Inhibition of the development of microorganisms (bacteria and fungi) by extracts of marine algae from Brittany, France. Authors; Authors and affiliations. C. Hellio ...
Appl Microbiol Biotechnol (2000) 54: 543±549
Ó Springer-Verlag 2000
ORIGINAL PAPER
C. Hellio á G. Bremer á A. M. Pons á Y. Le Gal N. Bourgougnon
Inhibition of the development of microorganisms (bacteria and fungi) by extracts of marine algae from Brittany, France
Received: 30 December 1998 / Received revision: 25 April 2000 / Accepted: 1 May 2000
Abstract The inhibitory eects of aqueous, ethanolic and dichloromethane fractions from 16 marine algae from the Atlantic shores of North-East Brittany, France, have been investigated against microorganisms frequently associated with immersed surfaces. The extracts were tested in vitro against isolates of marine fungi, bacteria and yeasts potentially involved at dierent stages in the formation of bio®lms in the sea. The high levels of inhibitory activity of nine extracts against marine fungi and Gram-positive bacteria and their apparent absence of toxicity against larvae of oysters and sea urchins suggests a potential for novel active ingredients.
Introduction Materials immersed in seawater are acted upon by a series of physical, chemical and biological events which result in the formation of a bio®lm complex, including a layer of attached organisms (Characklis 1981; Abarzua and Jakubowski 1995). Immediately after immersion,
C. Hellio (&) á Y. Le Gal Station de Biologie Marine, FRE 2125, MuseÂum National d'Histoire Naturelle-ColleÁge de France, BP 225, 29182 Concarneau, France e-mail: hellio@sb-rosco.fr G. Bremer School of Biological Science, University of Portsmouth, King Henry Building, Portsmouth PO1 2DY, UK A. M. Pons Laboratoire de Genie ProteÂique et Cellulaire, Equipe de Microbiologie, Avenue Marillac, Universite de La Rochelle, 17042 La Rochelle Cedex 01, France C. Hellio á N. Bourgougnon Laboratoire de Biologie et Environnement Marins, Avenue Marillac, Universite de La Rochelle, 17042 La Rochelle Cedex 01, France
surfaces are coated with a glycoproteinaceous ®lm which favours colonisation by bacteria, fungi, diatoms, protozoa and other microorganisms. Macrophytes and invertebrates such as barnacles and blue mussels subsequently attach to this ®lm (Davis et al. 1989). The control of biofouling is of particular concern in modern marine engineering and shipping operations and is one of the most important problems currently facing marine technology (Hattori and Shizuri 1996). Uncontrolled settlement and adhesion of marine invertebrates and algae to the hulls of ships increases frictional drag, with a corresponding decrease in speed, manoeuvrability and fuel eciency. Fouling and corrosion are both major problems in the protection of ships' hulls, cooling systems of power plants, aquaculture systems and other submarine infrastructures. Generally, the development of fouling can be prevented by means of antifouling paints containing one or more toxic compounds such as organotin derivatives in a paint matrix (Vallee-Rehel et al. 1998). However, environmental and human health problems are associated with these metallic compounds (Martin et al. 1981; Gibbs et al. 1987, 1988; Gibbs 1993; Peterson et al. 1993). In response to increasing scienti®c evidence on the toxicity and occurrence of organotin residues from antifouling paints in an aquatic environment, the application of triorganotin antifouling compounds to boats of less than 25 m length has been banned in many countries since 1987 (Voulvoulis et al. 1999). One of the most promising alternatives to heavymetal-based paints is oered by the development of antifouling coatings in which the active ingredients are compounds naturally occurring in marine organisms and operating as natural anti-settlement agents (HoÈlmstrom and Kjelleberg 1994; Kjelleberg and Steinberg 1994). Sessile marine invertebrates such as sponges, ascidians and corals are remarkably free from settlement by fouling organisms. It has been stated (Fusetani 1991) that these organisms secrete chemicals that prevent larvae of other marine organisms from settling and growing on them.
