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Chemical defense against fouling in the solitary ascidian Phallusia nigra.

Introduction

In marine benthic environments, any exposed, undefended. long-lived surface becomes fouled (Wahl, 1989). The fouling process consists of multiple steps starting with the development of an organic film by adhesion of macromolecules, followed by the settlement of microorganisms and the development of a microbial biofilm. The settlement of unicellular and multicellular eukaryotes establishes the mature fouling community composed of prokaryotes, fungi, diatoms, algae, protists, and invertebrates (Krug, 2006). Fouling also occurs on the surfaces of living organisms, and sessile marine invertebrates are particularly susceptible to fouling (Wahl, 1989). Although fouling organisms can be beneficial to their hosts, for example by offering camouflage and by providing protection from predators by their secondary metabolites (Laudien and Wahl, 1999), the interaction is largely considered harmful. Among the negative effects are increased hydrodynamic drag on the host, which can lead to dislodgment of sessile hosts; and disruption of the host's feeding, either by direct competition for food particles or by changing the water flow to the host, which is especially important for filter-feeding organisms (Stoecker, 1978; Wahl, 1989; Krug, 2006).

Ascidians (phylum: Chordata, subphylum: Tunicata) are soft-bodied, sessile, filter-feeding organisms. They occur in all oceans in colonial and solitary forms. Fouling on ascidians can lead to the obstruction of the inhalant and exhalant siphons as well as to the clogging of the branchial basket, important to both respiration and feeding (Lambert, 1968; Koplovitz et al., 2011). Both naturally fouled and non-fouled ascidian species have been reported from various geographical regions (e.g., Stoecker, 1980; Uriz et al., 1991; Davis and White, 1994). Fouled ascidian species might use behavioral responses (e.g., actively closing the siphons or squirting water and particles from the branchial basket through apertures [Hoyle, 1953]) and physical responses (e.g., regular tissue sloughing [Goodbody, 1962] and mucus secretion [Wahl et al., 1998]) to overcome fouling. Short-lived ascidians might be able to grow and reproduce before becoming fouled (Stoecker, 1980). Among the taxa reported as epibionts on ascidians are bacteria, algae, bryozoans, hydroids, soft corals, and sponges (Stoecker, 1980; Davis and White, 1994; Wahl et al., 1994).

Chemical defense against fouling has been found in several ascidians and against various fouling organisms. Ascidian crude extracts have been shown to inhibit fouling by barnacles and bryozoans (Davis and Wright, 1989; Bryan et al., 2003) as well as the growth and settlement of environmental bacteria (Wahl et al., 1994; Bryan et al., 2003) and diatoms (McClintock et al., 2004; Koplovitz et al., 2011). Furthermore, antifouling and antibacterial compounds have been isolated from ascidians (Davis and Wright, 1990; Tsukamoto et al., 1994; Davis and Bremner, 1999). Tunic surface-associated bacteria from ascidians have been shown to produce antibacterial and antifouling compounds in laboratory cultures (Olguin-Uribe et al., 1997; Holmstrom et al., 1998, 2002), and symbiotic bacteria have been assumed to be the real producers of some known bioactive compounds originally isolated from their ascidian hosts (Simmons et al., 2008).

In the present study, we investigated the antifouling chemical defense potential of the solitary tunicate Phallusia nigra (= Ascidia nigra) (Savigny, 1816). P. nigra is a cosmopolitan species occurring in the West Atlantic from Florida throughout the Caribbean (Bermuda, Jamaica) to Brazil, throughout the Mediterranean Sea, and in the Red Sea from Yemen to the northern tip of the Gulf of Aqaba. Along the Israeli Mediterranean Sea, it is commonly found at depths between 1 and 30 m as part of the fouling community on hard substrates such as rocks or man-made structures (Shenkar and Loya, 2009). Even though P. nigra lives up to 2 years (Goodbody, 1962) and in environments with high fouling pressure (Goodbody, 1965), it has frequently been described as free of epibionts (Goodbody, 1962; Stoecker, 1978, 1980; authors' own observations (Fig. 1A); see also figure 2b in Shenkar and Loya, 2009). Early studies suggested that P. nigra regularly sloughs off its external layers and thus cleans itself from epibionts (Flecht, 1918; Goodbody, 1962). High vanadium concentrations and low tunic surface pH (<2) were also suggested as defenses against fouling organisms (Stoecker, 1978). These assumptions were challenged by later studies as ascidian species containing high vanadium concentrations, comparable to those in P. nigra, were found to be fouled (Stoecker, 1980). Tunic surface acidity as an antifouling mechanism in ascidians was also later questioned due to the presence of heavily fouled ascidians with low tunic surface pH (Parry, 1984; Davis and Wright, 1989).

