Inhibition of marine bacteria by extracts of macroalgae: potential use for environmentally friendly antifouling paints

Marine Environmental Research 52 (2001) 231±247 www.elsevier.com/locate/marenvrev

Inhibition of marine bacteria by extracts of macroalgae: potential use for environmentally friendly antifouling paints C. Hellio a,b,*, D. De La Broise c, L. Dufosse c, Y. Le Gal a, N. Bourgougnon b a

FRE 2125 CNRS-MNHN-UBO, Station de Biologie Marine, MuseÂum National d'Histoire Naturelle-ColleÂge de France, BP 225, 29182 Concarneau, France b Laboratoire de Biologie et Environnement Marins, Avenue CreÂpeau, Universite de La Rochelle, 17042 La Rochelle cedex 01, France c Laboratoire de Microbiologie AppliqueÂe EA 2651, Universite de Bretagne Occidentale, site IUP-IIA, PoÃle Universitaire de Creac'h Gwen, 29000 Quimper, France Received 5 September 2000; received in revised form 10 December 2000; accepted 15 December 2000

Abstract Although a total ban on the use of TBT coatings is not expected in the short term, there is a growing need for environmentally safe antifouling systems. A search for new non-toxic antifoulants has been carried out among marine macroalgae. Antifouling activity of aqueous, ethanolic and dichloromethane extracts from 30 marine algae from Brittany coast (France) was examined in vitro against 35 isolates of marine bacteria. About 20% of the extracts were found to be active. The high levels of inhibitory activities against bacteria recorded in some extracts and the absence of toxicity on the development of oyster and sea urchin larvae and to mouse ®broblast growth suggests a potential for novel active ingredients in antifouling preparations. # 2001 Elsevier Science Ltd. All rights reserved. Keywords: Algal extract; Antibacterial activity; Antifouling; Bio®lm; Marine bacteria; Natural product; Toxicity

* Corresponding author. Present address: Department of Marine Sciences and Coastal Management, Ridley Building, Newcastle University, Newcastle upon Tyne, NE1 7RU, UK. Fax: +44-191-222-7891. E-mail addresses: [email protected] (C. Hellio), nbourgou@univ- lr.fr (N. Bourgougnon). 0141-1136/01/$ - see front matter # 2001 Elsevier Science Ltd. All rights reserved. PII: S0141-1136(01)00092-7

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1. Introduction Vessels of all sizes, both commercial and recreational, use antifoulant paints to control the biological growth of target organisms on submerged surfaces. Bacteria are the ®rst organisms to foul surfaces exposed to sea water (Little, 1984). Their subsequent multiplication and exopolymer production lead to the formation of bio®lms (Abarzua & Jakubowski, 1995; Characklis, 1981) which are believed to in¯uence the settlement of subsequent colonisers in the fouling process (Davis, Targett, McConnell, & Young, 1989). The formation of slime ®lms attached to submerged structures has important economic and ecological implications. It destroys or alters the toxic property of the paint ®lms and acts as attractant to higher fouling organisms. Many macrofoulers prefer slimed surfaces and can discriminate between slimed and clean surfaces on their pre-settling stage (Crisp, 1984; Little, 1984). Moreover, many sedentary organisms need a slimed surface to undergo ®nal metamorphosis and attachment. Thus, the presence of a primary slime surface attracts a large number of fouling organisms. In this way, bacterial attachment can indirectly in¯uence the fouling and corrosion of man-made surfaces (Corpe, 1970). Fouling of ships results in increased frictional resistance leading to loss of speed, fuel eciency is then decreased and consequently pollution increases (Hall, Giddings, Soloman, & Balcomb, 1999). The fouling of the hulls of ships is a long-standing problem and a wide variety of technologies have been applied to reduce fouling (Hall et al., 1999). Research in the 1960s showed the e€ectiveness of organotin as antifoulants (Rexrode & Spatz, 1997). Subsequently, however, tri-substituted organotin compounds were found to be highly toxic to non-target aquatic life and to persist and to bio-accumulate (Fent & Hunn, 1997). This can pose risks to aquatic ecosystems and through human health food chain bioaccumalation and biomagni®cation (Boyer, 1989; Yamamoto, 1994). The use of tributyltin (TBT) as an active ingredient in antifouling paints has been progressively restricted since 1982 when France was ®rst to ban this compound to protect oyster-growing areas (Tolosa, Readman, Balevoet, Ghilini, Bartocci, & Horvat, 1996). One of the most promising alternatives to heavy metal based paints is o€ered by the development of antifouling coatings in which the active ingredients are compounds naturally occurring in marine organisms and operating as natural antisettlement agents (HolmstroÈm & Kjelleberg, 1994; Kjelleberg & Steinberg, 1994). Sessile marine macroalgae are remarkably free from settlement by fouling organisms. 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, Steinberg, Willemsen, Dworjanyn, Gabelish, & King, 1995). But while some marine algae are heavily fouled, other species living in the same ecological niche are rarely epiphytized, indicating the possible presence of antifouling mechanisms (Hellio, Bourgougnon, & Le Gal, 2000a; Hellio, Bemer, Pons, Le Gal, & Bourgounon, 2000b). There have been a number of reports of antibacterial activity from marine plants (Bernard & Pesando, 1989; Burkholder, Burkholder, & Almodovar, 1960; Caccamese, Azzolina, Furnari, Coraci, & Grasso, 1980; Caccamese, Toscano, Furnari, &