544
Examination of the interactions and competition between species in marine ecosystems has revealed the signi®cant role played by chemicals synthesised by these organisms in mediating these interactions (Bakus et al. 1986; Bazzaz et al. 1986). The surfaces of sessile benthic marine algae are particularly susceptible to fouling because they are restricted to the photic zone where conditions for the growth of fouling organisms are optimal (De Nys et al. 1995). But while some seaweeds are heavily fouled, other species living in the same ecological niche are rarely epiphytised, indicating the possible presence of antifouling mechanisms. Such a process may be solely of a physical nature, but there is some increasing evidence indicating that algae also deter fouling through chemical means. Studies on the antifouling mechanisms utilised by marine sedentary organisms may therefore provide valuable information for fouling control in marine technology. This study is a part of our global research programme committed to ®nding eective natural products against marine bio®lm formation. In this work, a series of aqueous, ethanol and dichloromethane extracts of 16 marine algae from the North-East Atlantic coast of France were tested for their in vitro activity against marine fungi, yeast and bacteria. The microorganisms chosen are representative of the microfouling community and constitute the ®rst colonisers found on immersed surfaces.
Materials and methods Preparation of the extracts Fourteen marine algal species were collected in April 1997 from the East Coast of France (Concarneau Bay, 47°52¢N, 3°55¢W, Brittany); these included Ulva lactuca, Enteromorpha intestinalis (Ulvaceae), Cladophora rupestris (Cladophoraceae), Pelvetia canaliculata, Fucus vesiculosus, Ascophyllum nodosum (Fucaceae), Sargassum muticum (Sargasseae), Laminaria ochroleuca (Laminariaceae), Ectocarpus siliquosus (Ectocarpaceae), Chrondus crispus (Gigartinaceae), Laurencia pinnati®da, Polysiphonia lanosa (Rhodomelaceae), Ceramium rubrum (Ceramiaceae), Cryptopleura ramosa (Delesseriaceae). After collection, the samples were rinsed with sterile seawater to remove associated debris. The cleaned material was then surface-dried by pressing it brie¯y between sheets of paper towel and air-dried in the shade at 30 °C for 24 h. The surface micro¯ora was removed by washing the algal samples for 10 min with 30% ethanol. Chlamydomonas reinhardtii (Chlorophyceae) and Dunaliella tertiolecta (Chlorophyceae) were grown axenically in aerated batch culture at 18 °C under continuous illumination (4,000 lx white ¯uorescent lamps). The growth medium consisted of distilled water or natural seawater, respectively, enriched with Guillard's F2 medium (Guillard and Ryther 1962) deprived of thiamin, biotin and B12 vitamins (Hellio and Le Gal 1999). Mass cultivation for extraction required 5-l batch cultures. Cells were harvested by centrifugation (2,000 g, 20 min, 4 °C) and the resulting pellet was equilibrated for 20 min in 15 ml of extraction buer (Tris 20 mM, pH 7.8, containing 1 mM benzamidine, 5 mM eaminocaproic acid, 0.5 mM dithiotreitol, 200 lM phenyl methyl sulfonyl ¯uoride) and centrifuged again (Hellio and Le Gal 1998). The resultant cell pellet was immediately subjected to extraction procedures. Extracts A (aqueous), B (ethanol), and C (dichloromethane) were obtained as previously described by Hellio et al. (2000).
