. T. Staley & S. T. Williams. 1994. International edition: Bergey's manual of determinative bacteriology. 9th ed. Maryland: Williams & Wilkine.

Hellio, C., J.-P. Marechal, B. Veron, G. Bremer, A. S. Clare & Y. L. Gal. 2004. Seasonal variation of antifouling activities of marine algae from the Brittany Coast (France). Mar. Biotechnol. 6:67-82.

Hylleberg, J. & R. N. Kilburn. 2002. Annotated inventory of molluscs from the Gulf of Mannar and vicinity. In: J. Hylleberg & A. Nateewathana, editors. Zoogeography and inventory of marine molluscs encountered in Southern India. PMBC, Thailand. Phuket. Mar. Biol. cent. spec. publ. 29. pp. 19-70.

Jayaseeli, A. A., T. Prem Anand & A. Murugan. 2001. Antibacterial activity of 4 bivalves from Gulf of Mannar. Phuket. Mar. Biol. Cent. Spec. Publ. 25(1):215-217.

Kamiya, H., K. Muramoto & K. Ogata. 1984. Antibacterial activity in the egg mass of a sea hare. Experimentia 40:947.

Kamiya, H., K. Muramoto, R. Goto & M. Yamazaki. 1988. Characterization of the antibacterial and antineoplastic glycoproteins in a sea hare Aplysia juliana. Nippon Suisan Gakkaishi 54:773-777.

Kisugi, J., H. Kamiya & M. Yamazaki. 1987. Purification and Characterization of Aplysianin E, an antitumour factor from sea hare eggs. Cancer Res. 47:5649-5653.

Koh. 1997. Do scleratinian corals engage in chemical warfare against microbes? J. Chem. Ecol. 23:379-398.

Lord, F. 1986. Are the contents of egg capsules of the marine gastropod Nucella lapillus (L) axenic. Am. Malac. Bull. 4:201-203.

Matsunaga, S., N. Fusetani & K. Hashimoto. 1986. Kabiramide C, a novel antilungal macrolide from nudibranch egg masses. J. Am. Chem. Soc. 108:847-849.

Mitta, G., F. Vandenbulcke, F. Hubert, M. Salzet & P. Roche. 2000. Involvement of mytilins in mussel antimicrobial defense. J. Biol. Chem. 275:12954-12962.

Murugan, A. & M. Santhana Ramasamy. 2003. Biofouling deterrent natural product from the ascidian Distaplia nathensis. Indian J. Mar. Sci. 32:162-164.

Murugan, A., J. K. Patterson Edward & K. Ayyankannu. 1991. Ecological, biochemical and antibacterial data from India. Phuket Mar. Biol. Cent. Spec. Publ. No. 9:108-110.

Newbold, R. W., P. R. Jensen, W. Fenical & J. R. Pawlik. 1999. Antibacterial activity of Caribbean sponge extracts. Aquat. Microb. Ecol. 19: 279-284.

Patterson Edward, J. K. & A. Murugan. 2000. Screening of cephalopods for bioactivity. Phuket Mar. Biol. Cent. Spec. Publ. No. 21 (1):253-256. Pawlik. J. R., 1993. Marine invertebrate chemical defenses. Chem. Rev 93:1911-1922.

Pawlik, J. R., M. R. Kernan, T. F. Molinski & M. K. Harper. 1988. Defensive chemicals of the Spanish Dancer nudibranch Hexabranchus sanguineus, and its egg ribbons, Macrolids derived from a sponge diet. J. Exp. Mar. Biol. Ecol. 119:99-109.

Prem Anand, T., J. Rajaganapathi & J. K. Patterson Edward. 1997. Antibacterial activity of Marine Molluscs from Portonovo region. Indian J. Mar. Sci. 26:206-208.

Przeslawski, R. & K. Benkendorff. 2005. The role of surface fouling in the development of encapsulated gastropod embryos. J. Mollusc. Stud. 71: 75-83.

Rajaganapathi, J. 1996. Studies on antibacterial activity of five marine molluscs. M.Sc. thesis. Annamalai University, Parangipettai. India, pp. 43.

Riguera, R. 1997. Isolating bioactive compound from marine organisms. J. Mar. Biotechnol. 5:187-193.

Rinehart, K. L., P. D. Shaw, L. S. Shield, J. B. Gloer, G. C. Harbour, M. E. S. Koker, D. Samain, R. E. Schwartz, A. A. Tymiak, E. G. Swynenberg, D. A. String Fellow, J. J. Vavva, J. H. Coats, G. E. Zurenko, S. L. Kuentzel, L. H. Li, G. J. Bakus, R. C. Brasca, L. L. Craft, D. N. Young & J. L. Connot. 1981. Marine natural products as a source of antiviral, antimicrobial and antineoplastic agents. Pure Appl. Chem. 53:795-817.

Rogers, C., R. De Nys, T. Charlton & P. Steinberg. 2000. Dynamics of algal secondary metabolites in two species of sea hare. J. Chem. Ecol. 26:721-744.

Sanduja, R., A. J. Weinheimer, K. L. Euler & M. Alam. 1985. Unusual occurrence of faloplumierin, an antibacterial agent in the marine mollusc Nerita albicilla. J. Natl. Prod. 48:335-336.

Santhana Ramasamy, M. & A. Murugan. 2003. Chemical defense in ascidians Eudistoma viride and Didemnum psammathodes in Tuticorin, Southeast coast of India: Bacterial epibiosis and fouling deterrent activity. Indian J. Mar. Sci. 32(4):337-339.

Satyamurti, S. J. 1952 a & b. The mollusca of Krusadi Island (in the Gulf of Mannar). I. Amphineura and Gastropoda; II. Scaphapoda, Pelecypoda and Cephalopoda. pp. 466.

Scheltema, R. S. 1974. Biological interactions determining larval settlement of marine invertebrates. Thalassia Jugosl. 10:263-269.

Slattery, M., J. B. McClintock & J. N. Heine. 1995. Chemical defenses in Antarctic soft corals: Evidence for antifouling compounds. J. Exp. Mar. Biol. Ecol. 190:61-77.

Steinberg, P. D., R. De Nys & S. Kjelleberg. 2002. Chemical cues for surface colonization. J. Chem. Ecol. 28(10):1935-1951.

Tang, R. J. & J. J. Cooney. 1998. Effects of marine paints on microbial biofilm development at three materials. J. Ind. Microbiol. Biotechnol. 20:275-280.

Wahl, M. 1995. Bacterial epibiosis on Bahamian and Pacific ascidians. J. Exp. Mar. Biol. Ecol. 191:239-255.

Wahl, M., K. Kroger & M. Lenz. 1998. Nontoxic protections against epibiosis. Biofouling 12(13):205-226.

Wright, A. E. 1998. Isolation of marine natural products. In: R. P. J. Cannell, editor. Methods in biotechnology, vol. 4. Natural product isolation. New Jersey: Humana press Inc. pp. 365-408.

Yamazaki, M. 1993. Antitumor and antimicrobial glycoprotens from seahares. Comp. Biochem. Physiol. 105C:141-146.

Yamazaki, M., J. Kisugi, H. Kimura & D. Mizuno. 1985. Purification of antineoplastic factor from eggs of a seahare. FEBS Lett. 185:295-298.

Yamazaki, M., J. Kisugi, M. Ikenami, H. Kamiya & D. Mizuno. 1984. Cytolytic factor in eggs of the seahare Aplysia kutodai. Gann. 75:269-274.


Suganthi Devadason Marine Research Institute, 44-Beach Road, Tuticorin-628 001, Tamil Nadu, India

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