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Cormaci, 1985; Devi, Solimabi, D'Souza, Sonak, Kamat, & Singbal, 1997; Hellio et al., 2000b; Navqi et al., 1981; Pesando & Caram, 1984). However, few data are available on the activity of marine algae against fouling bacteria (Devi et al., 1997). As part of the ongoing research programme to look for biologically active substances from marine algae against marine bio®lm formation, aqueous, ethanolic and dichloromethane extracts of 30 algae from the North East Atlantic Coast (France) were tested for their in vitro antibacterial activity. Upsetting the development of the microfouling community by non-polluting marine natural antifouling compounds may be the most desirable way of breaking the fouling chain and so indirectly reducing the macrofouling settlement on man-made structures. 2. Materials and methods 2.1. Preparation of the extracts Thirty marine algal species were collected in April 1998 from the East coast of France (Concarneau Bay, Brittany, 47 52N±3 55W) these included: (1) Enteromorpha intestinalis; (2) Ulva lactuca (Ulvales); (3) Cladophora rupestris (Cladophorales); (4) Ascophyllum nodosum; (5) Fucus serratus; (6) F. spiralis; (7) F. vesiculosus; (8) Himanthalia elongata; (9) Pelvetia canaliculata; (10) Sargassum muticum (Fucales); (11) Ectocarpus siliculosus (Ectocarpales); (12) Alaria esculenta; (13) Chorda ®lum; (14) Laminaria digitata; (15) L. ochroleuca; (16) Saccorhiza polyschides (Laminariales); (17) Chondrus crispus; (18) Gigartina stellata (Gigartinales); (19) Gelidium latifolium (Gelidiales); (20) Palmaria palmata (Palmariales); (21) Dilsea carnosa (Cryptonemiales); (22) Bornetia secundi¯ora; (23) Ceramium rubrum; (24) Cryptopleura ramosa; (25) Delesseria sanguinea; (26) Grithsia ¯oculosa; (27) Halurus equisetifolius; (28) Laurencia pinnati®da; (29) Plumaria elegans; (30) Polysiphonia lanosa (Ceramiales). After collection, the samples were rinsed with sterile seawater to remove associated debris. Epiphytes were removed of the algae. The cleaned material was then surface dried by pressing it brie¯y between sheets of paper towelling and air dried in the shade at 30 C during 24 h. The surface micro¯ora was removed by washing the algal samples for 10 min with 30% ethanol (Hellio et al., 2000a). One part of the dried algae was suspended by stirring in distilled water (50 g/l of dried weight) with an Ultra-turrax (2 h) at 4 C. After centrifugation (30 min, 30 000 g, 4 C) and ®ltration (Whatman cat no. 1 822 047), the supernatant was lyophilised and the aqueous extract was obtained (Extract A). For organic extracts, the dried algae was suspended by stirring in ethanol 95 (200 g/300 ml). After centrifugation (30 min, 30 000 g, 4 C), the resultant pellet was reextracted ®ve times in the same way. The alcoholic extracts were combined and evaporated under vacuum at low temperature (<40 C). Distilled water (100 ml) was then added and partitioned with methylene chloride (4100 ml). The aqueous phases were collected, lyophilised, re-suspended in absolute ethanol (100 ml), ®ltered and concentrated under vacuum at low temperature (Extract B). The organic phases

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were collected, dried during 24 h under Na2SO4, ®ltered and concentrated under vacuum at low temperature (Extract C). These three phases were stored at 40 C before use (Hellio et al., 2000a). 2.2. Collection of marine bacteria Thirty-®ve marine bacterial strains were obtained from the Culture Collection of the University of Quimper (LUMAQ; France; Table 1).