The role of secondary metabolites in the defense of P. nigra, especially against fouling, is poorly understood. Results from predator feeding-deterrence assays with chemical extracts indicated that while whole-tissue extracts did not deter fish feeding, active (i.e., deterrent) compounds were present in the extracts from gonads and the combined tissue of the branchial basket and gut (Pisut and Pawlik, 2002). Furthermore, crude extracts of P. nigra were active against various standard laboratory bacteria (Jaffarali et al., 2008; Amutha et al., 2010), but not against four known pathogens of marine invertebrates (Odate and Pawlik, 2006).

Given the unclear role of secondary metabolites in the fouling defense of P. nigra, its clean surface, and the good track record of ascidians as a source of antifouling compounds, we investigated the antifouling activity of the secondary metabolites of P. nigra in laboratory and field assays. For comparison, we assayed organic extracts from the solitary ascidian Herdmania momus (Savigny, 1816) in the laboratory assays. This latter species is usually fouled in nature (Davis and White, 1994; Shenkar and Loya, 2009; authors' own observation (Fig. 1D)) and is sympatric with P. nigra along the Israeli Mediterranean and Red Sea coasts. Since every step in the fouling process affects the subsequent ones (Krug, 2006; Wahl, 2008), we first tested the antibacterial activity of extracts against a panel of sympatric and laboratory bacteria, and used scanning electron microscopy to determine natural bacterial fouling on the tunic surface of P. nigra and H. momus. The ability of extracts to inhibit metamorphosis of a representative macro-fouling invertebrate was examined against larvae of the sympatric bryozoan Bugula neritina (Linnaeus, 1758). General toxicity of crude extracts was tested against larvae of the brine shrimp Artemia salina (Linnaeus, 1758). Finally, extracts of P. nigra from the Red Sea and the Mediterranean Sea were exposed to natural fouling communities in field assays in their respective locations. In order to decouple the antifouling effect of low pH from that of secondary metabolites, the pH of the extracts was monitored in all assays, and control experiments were performed for all laboratory assays.

Materials and Methods

Sample collection and extraction

The ascidians Phallusia nigra and Herdmania momus were collected by scuba from various sites along the Israeli Mediterranean coast at depths from 5 to 30 m. P. nigra samples used in Red Sea experiments were collected in Eilat, northern Red Sea, Israel. After each dive, the volume of the collected ascidians was determined by seawater displacement. Samples were then stored at -20[degrees]C until extraction took place.

Frozen ascidians of known (wet) volume were cut into 1-[cm.sup.3] cubes and covered with a mixture of 1:1 dichloromethane and methanol in a flask. The flask was kept at 4[degrees]C for 24 h, during which it was periodically shaken to ensure that the ascidian tissue was equally exposed and extracted by both solvents. The resulting phases methanol:water and dichloromethane were separated, and both phases were filtered (Whatman, 8-[micro]m pore size). Each phase was evaporated to dryness using a rotary evaporator (Buchi Rotavapor R-114) with a water bath at a temperature below 40[degrees]C. Extracts of both phases were combined into the same glass bulb. The extraction was repeated a second time, followed by a final extraction with methanol at 4 [degrees]C for 6 h. All phases were combined in the same glass vial, resulting in the crude extract, which was weighed and stored at -20 [degrees]C until use in the assays. Natural concentrations were calculated by dividing the crude extract weight by the wet ascidian volume.

Scanning electron microscopy

Collected ascidians were rinsed with 0.22-[micro]m-filtercd seawater, fixed in 2.5% glutaraldehyde in seawater, and kept at 4[degrees]C until further preparation. From three individuals of each species, three 1 X 1-cm samples were taken and washed with PBS, dehydrated in an EtOH gradient followed by critical-point drying with liquid C[O.sub.2], and coated with gold-palladium. The tunic surfaces of the samples were scanned for bacteria at magnifications ranging from 100X to 7000 X using a Jeol SEM JSM 840A.