Evaluation of antibacterial and anti-yeast activities All the microorganisms studied here have been found associated with immersed marine surfaces, including the non-marine species. Bacteria were grown on nutritive agar plates except for Streptococcus sp., which was grown on 5% sheep blood agar. Yeasts were grown on Sabouraud dextrose agar plates. Antibacterial testing of the extracts was performed using a modi®ed well-agar diusion method adapted from Tagg and McGiven (1971), against ®ve Gram-negative bacteria (B1±B5), ®ve Gram-positive bacteria (B6±B10), four marine bacteria isolated from the surface of Mytilus edulis (B11±B14) and ®ve yeast strains (Y1±Y5) (Table 1). The plate assays containing a ®rst layer (10 ml) of medium agar 12% (w/v) were overlaid with a second layer of 5 ml medium agar 6% seeded with 5 ll of a suspension of the target microorganisms (106 cfu/ml). A variety of media were used for the dierent microorganisms studied: for bacteria (B1±B10) the ®rst agar layer was composed of BHI (brain±heart infusion; Difco, France) and the second layer of M63 (Miller 1972); for yeasts (Y1± Y5), Sabouraud (Difco) was used; for marine bacteria (B11±B14), Marine Broth Agar (Sigma, France) was used. Sterile glass rings (4 mm internal diameter) were placed on this bilayered agar. Extracts (30 lg) diluted in DMSO 5% and ®ltered (Millex-GV unit 0.22 lm Millipore pore size) were placed in the glass wells and allowed to diuse for 2 h at 4 °C. After incubation for 18 h at 37 °C for B1±B6 and B9±B10, at 30 °C for B7±B8 and Y1±Y5, and at 20 °C for B11±B14, the activity was evaluated by measuring the diameter (D, mm) of the inhibition zones around the rings. The assay was scored as follows: highly positive (+++) if D was >10 mm, positive (++) if D was 8±10 mm, weakly positive (+) if D was 4±8 mm, and (±) for no activity. Control tests with the solvents were performed for every assay and showed no inhibition of the microbial growth. In addition, the biocides TBTO (1 ppm) and CuSO4 (1 ppm) were used to check the sensitivity; these two substances are present in the formulation of commercial antifouling coatings. All inhibition assays were carried out in duplicate. Evaluation of antifungal activity The antifungal activity of the extracts was tested against ®ve strains of marine fungi (F1±F5) (Table 1) using an agar diusion method. F1 and F2 are unidenti®ed marine fungi isolated respectively from sand collected from Malaysia and from driftwood collected from the Fleet estuary, UK. F3, F4 and F5 were isolated from driftwood collected respectively from Dinas (Wales), Denmark and Galway (Eire). The fungal strains were maintained on maize meal agar (Oxoid) slopes. Thirty micrograms of each sample was diluted in DMSO 5% and ®ltered (Millex-GV unit 0.22 lm Millipore pore size) and then incorporated into 6 ml of maize meal agar 12%, pH 6 (Sigma). Then the plate assay was inoculated aseptically at the centre with an 8 mm diameter agar plug of mycelia. After incubation at 25 °C for 1 week for F1 and F3, 3 weeks for F2, and 4 weeks for F4 and F5, the activity was evaluated by measuring the diameter (mm) of the fungal colonies. Control tests containing the solvents loaded in the rings were performed for every assay and showed no inhibition of fungal growth. In addition, standard glass rings of TBTO (1 ppm) and CuSO4 (1 ppm) were used to check the sensitivity. All inhibition assays were done in duplicate. Determination of the minimum inhibitory concentrations Determination of the minimum inhibitory concentrations (MICs) was carried out for the bacteria (National Committee for Clinical Laboratory Standards 1993) and fungi (Shadomy et al. 1985) by the macrodilution method for testing the antimicrobial activity of the algal extract. The extracts concentration-tested were 96, 48, 32, 16, 8 and 4 lg/ml. Microorganisms (2 ´ 108 cfu/ml) were placed in a liquid medium consisting either of MH (Mueller Hinton) medium for B3, B6±B10 and Y1±Y2 (RPMI 1,640 with L-glutamine buffered to pH 7 with 0,165 MOPS buer), or marine broth medium