Table 1 Characteristics of the marine bacteria Sample number

Date

Position

Sample composition

shape

Gram

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35

25/05/1998

47 430 95N 4 010 85W

25/05/1998 25/05/1998 25/05/1998

47 430 95N 4 010 85W 47 430 95N 4 010 85W 47 420 64N 4 000 85W 47 420 64N 4 000 85W

Rod Rod Coccus Diplococcus Rod Coccobacillus Rod Coccobacillus

+ +

25/05/1998 25/05/1998

47 420 45N 3 59 98W

06/07/1998

47 420 44N 4 020 95W

06/07/1998

47 420 54N 4 010 65W

07/10/1998

47 430 07N 3 570 17W

07/10/1998 07/10/1998

47 430 11N 3 570 13W 47 420 90N 3 570 23W

07/10/1998

47 430 02N 3 570 13W

Enteromorpha sp. Enteromorpha sp. Dilsea carnosa Dumontia sp. Gelidium corneum Gelidium corneum Sand Sand Sand Sand Sand Gigartina sp. Gigartina sp. Corallina sp. Corallina sp. Corallina sp. Decaying algae mixture Decaying algae mixture Decaying algae mixture Decaying algae mixture Enteromorpha sp. Cladophora rupestris Cladophora rupestris Laminaria sp. Laminaria sp. Laminaria sp. Laminaria sp. Ulva sp. Ulva sp. Ulva sp. Ulva sp. Ulva sp. Green algae in highly salt water Green algae in highly salt water Green algae in highly salt water

07/10/1998

47 420 98N 3 570 10W

07/10/1998

47 420 98W 3 570 24W

Diplococcus Rod Rod Rod Rod Rod Rod Rod Coccus Rod Rod Rod Rod Rod Rod Rod Rod Coccus Rod Rod Rod Rod Rod Coccus Rod Long rod

+ + + + +

+ + + + + + + + +

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2.3. Evaluation of antibacterial activities Antibacterial testing of the extracts was performed by the disc di€usion technique in agar plated petri dishes (Devi et al., 1997). Whatman ®lter paper discs (6 mm diameter) were initially sterilised at 10104 Pa pressure for 15 min. A sample consisting of algal extracts (30 mg) diluted in DMSO 5% and ®ltered (Millex-GV unit 0.22 mm Millipore pore size) was loaded on each of these discs and allowed to dry at room temperature during 3 h. Discs were then placed on agar dishes. Bacterial cultures were grown in liquid DIFCO 2216 marine broth overnight, and 0,1 ml samples of the culture (106 CFU/ml) were spread over the agar. After incubation for 4 days at 20 C, the activity was evaluated by measuring the diameter (D, in mm) of the inhibition zones around the discs. Control tests with the solvents were performed for every assay and showed no inhibition of the microbial growth. In addition, the biocides TBTO (bis (tributyltin) oxide; 1 ppm) and CuSO4 (copper sulfate; 1 ppm) were used as positive controls to check the sensitivity (Hellio et al., 2000b). All inhibition assays were carried out in duplicate. 2.4. Determination of the minimum inhibitory concentrations (MICs) Determination of MICs were made by the macrodilution method (NCCLS, 1993) for testing the antimicrobial activity of the algal extract. Extract concentrations tested were 96, 48, 32, 16, 8 and 4 mg/ml. Micro-organisms (2.108 CFU/ml) were placed in a liquid medium consisting in DIFCO 2216 marine agar containing the algal extracts to test for an incubation of 48 h at 20 C. MIC represents the lowest concentration that inhibits the organism's growth. 2.5. Cytotoxicity determination:continuous drug exposure 2.5.1. Cell culture and treatment The 3T3 cells line (ECACC 8611040, mouse ®broblasts) were plated in 96-well plates in minimum essential medium Eagle (MEM) supplemented with 10% foetal bovine serum (FBS, Eurobio) which contained Penicillin (100 mg/ml), sodium pyruvate (1 mM) and l-glutamine (2 mM). 2.5.2. Cytotoxicity tests The cytotoxic e€ects of the biocides (TBTO and CuSO4) and of the algal extracts were assessed after 48 h of exposure, using the Neutral Red and the MTT tests. These assays were performed as previously described (Fautrel et al., 1991). Cytotoxicity data were standardised as the percentage of control cell viability as previously described (De Sousa et al., 1998). 2.6. Toxicity tests on larval oyster (Crassostrea gigas) and larval sea urchin (Echinus esculentus) Toxicity tests were realised as previously described (Hellio et al., 2000b).