Antibacterial assay

Crude organic extracts from P. nigra and H. momus were tested for antibacterial activity against eight environmental bacteria and four standard laboratory bacteria (Table 1). Isolation and identification of the environmental bacteria from the Israeli Mediterranean coast are described in Haber et al. (2011). The test protocol was adapted from Kelman et al. (2001). Test bacteria were cultured overnight in liquid medium (Marine Broth 2216 and LB for environmental and laboratory bacteria, respectively). Culture densities were measured at 620 nm using a Turner SP-830 spectrophotometer. The cultures were adjusted to an optical density of 0.04 and diluted 1:100. From the diluted cultures, 100 [micro]l was added to a 96-well plate containing 100 [micro]l of appropriate liquid medium, DMSO (final concentration 0.5% v/v), and a series of extract concentrations (ranging from 0.625 to 20 mg [ml.sup.-1], which represents 3% to 100% and 6% to 200% of natural volumetric concentration of P. nigra and H. momus, respectively, and including a solvent control with 0 mg mf'). Each extract concentration and control was assayed in triplicate. Initial optical density was measured at 620 nm on a GENESIS Workstation 200 (Tecan) using SPECTRA-Fluor Plus (Tecan), after 5 s of shaking and 2 s of settling. Plates were incubated at 30[degrees]C with shaking (80 rpm). Optical density measurements were repeated after 24 and 48 h. Minimal inhibitory concentrations were determined as the smallest amount of extract needed to inhibit growth by more than 85% compared to growth in the controls without extract.

Bugula neritina larvae assay

Colonies of B. neritina were collected from marinas in Tel Aviv and Herzelia, Israel (Mediterranean Sea). After transfer to the laboratory, colonies were maintained in an aquarium. To promote larval release, colonies were kept in the dark for 24 h and then exposed to light (Dahms et al., 2007). The released larvae were collected during the next 30 min by using a glass pipette and were transferred into 0.22-[micro]m-filtered seawater. Larvae were then immediately pipetted into the assay wells. The experiment was conducted in 24-well tissue-culture plates (Corning). Extracts were dissolved in methanol and diluted to the required concentrations (see below). An aliquot of 2.7 ml of dissolved extract was pipetted into each well and evaporated to dryness. Methanol only was used for control wells. Three to four larvae were pipetted into each well with 2.7 ml of seawater. Larval vitality state was recorded as alive-swimming, alive-metamorphosed, and dead after 0, 24, and 48 h. The effect of P. nigra and H. momus extracts at concentrations between 0.05 and 2 mg [ml.sup.-1] was determined by comparison with the larval development in the control wells. At least 12 wells were analyzed for every tested concentration and controls, and the percentage of metamorphosed larvae was calculated for each well. As obtained data were not normally distributed, significance of observed differences between wells containing P. nigra and H. momus extracts at the various concentrations and controls was determined using a Kruskal-Wallis test, followed by multiple comparison tests once significant differences were found. All statistical analyses were done using Statistica 9 software.

Brine shrimp assay

Toxicity of P. nigra and H. momus crude extracts was tested against nauplii of the brine shrimp Artemia salina. Brine shrimp were hatched from eggs in an aquarium and collected using a glass pipette into 0.22-[micro]m-filtered artificial seawater (Instant Ocean). Extracts were dissolved in DMSO and diluted to the required concentrations (final DMSO concentration in the assay 1% v/v). Ten to fifteen brine shrimp larvae were transferred to a 2-ml Eppendorf tube with a minimum amount of artificial seawater. The volume was adjusted to 1.2 ml with artificial seawater, and 0.3 ml of extract was added. A DMSO control without extract was prepared accordingly. Each extract concentration and the control were tested in triplicate. The number and vitality state--alive (any movement of the larvae) or dead)--of the brine shrimp larvae in each tube were recorded after addition of extract or DMSO control at 0, 24, and 48 h. The extracts of both ascidian species were tested at natural whole-tissue volumetric concentration and 2 mg [ml.sup.-1] (= 10% and 20% of natural whole-tissue volumetric concentration for P. nigra and H. momus, respectively). In addition, a range of concentrations (0.01-5 mg [ml.sup.-1]) was tested for the P. nigra crude extract to determine the LC50, which was calculated by trimmed Spearman-Kaber estimation using the TSK program ver. 1.5 as provided by the ecological exposure research division of the U.S. Environmental Protection Agency (www.epa.gov/eerd/stats2.htm as accessed 7 May, 2012).