545 Table 1 List of microorganisms used for our experiments B1 B2 B3 B4 B5 B6 B7 B8 B9 B10 B11 B12 B13 B14 Y1 Y2 Y3 Y4 Y5 F1 F2 F3 F4 F5
Organism
Characteristics
Strain
Escherichia coli K12 Klebsiella pneumoniae Serratia marcescens Proteus vulgaris Pseudomonas aeruginosa Bacillus subtilis Bacillus cereus Bacillus megaterium Streptococcus sp. Staphylococcus aureus Unidenti®ed marine bacteria Unidenti®ed marine bacteria Unidenti®ed marine bacteria Unidenti®ed marine bacteria Candida brusei Candida albicans Saccharomyces cerevisiae Candida tropicalis Issatchenkia orientalis Unidenti®ed marine fungus Unidenti®ed marine fungus Corollospora maritima Lulworthia sp. Dendryphiella salina
Gram-negative Gram-negative Gram-negative Gram-negative Gram-negative Gram-positive Gram-positive Gram-positive Gram-positive Gram-positive Isolated mixed Gram-positive Isolated mixed Gram-negative Isolated Gram-positive Isolated Gram-negative
ATCC 23176a CIP 53.153b CIP 67.55 LC 0006c ATCC 27853 CIP 5262 LC 0035 CIP 6620T CIP 55120 ATCC 25923 LC 18 LC 28 LC 35 LC 36 LC 0001 LC 002 LC 0003 DSM 1346d DSM 6128 LC LC LC LC LC
Isolated Isolated Isolated Isolated Isolated
a
ATTC: American Type Culture Collection CIP: Collection de l'Institut Pasteur c LC: Laboratory Collection d DSM: Deutsche Sammlung von Mikroorgansmen b
for B13, or maize meal medium for F1±F5, containing the algal extracts for testing. After incubation for 20 h at 37 °C for B3, B6, B9 and B10; 20 h at 30 °C for B7, B8, Y1 and Y2; 20 h at 20 °C for B13 and 2 days at 25 °C for the marine fungi, MIC represents the lowest concentration that inhibits the organism's growth. Toxicity tests on larval oyster (Crassostrea gigas) and larval sea urchin (Echinus esculentus) The larvae used came from adult animals collected from the sea. Oysters came from the Belon river (15 July 1999, Brittany, France) and the sea urchins were collected in the Glenans Islands (10 August 1999, Brittany, France). The oysters' gametes were obtained after several thermal shocks (18 °C and 30 °C; Calabrese et al. 1973) and 2 ml of a dense suspension of sperm was mixed with 200 ml of a suspension of eggs. After a contact period of 45 min, the absence of polyspermy was veri®ed by microscopic observation. Fertilisation occurred during a 3-h period, at 18 °C, in natural sterile seawater (®ltrated at 0.2 lm). The larval concentration was determined by microscopic observation using a haemocytometer (Malassez Cell) and a concentration of 100 larvae/ml (Martin et al. 1981) was employed for each test. The same protocol was used for sea urchin larvae. In order to determine the toxicity of the algal extract on the larval development of oysters and sea urchins, larvae were incubated with six dierent concentrations of algal extracts: 10, 50, 125, 250, 500 and 1,000 lg/ml. TBTO and CuSO4 (1, 10, 25, 50, 75 and 100 lg/ml) were used as positive controls. Five replicates were used in order to reduce the eect of the natural variability between individuals. Results are expressed as an average of the replicate results. A population control comprised larvae reared in seawater. After 24 h at 24 °C, larvae reached the D stage (con®rmed by microscopic observation). Larval development was then stopped by the addition of 100 ll of formol to the culture medium. The rate of mortality is determined by the formula: MR (NL0 ) NLV24)/ NL0 ´ 100 with MR being the mortality rate, NL0 the number of larvae in the medium at the hour 0 and NLV24 the number of viable larvae present in the medium after 24 h incubation.