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3. Results 3.1. Antibacterial activity Ninety crude extracts of 30 marine algae were tested against 35 strains of potentially marine fouling bacteria. The results of the screening tests are summarised in Table 2. Among the 90 algal extracts tested, only 18 extracts showed antibacterial activities, these are Enteromorpha intestinalis (1B), Cladophora rupestris (3C), Ascophyllum nodosum (4B, 4C), Sargassum muticum (10B, 10C), Ectocarpus siliculosus (11B, 11C), Laminaria ochroleuca (15B), Chondrus crispus (17B), Gelidium latifolium (19C), Palmaria palmata (20C), Ceramium rubrum (23C), Cryptopleura ramosa (24C), Laurencia pinnati®da (28B, 28C), Polysiphonia lanosa (30B, 30C). Aqueous algal extracts (A) did not show any inhibition of the growth of marine bacteria (results not shown). On the other hand, crude ethanolic and dichloromethane fractions appeared to be speci®c against growth of marine bacteria. No signi®cant difference was observed in activity against marine bacteria between ethanol extracts (Extract B) and dichloromethane extracts (Extract C) for each alga. Among the 18 active extracts, three extracts (3C, 19C and 20C) showed speci®c inhibition against only Gram-negative bacteria (17, 18, 19, 24, 25, 27), 12 extracts (1B, 4C, 10C, 11B, 11C, 15B, 17B, 23C, 24C, 28B, 28C and 30B) showed speci®c inhibitory activities against only Gram-positive bacteria (1, 2, 12, 13, 20, 22, 23, 26, 29, 30, 32). Three extracts (4B, 10B and 30C) inhibited simultaneously the growth of Gram-positive and Gram-negative marine bacteria. Among the 35 marine bacteria strains tested, 17 strains showed some growth inhibition by algal extracts. In contrast, ®ve strains (14, 15, 16, 34 and 35) of Grampositive bacteria and 13 strains of Gram-negative bacteria (3, 4, 5, 6, 7, 8, 9, 10, 11, 21, 28, 31 and 33) were not inhibited. TBTO and CuSO4 used as positive controls, were found to be very toxic (>5±6 mm zone of inhibition) against the growth of the most strains of bacteria tested. In contrast three strains (1, 2 and 22) showed tolerance to these reagents. Similar levels of inhibition were found in some of the algal extracts. DMSO 5% was used as control and showed no inhibition of the growth of the marine bacteria tested. MICs were performed for the algal extracts exhibiting the higher levels of activity (Table 3). Some crude extracts showed MICs at concentrations as low as 24 mg/ml. The ethanolic fraction of Ascophyllum nodosum (4B) is the most active extract against 10 strains of marine bacteria with a MIC of 24 mg/ml. The dichloromethane fraction of Polysiphonia lanosa (30C) lead to the inhibition of seven sensitive strains of bacteria at 24 mg/ml. The dichloromethane fraction of Ectocarpus siliculosus (11C), ethanolic fraction of Chondrus crispus (17B) and dichloromethane fraction of Laurencia pinnati®da (28C) lead to the inhibition of six of the sensitive strains at a concentration as low as 24 mg/ml. The results of the MIC studies show that the most sensitive strains of bacteria: strains number 13, 23 and 29 are the most sensitive to the algal extracts, as they are each inhibited by eight di€erent extracts at a concentration of 24 mg/ml. The MIC levels for the biocides are for the most part of strain of 6 mg/ml, except for the strains 1, 2 and 23 which exhibited MIC >96 mg/ml.