Field experiments

Agar gel surfaces containing the crude extract of P. nigra were used as substrate for the settlement of fouling organisms in natural settings. The experimental protocol was adapted from reports using similar methods (Slattery et al., 1995; Henrikson and Pawlik, 1995). Agar solution (5% w/v Difco, granulated agar) was autoclaved and kept at 50[degrees]C until extract addition. The dry extract obtained from 50 ml of ascidian tissue was dissolved in 3 ml of methanol and mixed into 50 ml of agar solution, resulting in natural volumetric extract concentration in the gel. Control gels contained only 3 ml of methanol. To prepare the settlement assay, 15 ml of agar solution was poured into each well of a six-well tissue-culture plate. In each plate three wells contained agar with extract, and three wells were filled only with control agar. The order of the control and treatment wells was randomized. The plates were secured to plastic meshes, which were attached with cable ties to piers at a depth of 0.5-1 m, next to naturally fouled surfaces at the Interuniversity Institute in Eilat (Red Sea) and Herzelia Marina (Mediterranean Sea). In each experiment the plates were deployed and monitored weekly for the occurrence of damages and disturbances to the plates for a period of 4 weeks to allow for settlement and development of fouling organisms. After retrieval, plates were examined for macrofouling under a stereoscope, and the number of epibionts in each well was recorded. Fouling organisms were divided into three main categories: macroalgae, barnacles, and sessile polychaetes (Serpulidae). To test for seasonal variation of fouling organisms, this experiment was conducted eight times throughout the year in the Mediterranean and twice in the Red Sea (dates of the experiments are given in the figures).

The three groups of settled epibionts (macroalgae, barnacles, and polychaetes) were analyzed separately. Since the number of settled epibionts in each well was too low for statistical analysis, the data from all wells of the same treatment in each plate were combined. In each plate the distribution of settled epibionts on wells containing extract was compared to their distribution in control wells by a replicated goodness of fit test (Sokal and Rolf, 1995). For each group of epibionts, only experiments in which settlement occurred were included in the analysis.

During the deployment of the agar settlement surfaces, the extract slowly diffused out of the agar. To determine the concentration of extract in the agar from the initial deployment until the end of the experiment, six tissue-culture multi-well plates with agar gel containing extract and controls were prepared and deployed. The first plate was tested before deployment and the others after 1, 7, 14, 21, and 28 days. From each plate the three agar gels containing the extract were removed from the wells, combined, and then re-extracted. We used the same protocol as in the initial extraction, except that a butanol/water separation phase was added to the original extraction protocol to remove salts from the extracts. The same procedure was applied to the matching control wells. Once salts were removed, the weight of the extract from control wells was subtracted from the weight of extract in the matching treatment wells. The resulting "net" extract weights indicated the change in the amount of P. nigra extract due to leakage from the agar gels throughout the experiment. This experiment was performed twice, once in May and once in October during the research duration. Thin-layer chromatography (TLC) was used to verify that the same chemical compounds were found in the extract throughout the settlement experiments. Silica plates (Kieselgel 60 [F.sub.254], Merck) were loaded with samples from treatment and control wells in chronological order of their deployment. A toluene/dichloromethane 1:1 mixture was used as the mobile phase. TLC profiles were compared after visualization under UV light (254 nm) and after developing the TLC plate with vanillin stain (6 g vanillin in 1.5 ml concentrated sulfuric acid and 95 ml 96% ethanol).

Measurement of pH and pH control experiments

The pH in the assay wells was recorded at the end of all laboratory assays using pH indicator paper (Merck 0-14 and Merck 5.0-10.0). In the field assay, the pH of the agar mixtures was measured during plate preparation, and the surface pH of the plates was measured again after 24 h of deployment. For laboratory assays, control experiments were performed with media adjusted to pH 5.5-8.5 (antibacterial assay) and seawater adjusted to pH 6.0-8.0 (B. neritina larvae assay, brine shrimp assay) in 0.5-unit increments according to the range of pH measured at the end of the assays. These measurements and experiments were performed to ensure the obtained results were not influenced by extract acidity.

Results

Natural bacterial fouling on ascidian tunics and antibacterial assay

Tunic samples of Herdmania momus were heavily overgrown by epibionts. As this did not allow a quantitative comparison between both tunicate species, only a qualitative screen of the samples was performed. Scanning electron microscopy examination showed that the tunic of Phallusia nigra was largely bacteria-free, with the exception of a few clusters of coccoid-shaped bacteria. In contrast, various bacteria were clearly evident on the tunics of the sympatric H. momus (Fig. 1).