Results Crude extracts of the 16 marine algae tested against bacteria, yeasts and fungi, showed varied antimicrobial activity. The results of the primary screening tests are summarised in Table 2. Although these compounds exhibited a broad-spectrum activity as a group, there were important variations in the eects of the dierent extracts on dierent fouling microorganisms. Aqueous extracts (extract A) did not show inhibition of the growth of bacteria, yeasts and fungi (results not shown). On the other hand, crude ethanolic and dichloromethane fractions appeared to be speci®c, in particular against the growth of the Gram-positive bacteria tested. No signi®cant dierence in activity against dierent microorganisms was observed between ethanol extract (extract B) and dichloromethane extract (extract C) for each alga. Of the 48 algal extracts tested, none gave any inhibition against the following strains: Escherichia coli K12 (B1), Klebsiella pneumoniae (B2), Pseudomonas aeruginosa (B5), Gram-negative marine bacteria (B12 and B14), Saccharomyces cerevisiae (Y3), Candida tropicalis (Y4) and Issatchenkia orientalis (Y5). Eleven extracts exhibited high levels of activity against Gram-positive bacteria and fungi and these were at the same levels as positive controls of CuSO4 or TBTO (1 ppm). Comparable inhibition zones were only obtained for Enteromorpha intestinalis (extract B), Laurencia pinnati®da (extracts B and C) and Polysiphonia lanosa (extracts B and C), which showed strong inhibition of the Gram-positive bacteria. Chlamydomonas
546 B13 (Gram-positive bacteria), Y1 (C. brusei), Y2 (C. albicans), F1 (unidenti®ed marine fungus), F2 (unidenti®ed marine fungus), F3 (C. maritima), F4 (Lulworthia sp.) and F5 (D. salina). No activity was recorded against B1 (E. coli K12), B2 (K. pneumoniae), B4 (P. vulgaris), B5 (P. aeruginosa), B12 (mixed Gram-negative marine bacteria), B14 (Gram-negative marine bacteria), Y3 (S. cerevisiae), Y4 (C. tropicalis) and Y5 (I. orientalis)
Table 2 Bioassay for antimicrobial activity. Level of activity is denoted by +++ highly positive (10±12 mm zone of inhibition), ++ positive (8±10 mm zone of inhibition), + weakly positive (4±8 mm zone of inhibition) and ) negative (no zone of inhibition). Algal extracts B and C represent the ethanol and the dichloromethane fractions, respectively. B3 (S. marcescens), B6 (B. subtilis), B7 (B. cereus), B8 (B. megaterium), B9 (Streptococcus sp.), B10 (S. aureus), B11 (mixed Gram-positive marine bacteria), Organism
Extract B3
B6
B7
B8
B9
B10
B11
B13
E. intestinalis C. reinhardtii C. reinhardtii S. muticum S. muticum L. ochroleuca A. nodosum A. nodosum C. crispus L. pinnati®da L. pinnati®da P. lanosa P. lanosa C. ramosa DMSO 5% TBTO 1 ppm CuSO4 1 ppm
B B C B C B B C B B C B C C
++ ) ) ) ) ) ) ) ) + +++ ++ + ) ) +++ +++
+ ) ) ) ) ) ) ) ) ++ ++ ++ ++ ) ) +++ +++
+++ ) ) ) ) ) ) ) ) + ++ +++ ++ ) ) +++ +++
+ ) ) ) ) ) ) ) ) + ++ + ++ ) ) +++ +++
+ ) ) ) ) ) ) ) ) + +++ ++ ++ ) ) +++ +++
++ ) ) ) ) ) ) )
+++ ) ) ) ) ) ) )
) ) ) ) + ) ) ) ) ) ) ) + ) ) +++ +++
+ ) ) ) ) ) ) ) + + + + ++ ++ + +++ ++ ) ++ +++ ) ) ) ) ) ) ) +++ +++ +++ +++ +++ +++
Organism
Extract
E. intestinalis C. reinhardtii C. reinhardtii S. muticum S. muticum L. ochroleuca A. nodosum A. nodosum C. crispus L. pinnati®da L. pinnati®da P. lanosa P. lanosa C. ramosa
B B C B C B B C B B C B C C
B3
Y2
F1
F2
F3
F4
F5
) + ) ) ) ) ) ) ) ) ) ) ) ) ) +++ +++
) ) ) ++ ) +++ +++ ) ++ ) ++ ) ) +++ ) +++ +++
) ) ) ++ ) +++ +++ ) ++ ++ ) ) ) ) ) +++ +++
) ) ++ ) ) ) ++ +++ +++ +++ +++ ) ) +++ ) +++ +++
) ) ++ + ) ++ ++ + +++ ++ ++ ) ) ++ ) +++ +++
) ) ++ + ) + +++ ++ ++ ++ ++ ) ) ++ ) +++ +++
extracts B and C of Ascophyllum nodosum, extract B of Chondrus crispus and extract C of Cryptopleura ramosa showed MICs of only 24 lg/ml for the inhibition of at least two strains of marine fungi (Table 3). Toxicity tests against non target species were investigated for those extracts showing the highest levels of antimicrobial activity: extracts B of Ascophyllum nodosum, Chondrus crispus, Enteromorpha intestinalis, Laminaria ochroleuca, Laurencia pinnati®da and Polysiphonia lanosa, and extracts C of A. nodosum, Cryptopleura ramosa, L. pinnati®da and P. lanosa. Results are shown in Fig. 1 (toxicity on oyster larvae) and Fig. 2 (toxicity on sea urchin larvae). TBTO and CuSO4, used as positive controls, were found to be very toxic to the larvae studied and no viable larvae were detected at 50 lg/ml concentrations. In contrast, the algal extracts appeared to be non-toxic to invertebrate larvae even at concentrations up to 1,000 lg/ml.