Table 2 Bioassay for the bactericidal activity of algal extractsa 2

12

13

++ ± ++ ++ ++ ++ ++ + ++ ++ ± ± ++ + + ++ ++ ++

++ ± ++ ++ ++ ++ ++ + ++ ++ ± ± ++ + + ++ ++ ++

+++ ± +++ +++ ++ ++ + +++ + +++ ± ± +++ ++ + +++ ++ ++

+++ ± +++ ++ ++ +++ + +++ +++ +++ ± ± ++ ++ + +++ ++ +++

TBTO ± CuSO4 ±

±

±

17 ± ++ ++ ± ++ ± ± ± ± ± ++ ++ ± ± ± ± ± ++

18

19

± ± +++ +++ +++ +++ ± ± ++ ++ ± ± ± ± ± ± ± ± ± ± ++ ++ +++ ++ ± ± ± ± ± ± ± ± ± ± +++ +++

20 ++ ± ++ ++ ++ ++ ++ + ++ ++ ± ± ++ + + ++ ++ ++

22 ++ ± ++ ++ ++ ++ ++ + ++ ++ ± ± + + + ++ ++ ++

23

24

+++ ± ± +++ +++ +++ ++ ± ++ ++ +++ ± + ± +++ ± + ± +++ ± ± ++ ± +++ +++ ± ++ ± + ± +++ ± ++ ± +++ ++

+++ +++ +++ +++ +++ +++ +++ ± +++ +++ +++ +++ +++ +++ +++ ±

25 ± ++ ++ ± ++ ± ± ± ± ± ++ ++ ± ± ± ± ± ++

26

27

+++ ± ± +++ +++ +++ ++ ± ++ ++ ++ ± + ± +++ ± ++ ± +++ ± ± ++ ± ++ ++ ± ++ ± + ± +++ ± ++ ± +++ +++

29 ++ ± ++ ++ ++ ++ ++ + + ++ ± ± ++ + + ++ ++ ++

30

32

+++ ± +++ ++ ++ +++ + +++ +++ +++ ± ± ++ + + +++ + +++

+++ ± +++ ++ ++ +++ + +++ ++ +++ ± ± ++ + + +++ + ++

+++ +++ +++ +++ +++ +++ +++ +++ +++ +++ +++ +++ +++ +++

a The results of 30 mg of algal extracts (rows) tested for their antimicrobial activities against strains of marine bacteria (columns). Activity was measured by the diameter of the inhibition zone. ±, no zone of inhibition; +, 1±3 mm zone of inhibition; ++, 3±5 mm zone of inhibition; +++, 5±6 mm zone of inhibition. Algal extracts B and C represent respectively the ethanol and the dichloromethane fraction (3-C. rupestris, 4-A. nodosum, 10-S. muticum, 11-E. siliculosus, 15-L. ochroleuca, 17-C. crispus, 19-G. latifolium, 20-P. palmata, 23-C. rubrum, 24-C. ramosa, 28-L. pinnati®da, 30-P. lanosa). The Gram-positive bacteria are strains number 1, 2, 12, 13, 20, 22, 23, 26, 29, 30 and 32 and the Gram-negative bacteria are the strains number 17, 18, 19, 24, 25 and 27. No activity were recorded against strains number 3, 4, 5, 6, 7, 8, 9, 10, 11, 14, 15, 16, 21, 28, 31, 33, 34 and 35.

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1B 3C 4B 4C 10B 10C 11B 11C 15B 17B 19C 20C 23C 24C 28B 28C 30B 30C

1

237

238

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Table 3 Determination of the minimum inhibitory concentration (MIC)a