Chemical extract of the two ascidian species showed very different activity profiles in the antibacterial assay (Table 1). The P. nigra extract inhibited the growth of all environmental test bacteria at 5-10 mg ml-1 (25%-50% of natural whole-tissue volumetric concentration) and the two gram-positive laboratory test bacteria at 20 mg [ml.sup.-1]. The H. momus extract inhibited only the gram-negative environmental alpha-proteobacterium Loktanella sp. ESY23 at 10 mg [ml.sup.-1], a concentration slightly above natural whole-tissue volumetric concentration.

The pH in tests with laboratory bacteria conducted in LB medium was 8.5 for all extract concentrations of both ascidian species and the controls. Growth of laboratory bacteria was not inhibited in LB medium adjusted to pH 5.5-8.5. In tests with environmental bacteria cultured in Marine Broth 2216, the pH was 8.5 in the controls and all H. momus concentrations. The pH of extracts of P. nigra in Marine Broth 2216 at 20, 10, and 5 mg [ml.sup.-1] was 5.5, 6.5, and 7.5, respectively. Marine Broth 2216 adjusted to pH 5.5 to 8.5 did not inhibit the growth of half of the environmental bacteria. The growth of the remaining four bacteria strains ESY10, ESY12, ESY17, and ESY23 was only inhibited at pH 5.5, but not at pH 6.0 to 8.5. The pH at the minimal inhibitory concentrations for these bacteria was 6.5 (ES Y12, ESY17) and 7.5 (ESY10, ESY23), respectively.

Bugula neritina assay

Significant differences in the metamorphosis of B. neritina larvae between treatments were found at all concentrations tested, which were well below natural whole-tissue concentrations (Kruskal-Wallis test [H.sub.2] > 20, n = 48, P < 0.005 for all concentrations). These differences were due to the lower number of metamorphosed larvae in wells with the ascidian extracts compared to the controls (Fig. 2). The average number of metamorphosed larvae was always lower in tests with P. nigra crude extracts than in the corresponding tests with H. momus crude extracts. However, this difference was only significant at concentrations of 0.1 and 0.25 mg [ml.sup.-1] (Kruskal-Wallis test, [H.sub.2] = 25.718, P = 0.025 and Kruskal-Wallis test, [H.sub.2] = 27.561, P = 0.030, respectively). At the lowest tested concentration of 0.05 mg [ml.sup.-1] more than 50% of the B. neritina larvae metamorphosed in wells with both extracts (52% and 62% in wells with P. nigra and H. momus extracts, respectively).

Throughout the experiment pH 7 was recorded in all wells containing P. nigra extracts, pH 7-7.5 in wells containing H. momus extract, and pH 8 in control wells. Control experiments performed in seawater adjusted to pH 6.0 to 8.0 showed larval metamorphosis and settlement was significantly inhibited only at pH 6, but not at pH 6.5-8.0 (Kruskal-Wallis test, [H.sub.4] = 24.78, P = 0.0001, multiple comparisons test P < 0.03 to <0.001).

Brine shrimp assay

H. momus and P. nigra crude extracts killed all brine shrimp larvae at their respective natural whole-tissue volumetric concentration of 9 and 20 mg [ml.sup.-1]. At 2 mg [ml.sup.-1], however, only the P. nigra crude extract was active and killed on average 82.2% ([+ or -] 10.2% STD), whereas mortality in the H. momus crude extract was almost the same as in the control (Table 2). Based on the tested range of concentrations (Table 2) the L[C.sub.50] of the P. nigra crude extract was found to be 1.11 mg [ml.sup.-1] with a 95% confidence of 0.94-1.32 mg [ml.sup.-1] (trimmed Spearman-Kaber estimation). The seawater pH after addition of the P. nigra crude extracts was between 6 and 7. Survival of brine shrimp was not affected in seawater adjusted to pH 6.0-8.0.

Field assay results

P. nigra crude extract was tested at natural concentration in the field against various epibionts present in the ascidians' natural environment. During the 28 days of the Mediterranean field experiment, as much as 52% of the original extract weight leaked out from the gel surfaces (Fig. 3). Qualitative TLC analysis of the P. nigra extract content of these agar gels during the settlement trials showed that the same chemical pattern was present before the gels were immersed in seawater and throughout the the experiment. The TLC analysis of the control agar gels showed no comparable spots. Therefore all active secondary metabolites originally found in P. nigra extract were still present in the settlement gels at the end of the field experiments. During gel preparation, the pH for agar gels with H. momus extract was recorded as 6.1, agar with P. nigra extract was pH 4, and the pH of agar control gels was 5.5. However, the pH for all gels after a 24-h exposure to seawater was recorded as 7.5.