reinhardtii (extract C), Sargassum muticum (extract B), Laminaria ochroleuca (extract B), Ascophyllum nodosum (extracts B and C), Chondrus crispus (extract B) and Cryptopleura ramosa (extract C) gave high levels of antifungal activity. Table 2 shows that some marine fungi were particularly sensitive to the extracts B and C of Laurencia pinnati®da. In contrast to the ethanol extract of E. intestinalis, which did not display activity against marine fungi, the dichloromethane extract of L. pinnati®da exhibited broader activity and was active against 12 tested organisms including several isolates of marine fungi. All the extracts that exhibited activity against Gram-positive bacteria showed strong inhibitory activity (Table 3). In particular, strong antibacterial activity was observed for extract B of Enteromorpha intestinalis, extracts B and C of Polysiphonia lanosa and extract C of Laurencia pinnati®da. Moreover, extract B of Laminaria ochroleuca, Table 3 Minimum inhibitory concentration (MIC) in lg/ml. Algal extracts B and C represent the ethanol and the dichloromethane fractions, respectively. B3 (S. marcescens), B6 (B. subtilis), B7 (B. cereus), B8 (B. megaterium), B9 (Streptococcus sp.), B10 (S. aureus), B13 (Gram-positive bacteria), Y1 (C. brusei), Y2 (C. albicans), F1 (unidenti®ed marine fungus), F2 (unidenti®ed marine fungus), F3 (C. maritima), F4 (Lulworthia sp.) and F5 (D. salina)
Y1
B6
B7
B8
B9
B10
B13
Y1
48
96
24
96
96
24
96
96
96
96 24 48 96
48 48 48 48
96 48 24 48
96 48 96 48
96 24 48 48
96 48 48 24
96 96 96
Y2
F1
F2
48
48
24 24
24 24
24
24 48
96
48 24
F3
F4
F5
48
48 96
48 96
48 24 48 24 24
48 48 24 96 48 48
24 96 48 48 48 48
24
48
48
547 Fig. 1 Survival rate of larval oysters at dierent concentrations (1±1,000 lg/ml) of algal extracts
Fig. 2 Survival rate of larval sea urchins at dierent concentrations (1±1,000 lg/ml) of algal extracts
Discussion Marine fungi, bacteria and yeast are major organisms involved in the formation of the microlayer which is the ®rst step in the process of fouling (HoÈlmstrom and Kjellberg 1994). This primary bio®lm also provides a supporting substrate for the subsequent attachment of other fouling organisms; the deterrence of the formation of this layer is fundamental to eective control of further large-scale biofouling. In addition, marine bacteria and marine fungi are associated with corrosion products (Little et al. 1999).