1B 3C 4B 4C 10B 10C 11B 11C 15B 17B 19C 20C 23C 24C 28B 28C 30B 30C TBTO CuSO4

1

2

12

13

17

18

19

20

22

23

24

25

26

27

29

30

32

48 ± 48 48 48 48 48 96 48 48 ± ± 48 96 96 48 48 48

48 ± 48 48 48 48 48 96 48 48 ± ± 48 96 96 48 48 48

24 ± 24 24 48 48 96 24 96 24 ± ± 24 48 96 24 48 48

24 ± 24 48 48 24 96 24 24 24 ± ± 48 48 96 24 48 24

± 48 48 ± 48 ± ± ± ± ± 48 48 ± ± ± ± ± 48

± 24 24 ± 48 ± ± ± ± ± 48 24 ± ± ± ± ± 24

± 24 24 ± 48 ± ± ± ± ± 48 48 ± ± ± ± ± 24

48 ± 48 48 48 48 48 96 48 48 ± ± 48 96 96 48 48 48

48 ± 48 48 48 48 48 96 48 48 ± ± 96 96 96 48 48 48

24 ± 24 48 48 24 96 24 96 24 ± ± 24 48 96 24 48 24

± 24 24 ± 48 ± ± ± ± ± 48 24 ± ± ± ± ± 48

± 48 48 ± 48 ± ± ± ± ± 48 48 ± ± ± ± ± 48

24 ± 24 48 48 48 96 24 48 24 ± ± 48 48 96 24 48 24

± 24 24 ± 48 ± ± ± ± ± 48 48 ± ± ± ± ± 24

48 ± 48 48 48 48 48 96 96 48 ± ± 48 96 96 48 48 48

24 ± 24 48 48 24 96 24 24 24 ± ± 48 96 96 24 96 24

24 ± 24 48 48 24 96 24 48 24 ± ± 48 96 96 24 96 48

± ±

± ±

6 6

6 6

6 6

6 6

6 6

6 6

6 6

± ±

6 6

6 6

6 6

6 6

6 6

6 6

6 6

a The algal extracts were tested for their minimum inhibitory concentration of the sensitive strains of the marine bacteria. Minimum inhibitory concentrations are expressed in mg/ml. ±, CMI>96 mg/ml. Algal extracts B and C represent respectively the ethanol and the dichloromethane fraction (3-C. rupestris, 4-A. nodosum, 10-S. muticum, 11-E. siliculosus, 15-L. ochroleuca, 17-C. crispus, 19-G. latifolium, 20-P. palmata, 23-C. rubrum, 24-C. ramosa, 28-L. pinnati®da, 30-P. lanosa). The Gram-positive bacteria are strains number 1, 2, 12, 13, 20, 22, 23, 26, 29, 30 and 32 and the Gram-negative bacteria are the strains number 17, 18, 19, 24, 25 and 27.

3.2. Toxicity tests Four di€erent toxicity tests against non-target species were investigated for those extracts showing the highest levels of antibacterial activity: 1B, 3C, 4B, 4C, 10B, 10C, 11B, 11C, 15B, 17B, 19C, 20C, 23C, 24C, 28B, 28C, 30B and 30C. Results are shown in Figs. 1, 2, 3a, 3b, 4a and 4b respectively for the toxicity towards oyster larvae, sea urchins larvae and towards the mitotic and lysosomal activities of mouse ®broblasts. In a previous study, extracts 1B, 4B, 4C, 10B, 15B, 17B, 24C, 28B, 28C, 30B and 30C have been shown to be non-toxic towards larvae of sea urchin and oyster (Hellio et al., 2000b). TBTO and CuSO4 were found to be very toxic towards oyster and sea urchin larvae. No viable larvae were detected at 50 mg/ml. The same results were observed towards the mitotic and lysosomal activities of mouse ®broblasts. Among the 11 algal extracts tested for toxicity towards oyster and sea urchin larvae, two extracts of Ectocarpus siliculosus (11B, 11C) appeared to be toxic. They gave more than 50% of mortality of larval oysters when added at a concentration of

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Fig. 1. Percentage mortality of oyster larvae at di€erent concentrations (1±1000 mg/ml) of algal extracts.

125 mg/ml and of sea urchin larvae when added respectively at 500 mg/ml for 11B and at 125 mg/ml for 11C. The toxicity of ethanol and dichloromethane fractions of Ectocarpus siliculosus was found again on mouse ®broblasts cells. These two extracts (11B, 11C) gave a level of inhibition of the mitotic and lysosomal activities higher than 50% at 10 mg/ml. 50% inhibition of mitotic activity was observed at 50 mg/ml for the ethanolic fraction of Laminaria ochroleuca (15B). Fifteen extracts appeared non toxic to invertebrate larvae and on lysosomal and mitogenic activities even at concentrations up to 1000 mg/ml. These non toxic extracts are: Enteromorpha intestinalis (1B), Cladophora rupestris (3C), Ascophyllum nodosum (4A, 4B), Sargassum muticum (10B, 10C), Chondrus crispus (17B), Gelidium latifolium (19C), Palmaria palmata (20C), Ceramium rubrum (23C), Cryptopleura ramosa (24C), Laminaria pinnati®da (28B, 28C), and Polysiphonia lanosa (30B, 30C). 4. Discussion Marine bacteria are major organisms involved in the formation of the microlayer which is the ®rst step of the process of fouling (HolmstroÈm & Kjellberg, 1994). It is often assumed that the inhibition of the formation of the bio®lm will lead to the inhibition of the adhesion of the following fouling stages. Generally, fouling development can be prevented by means of antifouling paints based on copper oxide or tributyltin oxide containing one or more toxic compounds such as organotin

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Fig. 2. Percentage mortality of sea urchin larvae at di€erent concentrations (1±1000 mg/ml) of algal extracts.