In the Mediterranean settlement trials, barnacle settlement was most abundant from June to October, while very little settlement occurred from October till March (Fig. 4A). Barnacle settlement on P. nigra extract wells was significantly lower than on control gels throughout the experiment (replicated goodness of fit test, one-tailed, P < 0.001, df = 20). Settlement of polychaetes was most abundant between May and November, and none occurred from December till April (Fig. 4B). Statistical analysis of the results showed a significant difference in polychaete settlement between wells containing P. nigra extract and control wells (replicated goodness of fit test, one-tailed, P < 0.005, df = 19). Analysis of all statistic parameters of this test showed that this was due to a much lower polychaete settlement on gels containing the P. nigra extract, in all but one of the experiments. Algal settlement was noted from October to May (Fig. 4C). A significant difference was found between the amount of algae on P. nigra extract wells and on control wells (replicated goodness of fit test, one-tailed, P < 0.001, df = 8) due to a much lower algal settlement on gels containing the P. nigra extract, throughout all the experiments.

Barnacle settlement was observed in both Red Sea trials (Fig. 4D). Barnacles settled significantly less on gels with P. nigra extract than on control gels, throughout all the experiments (replicated goodness of fit test, one-tailed, P < 0.001, df = 6).

Discussion

Previous studies on the ascidian Phallusia nigra suggested that it keeps its surface free of fouling organisms by tissue sloughing (Hecht, 1918; Goodbody, 1962), high vanadium concentrations (Stoecker, 1978), and the acidic tunic surface (Stoecker, 1978; 1980; Odate and Pawlik, 2007). After confirming that tunics of Mediterranean P. nigra are largely free of fouling organisms, we examined the role of secondary metabolites in antifouling defense of P. nigra. Our results demonstrated that P. nigra crude extracts inhibited the growth of environmental bacteria, inhibited the metamorphosis of the bryozoan Bugula neritina, and reduced settlement by polychaetes, algae, and barnacles in field experiments. These activities were not due to low pH of the extracts, as shown by pH monitoring and control experiments. Therefore, secondary metabolites defend P. nigra against the settlement and growth of fouling organisms on its surface. A similar role for secondary metabolites has previously been shown in sponges, corals, bryozoans, and other ascidian species (Bryan et al., 2003; Haber et al., 2011; Fusetani 2012).

The idea that acids act as an antifouling mechanism in ascidians has already been challenged by Parry (1984) for two reasons: (i) the rapid dilution of acids by seawater and (ii) the presence of heavily fouled ascidian species with low tunic acidity (pH < 2), which was also later reported by Davis and Wright (1989). The tunic surface pH of P. nigra is normally around 4.5 and decreases upon mechanical stimuli to pH 1-2 due to the release of acids, mainly sulfuric acid, from bladder cells at the point of injury (Hirose et al., 2001). In aquarium experiments, the low pH remains stable for 5 min if the surrounding seawater is stirred, and then gradually climbs again over the next 6 min (Hirose et al., 2001). This mechanism might be involved in the defense against predation and infection (Hirose et al., 2001; Odate and Pawlik, 2007; but see Lopez-Legentil et al., 2006, as an example of the inefficiency of acidity in deterring tunicate predators), but due to the rapid dilution, acidity alone seems to be ineffective in preventing fouling and overgrowth by marine invertebrates (Davis and Wright, 1989). Our data suggest that P. nigra possesses antifouling secondary metabolites as the data from the pH monitoring during our experiments and our control experiments indicated that the observed antifouling activity of P. nigra extracts is not due to acidity. In the field assays, agar surfaces reached seawater pH after deployment for a single day, whereas the antifouling effect was still visible after 4 weeks of deployment. In the laboratory assays, the crude extracts of P. nigra showed activity at concentrations whose corresponding pH did not affect the assay organisms. This does not rule out the possibility of additive effects of acidity and secondary metabolites in P. nigra, but indicates that secondary metabolites active against fouling are present in this ascidian species.

Since its suggestion, the role of high vanadium concentrations as antifouling mechanism has been challenged. The concentration of vanadium in P. nigra is highest in the blood and internal organs but not in the ascidian tunic, which is the part exposed to potential epibionts (Odate and Pawlik, 2007) and where it should be stored if its main functional role is antifouling defense. Furthermore, other ascidians with comparably high vanadium concentrations are fouled (Stoecker, 1980; Davis and Wright, 1989). Odate and Pawlik (2007) already showed that non-acidic vanadium compounds do not deter predation nor inhibit microbial growth. Here, the P. nigra extracts were active in both field and laboratory assays at or near neutral pH. In the laboratory assays, vanadium was diluted compared to its natural volumetric concentration (e.g., up to 400 times in the B. neritina assay), while activity was still observed and pH was non-acidic. Therefore our results suggest that secondary metabolites present in the crude extracts are involved in the defense of P. nigra against fouling by bacteria, bryozoans, polychaetes, and barnacles.