The bio®lm deterrence strategy is seen as a viable alternative to chemical treatments of established colonies using bleaches and/or detergents or the provision of toxic surfaces, e.g. copper or organotin additives to paints to provide deterrent surfaces against primary colonisers. The antifouling agents incorporated into deterrent surfaces, if derived from naturally occurring substances, may be less environmentally damaging than the toxins currently employed and may have less activity against non-target species. Although marine macroalgae are already well documented as possessing generalised antibacterial and antifungal activity (Caccamese et al. 1980, 1985; Navqi
548
et al. 1980; Padma Sridhar et al. 1984; Pesando and Caram 1984; Bernard and Pesando 1989; Vlachos et al. 1996), there is also data available on their speci®c activity against fouling bacteria (Devi et al. 1997). Several pure active compounds have been isolated and their structure determined. For example, phlorotannins from brown algae have been implicated in antifouling (Sieburth and Conover 1965; McLachlan and Craigie 1966; Langlois 1975; Fletcher 1989). Philipps and Towers (1982) suggest that the brominated phenol lanosol exuded by the red alga Rhodomela larix may act as an antifoulant. De Nys et al. (1995) showed that the red alga Delisea pulchra produced a suite of unique secondary metabolites, furanones, that inhibit surface colonisation traits in epiphytic marine bacteria without toxicity. These compounds also speci®cally regulate bacterial fouling of the algal surface by interfering with lactone regulatory systems. KoÈnig and Wright (1997) show that the marine red algae Laurencia rigida remains relatively free from fouling organisms; this feature, coupled with their results in screening the dichloromethane extracts in antifouling bioassays, indicates a chemical deterrence of common marine fouling organisms, especially fungi, by the algal products. Considering the importance of microorganisms in the early stages of marine fouling, Ina et al. (1989) have studied the correlation between antibacterial and antifouling activities. They examined the antibacterial activity of compounds against Escherichia coli (Gramnegative), Staphylococcus aureus (Gram-positive) and Bacillus subtilis (Gram-positive). Compounds showing antifouling activity for as long as 60 days exhibited antimicrobial activity against the Gram-positive bacteria (Staph. aureus and B. subtilis) but were inactive against E. coli (Gram-negative). With few exceptions, antibacterial activity against Gram-positive bacteria seems to be a prerequisite for an eective antifouling compound. Devi et al. (1997) considered that such a preliminary screening method had the advantage of being simple, less time-consuming and requiring only small quantities of the materials for assays, thereby ensuring minimal removal of biodiversity from the environment. The method described is likely to be a suitable way forward as attempts are made to establish a simple, eective bioassay system for ®nding novel antifouling agents. Of the 48 algal extracts tested in this study against marine fungi, yeasts and marine bacteria, 12 extracts showed antimicrobial activity at low concentrations. Among the extracts obtained from the screening programme, the strong antibacterial activity and antifungal activities of the dichloromethane extract from Laurencia pinnati®da is worthy of further investigation. This extract inhibited activity against the Gram-positive bacteria, the marine bacteria and fungi with an inhibition zone between 3 and 12 mm. Extract B from Enteromorpha intestinalis and extracts B and C from Polysiphonia lanosa proved to be particularly eective against
the Gram-positive and marine bacteria. Moreover, these active extracts appeared to be non-toxic to oyster and urchin larvae in contrast with TBTO and CuSO4. These results con®rm that marine algae are a potential source of bioactive compounds against colonising microorganisms, which as natural non-toxic products may be more acceptable to legislators and environmental agencies. Upsetting the development of the microfouling community by non-polluting natural marine antifouling compounds may be the most desirable way of breaking the fouling chain and thus indirectly reducing the macrofouling settlement on man-made structures. There is also the potential for incorporating such compounds into broad-spectrum antifouling coatings containing a mixture of compounds, each of which will be most eective against a particular group of fouling organisms. The advantage of the utilisation of micro- and macroalgae for the isolation of bioactive agents is that algae can be cultivated in a short time in mass culture, independent of season. Furthermore, they are capable of manipulation to optimise the production of biogenic agents (Abarzua and Jakubowski 1995). This work is included in a general programme for ®nding new natural antifouling compounds extracted from algae. Our previous results indicate that extracts B and C of E. intestinalis; extracts B and C of S. muticum and extract B of P. lanosa speci®cally inhibited the activity of the phenoloxidase (an enzyme involved in mussel attachment by byssus formation) puri®ed from the foot of the mussel Mytilus edulis (Hellio et al. 2000). This con®rms that algal extracts can act at dierent stages of the fouling process, including micro- and macrofouling. However, further work is needed to identify these active compounds, to evaluate speci®c antimicrobial activity against the marine bacteria speci®cally implicated in biofouling and to examine the precise role of such activity in nature. Acknowledgements This work was supported by The Brittany Council and an European studentship. We thank Dr. Alain Plusquellec (IUT, Quimper, France) and Julie Clipston (School of Biological Sciences, University of Portsmouth, UK) for providing us with the strains of marine bacteria and marine fungi. Special thanks are given to Dr. Denis Fichet and Dr. Gilles Radenac (LBEM, University of La Rochelle, France) for their advice concerning the larval toxicity tests. We also thank Dr. Jean Pascal Berge (Ifremer, Nantes, France) for his helpful comments and criticisms of this manuscript.
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