derivatives in a paint matrix (Vallee-Rehel, Mariette, Hoarau, Guerin, Langlois, & Langlois, 1998). When submerged, antifouling paints release toxic compounds at levels (4 mg/cm2 of surface per day) that can cause adverse environmental e€ects (Gibbs, 1993; Gibbs, Bryan, Pascoe, & Burt, 1987; Gibbs, Pascoe, & Burt, 1988; Iwata, Tanabe, Mizuno, & Tatsukawa, 1995; Ohira, Matsui, & Nitta, 1996; Ponasik, Conova, Kinghorn, Kinney, Rittschof, & Ganem, 1998): tin-based paints have been linked to the death of dolphins, porpoises and whales in North Atlantic waters since the 1980s (Ponasik et al., 1998) and to production declines in the cultured oyster Crassosteras gigas in Arcachon Bay (France). Oysters are severely a€ected by TBT partly as a result of their ecient ®ltering ability which exposes them to high concentrations of xenobiotics, with consequence on human health (Peterson, Batley, & Scammell, 1993). In response to increasing scienti®c evidence on the toxicity and occurrence of organotin residues from antifouling paints in the aquatic environment, the application of triorganotin antifouling paints on boats of less than 25 m length has, since 1987, been banned in many countries (Voulvoulis, Scrimshaw, & Lester, 1999). Studies on the antifouling mechanisms utilised by marine sedentary organisms may therefore provide valuable information for fouling control in marine technology. The antifouling agents incorporated into the paints if derived from naturally occurring substances may be less environmentally damaging than the current toxins,

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Fig. 3. (a) Percentage of inhibition of mitotic activity of mouse ®broblasts at di€erent concentrations (1± 1000 mg/ml) of algal extracts (1B, 3C, 4B, 4C, 10B, 10C, 11B, 11C, 15B and 17B). (b) Percentage of inhibition of mitotic activity of mouse ®broblasts at di€erent concentrations (1±1000 mg/ml) of algal extracts (19C, 20C, 23C, 24C, 28B, 28C, 30B and 30C).

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Fig. 4. (a) Percentage of inhibition of lysosomal activity of mouse ®broblasts at di€erent concentrations (1±1000 mg/ml) of algal extracts (1B, 3C, 4B, 4C, 10B, 10C, 11B, 11C, 15B and 17B). (b) Percentage of inhibition of lysosmal activity of mouse ®broblast at di€erent concentrations (1±1000 mg/ml) of algal extracts (19C, 20C, 23C, 24C, 28B, 28C, 30B and 30C).

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in particular they may have less activity against non-target species (Hellio et al., 2000b). Macroalgae are already well documented as possessing antibacterial activities against terrestrial bacteria. Indeed, antibacterial activities were found in organic extracts of Fucus vesiculosus, Fucus endentalus, Fucus spiralis (Lustigman & Brown, 1991), Cytoseira balaerica, Zanardinia prototypus, Codium coralloides, Laurencia obtusa (Caccamese et al., 1980), Pelevetia caniculata (Glombitza & Klapperich, 1985), Laminaria saccharina, Desmarestia ligulata (Rosell & Srivastava, 1987), Sargassum wightii, Gracilaria corticata, (Sastry & Rao, 1994), Ascophyllum nodosum, Chondrus crispus, Enteromorpha intestinalis, Laminaria ochroleuca, Laurencia pinnati®da, Polysiphonia lanosa and Cryptopleura ramosa (Hellio et al., 2000b). In this study, the marine bacteria chosen are representative from the microfouling community and constitute the ®rst colonisers Ð and are target for antifouling compounds Ð found on immersed surfaces. These strains are involved in the process of colonisation of living (algae) or non-living (sediments, sand) surfaces in the tidal zone. Of the 90 algal fractions tested in this study against the 35 strains of marine bacteria, 12 extracts (1B, 3C, 4B, 4C, 10C, 15B, 17B, 20C, 23C, 28C and 30C) which showed in vitro antibacterial activity with a MIC as low as 24 mg/ml present no toxicity towards mollusc larval development or to the growth of mouse cells. Considering the variety of pathways that are possible for the chemical-induced cytotoxicity resulting in necrosis, a wide array of bioindicators are available and applicable (Tyson, 1987). In the present work, we used two di€erent end-points, that have been validated in a previous study (Fautrel et al., 1991), namely the MTT biotransformation and the Neutral Red incorporation, for mitochondrial and lysosomal functionality, respectively. Toxicity tests were also performed towards larvae of oyster and sea urchins which are the marine bioindicators most often studied. Three extracts showed toxic e€ects (11b, 11C and 15B) similar to TBTO and CuSO4 and are therefore not suitable for potential use in marine antifouling paints. The di€erence in toxicity between the mitotic and lysosomal tests showed that the different algal extracts do not have the same cellular targets. This shows the necessity of using several toxicity tests. The most active extracts are the organic fractions of one Fucaceae [Ascophyllum nodosum (4B)], one Gigartinaceae [Chondrus crispus (17B)] and two Ceramiaceae [Polysiphonia lanosa (30C)] and [Laurencia pinnati®da (28C)]. Considering the importance of marine bacteria in the early stages of marine fouling, Ina, Takasawa, Yagi, Yamashita, Etho, & Sakata (1989) studied the correlation between antibacterial and antifouling activities and showed that with few exceptions, antibacterial activity against Gram-positive bacteria seems to be prerequisite for antifouling compounds. In our study, among the 18 active non toxic extracts, three extracts (3C, 19C and 20C) showed speci®c inhibition against only Gram-negative bacteria (17, 18, 19, 24, 25, 27), 12 extracts (1B, 4C, 10C, 17B, 23C, 24C, 28B, 28C and 30B) showed speci®c inhibitory activities against only Grampositive bacteria (1, 2, 12, 13, 20, 22, 23, 26, 29, 30, 32). In a previous study (Hellio et al., 2000b) these extracts were tested for their activities against marine fungi and extracts 4C, 17B, 28B and 28C and shown to give a high level of inhibition towards these microorganisms. Three extracts (4B, 10B and 30C) inhibited the growth of both Gram-positive and Gram-negative marine bacteria, and they have been previously