Testing ecologically relevant concentrations is a major challenge in chemical ecology. We used whole-tissue volumetric concentrations, which probably underestimate the concentrations experienced by fouling organisms because compounds might be accumulated at the tunic surface. Yet even in the antibacterial assay, where activity was found at relatively high concentrations, it nonetheless was below natural (whole-tissue) volumetric concentration. The extracts examined here contained a large amount of salts, probably from seawater in the ascidians' tissues, which increased the extract weight. The salts are not responsible for the observed activities in the antibacterial assay since the sympatric H. momus did not produce positive results at the same concentrations as P. nigra. Although we tested the defense against bacterial fouling in a growth inhibition assay, bacterial fouling can also be prevented by compounds that are non- or weakly growth-inhibiting, as shown by prevention of bacterial attachment to the surface in other ascidians (Wahl et al., 1994), as well as for several sponge extracts and compounds (Kelly et al., 2003). A similar mechanism might also exist in the ascidian P. nigra given its bacteria-free tunic surface (Fig. 1), and could work together with physical defenses such as regular shedding of tunic tissue (Hecht. 1918; Goodbody, 1962).

The effect of P. nigra extract on Bugula neritina settlement may be a result of its direct effect on the larvae or a result of settlement cues from bacteria affected by the extract. Although the larvae were rinsed in filtered seawater and the assays were performed in filter-sterilized seawater, bacteria introduced with the larvae might have influenced settlement. However, B. neritina larvae are also known to settle rapidly and effectively on polystyrene plates without a biofilm (Maki et al., 1989; Dahms et al, 2004). Therefore, while we cannot rule out the influence of the antibacterial activity of the P. nigra extracts, we view these results as a direct inhibition of B. neritina settlement. The inhibitory effect of H. momus extracts on B. neritina larval settlement was somewhat surprising since this ascidian is normally covered by fouling organisms. In surveys conducted at southeastern Australian rock reefs, this ascidian was always fouled, although bryozoans were not among the reported fouling organisms (Davis and White, 1994). The observed activity was not due to a general toxicity of H. momus extract, as nauplii of the brine shrimp Artemia salina were not killed at 2 mg [m.sup.-1], the highest tested concentration in the B. neritina assay. A possible explanation is that H. momus extracts might rather be selectively active against various epibionts.

The P. nigra extract activities in the laboratory assays with larvae of the bryozoan B. neritina and the brine shrimp A. salina were found at relatively low concentrations, given the high salt portion in the extracts, indicating the potential for highly effective compound(s). The active compound (or compounds) could be an interesting candidate for potential commercial exploitation and deserve further investigation. The results of the field assays indicate that these compounds are likely lipophilic. Hydrophilic substances would have rapidly leaked out of the gel, together with acids and salts, as indicated by the rapid initial drop in extract weight in gels re-extracted after 24 h of deployment (Fig. 3). The presence of antifouling activity in the gel even after 4 weeks of deployment in the field points therefore to lipophilic active compounds. This suggestion is in line with theoretical arguments that antifouling compounds should be lipophilic in order to remain on an organism's surface and avoid rapid loss to the environment (Steinberg and de Nys, 2002).

In conclusion, the results reported here from field experiments, laboratory assays, and pH monitoring and control experiments show that secondary metabolites have a key role in keeping P. nigra surface free of epibionts. As in other ascidians (Wahl and Banaigs, 1991), this chemical antifouling defense may act alongside physiological and physical defenses suggested long ago (Hecht, 1918; Goodbody, 1962), but whose efficiency in P. nigra still need to be determined. The different activities observed in the extracts might also work additively--for example, the inhibition of epibiotic bacteria can lead to the absence of settlement cues necessary for macrofouling organisms. These fouling organisms would be affected directly by inhibitory secondary metabolites and indirectly by the absence of settlement cues. To evaluate this suggestion, further studies are needed and these would benefit from the isolation of the active secondary metabolites, which could be of biotechnological interest.

Acknowledgments

We thank Olga Bunis for valuable help with lab experiments.