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described as inhibitors of marine fungi (Hellio et al., 2000b). These seven extracts gave very promising results for a potential antifouling application. Three of the bacterial strains studied (1, 2 and 23) showed some tolerance to the antifouling products tested. A previous study indicated that the bacterial populations responded very strongly to a small addition of copper (Jonas, Gilmour, Stoner, Weir, & Tuttle, 1984; Tubbing & Admiraal, 1991; Vives Rego, Vaque, & Martinez, 1986). Heavy metal contamination is a persistent threat to aquatic communities and despite the sanitation of some industrial sources, several metals are now ubiquitous in soils and sediments (Tubbing, Santhagens, Admiraal, & Van Beleen, 1993). Typically, natural microbial populations respond to the input of heavy metals by developing resistance (Barkay, 1987; Duxbury & Bicknell, 1983). Vaccaro, Azam, and Hodson (1977) provided evidence that the response of heterotrophic bacteria to copper was e€ected by a simultaneous selection for bacterial species exhibiting an increased tolerance towards copper. The increasing number of observations of bacterial aquatic community resistances to heavy metals lead to the conclusion that it is a matter of urgency to ®nd new antifouling products. These results con®rm that marine algae are a rich source of antibacterial compounds and this feature may be important from the commercial perspective of developing novel antifouling coatings. Previous results indicate that extracts B and C of Enteromorpha intestinalis; extracts B and C of Sargassum muticum and extract B of Polysiphonia lanosa marked also speci®c inhibition of the activity of the phenoloxidase (enzyme involved in the mussel attachment by byssus formation) puri®ed from the foot of mussel Mytilus edulis (Hellio et al., 2000a). This con®rms that algal extracts can act at di€erent stages of the fouling process including micro and macrofouling. There is also a potential for incorporating such compounds in broad spectrum antifouling coatings containing a mixture of compounds, each of which will be most e€ective against a particular group of fouling organisms. Among the marine algal extracts tested, some appeared to be speci®c in their activity against several test bacteria, this point may be important for the development of novel antifouling coatings. Further work is needed to identify the compounds causing the activity, to evaluate speci®c antimicrobial activity against marine bacteria speci®cally implicated in biofouling and to examine the precise role of such activity in nature. Acknowledgements The work was carried out during tenure by Brittany Council and European studentship. We wish to thank both organisations. We thank Dr. Jean Pascal Berge (Ifremer, Nantes, France) for his helpful comments and criticisms of the manuscript. References Abarzua, S., & Jakubowski, S. (1995). Biotechnological investigation for the prevention of biofouling. I. Biological and biochemical principles for the prevention of biofouling. Journal of Marine Ecology and Progress Series, 123, 301±312.

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