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BOAZ MAYZEL *, MARKUS HABER, AND MICHA ILAN

Department of Zoology, George S. Wise Faculty of Life Sciences, Tel Aviv University, Tel Aviv 6997801, Israel

Received 25 September 2013; accepted 5 September 2014.

* To whom correspondence should be addressed. E-mail: bmayzel@ gmail.com

Table 1
Minimal inhibitory concentrations (in mg [ml.sup.-1) of
tunicate chemical extracts in antibacterial assay after 24 h

                                                             GenBank
Phylum; (Class)               Test bacterium                accn. no.

Gram positive
  Firmicutes                  Bacillus subtilis
  Firmicutes                  Staphylococcus aureus
  Actinobacteria              Kocuria sp. ESY10             GU479627
  Firmicutes                  Lysinibacillus sp. ESY09      GU059941
  Firmicutes                  Sporosarcina sp. NB90         GU479626
Gram negative
  Proteobacteria; ([gamma]-   Pseudomonas aerigunosa PA01
    Proteobacteria)
  Proteobacteria; ([gamma]-   Escherichia coli GM1655
    Proteobacteria)
  Proteobacteria; ([gamma]-   Pseudomonas sp. NB86          GU479630
    Proteobacteria)
  Proteobacteria; ([alpha]-   Sulfitobacter sp. ESY17       GU479629
    Proteobacteria
    Proteobacteria)
  Proteobacteria; ([gamma]-   Vibrio sp. ESY12              GU479628
    Proteobacteria)
  Proteobacteria; ([alpha]-   Erythrobacter sp. ESY19
    Proteobacteria
    Proteobacteria)
  Proteobacteria; ([alpha]-   Loktanella sp. ESY23
    Proteobacteria
    Proteobacteria)

Phylum; (Class)               Origin   P. nigra   H. momus

Gram positive
  Firmicutes                   Lab        20         --
  Firmicutes                   Lab        20         --
  Actinobacteria               Sed        5          --
  Firmicutes                   Sed        10         --
  Firmicutes                   Sed        5          --
Gram negative
  Proteobacteria; ([gamma]-    Lab        --         --
    Proteobacteria)
  Proteobacteria; ([gamma]-    Lab        --         --
    Proteobacteria)
  Proteobacteria; ([gamma]-    Seaw       10         --
    Proteobacteria)
  Proteobacteria; ([alpha]-    Seaw       10         --
    Proteobacteria
    Proteobacteria)
  Proteobacteria; ([gamma]-   Sed.w       10         --
    Proteobacteria)
  Proteobacteria; ([alpha]-    Seaw       5          --
    Proteobacteria
    Proteobacteria)
  Proteobacteria; ([alpha]-    Seaw       5          10
    Proteobacteria
    Proteobacteria)

The natural volumetric tissue concentrations of Herdmania momus
and Phallusia nigra were 9 and 20 mg [ml.sup.-1] respectively.
Origin of test bacteria is indicated by: Lab, Standard laboratory
test bacterium; Sed, Sediment; Sed.w, Sediment water; Seaw, Seawater.
--, no inhibition.

Table 2
Evaluation of crude extract toxicity on brine shrimp

Concentration           P. nigra                H. momus
     (mg           Average mortality        Average mortality
 [ml.sup.-1])      (%) ([+ or -] STD)      (%) ([+ or -] STD)      pH

0 (Seawater                            3.95 ([+ or -] 0.87)       8
  Control)
0 (DMSO                                4.5 ([+ or -] 0.07)        7.5
  Control)
0.01               0.00 ([+ or -] 0.0)            n.d.            7.5
0.05               0.00 ([+ or -] 0.0)            n.d.            7.5
0.10               2.22 ([+ or -] 3.9)            n.d.            7.0
0.25               3.13 ([+ or -] 4.4)            n.d.            7.0
0.50               3.33 ([+ or -] 5.8)            n.d.            7.5
1.00              34.95 ([+ or -] 22.1)           n.d.            7.0
2.00              82.22 ([+ or -] 10.2)    9.17 ([+ or -] 4.6)    n.d.
5.00             100.00 ([+ or -] 0.0)            n.d.            6.5
Natural          100.00 ([+ or -] 0.0)    100.00 ([+ or -] 0.0)   n.d.
  (volumetric)

pH after addition of Phallusia nigra extract as measured by pH paper.
The natural volumetric tissue concentrations of Herdmania momus and
Phallusia nigra were 9 and 20 mg [ml.sup.-1], respectively, n.d., not
determined.
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Date:Dec 1, 2014
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