Coral bleaching is the disruption of the symbiotic association between coral hosts and their photosynthetic microalgal endosymbionts, referred to as zooxanthellae (
12). Coral bleaching events of unprecedented frequency and global extent have been reported during the last two decades (
13). It has been suggested that coral bleaching is triggered by environmental factors which impose stress on the coral. The most frequently reported stress condition is increased seawater temperature (
5,
16). Thus, it is possible that global warming could result in alteration or destruction of coral reef systems. Consequently, it is essential to understand the mechanism(s) of coral bleaching.
Bleaching of the coral
Oculina patagonica from the Mediterranean Sea is the result of a bacterial infection (
17,
18). The causative agent,
Vibrio shiloi, was obtained in pure culture and was shown to cause bleaching in controlled aquarium experiments. Furthermore, it was shown that bacterium-induced bleaching by
V. shiloi could be inhibited by antibiotics. The bacterial infection and resulting coral bleaching were temperature dependent, occurring only at elevated seawater temperatures (25 to 30°C).
Using the
V. shiloi-O. patagonica model system to study coral bleaching, Toren et al. demonstrated that the first step in the infectious process was adhesion of
V. shiloi to a β-galactoside-containing receptor on the coral surface (
32). After
V. shiloi adheres to
O. patagonica, it penetrates into the exodermal layer of the coral (
2). However, the mechanism by which the bacterium kills the algae is unknown. Recently, we reported that
V. shiloisecretes extracellular materials that inhibit photosynthesis and bleach and lyse zooxanthellae isolated from corals (
4). The material responsible for inhibition of photosynthesis was heat stable and was produced only when the bacteria were grown at elevated seawater temperatures.
In the present paper we describe production, purification, and characterization of a proline-rich dodecapeptide from V. shiloi that rapidly inhibits photosynthesis of zooxanthellae in the presence of NH3.
MATERIALS AND METHODS
Bacterial strain and growth conditions.
V. shiloiAK1 (= ATCC BAA-91), isolated from bleached coral as previously described (
6), was used in this study. The strain was maintained on MB agar (1.8% marine broth [Difco] plus 0.9% NaCl solidified with 1.8% agar). After streaking, the plates were incubated at 30°C for 2 days and then allowed to stand for 1 week. For experiments described here, the bacteria were grown in MBT medium (1.8% marine broth, 0.75% tryptone, 0.9% NaCl), CA medium (0.75% Casamino Acids, 2% NaCl), and CAG medium (0.5% Casamino Acids, 0.5% glycerol, 2% NaCl) at 29°C with shaking.
Preparation of zooxanthellae from coral.
Intact colonies of the coral O. patagonica were collected from a depth of 1 to 3 m along the Mediterranean coast of Israel. Within 2 h of collection, each colony was split into several pieces, and the pieces were placed into 2-liter aerated aquaria containing filtered (pore size, 0.45 μm) seawater that were maintained at 25°C. The aquaria were illuminated with a fluorescent lamp by using cycles consisting of 12 h of light and 12 h of darkness. To obtain zooxanthellae, a healthy coral fragment (surface area, ∼1 cm2) was removed from an aquarium and rinsed gently with filter-sterilized seawater, and then the tissue was disrupted with a dental water pick by using ca. 50 ml of sterile seawater. The suspension was centrifuged for 30 min at 2,000 × g . The pellet, resuspended in 1 ml of seawater, was then centrifuged in an Eppendorf centrifuge (model 5402) for 4 min at 10,000 rpm. The pellet, resuspended in 1 ml of seawater, was then centrifuged for 21 min at 1,200 rpm. The final pellet was resuspended in seawater to a concentration of ca. 5 × 106 algae per ml (based on hemacytometer counts). Fresh zooxanthella preparations were used in all experiments.
Purification of toxin P.
Cultures of V. shiloi, grown in MBT medium for 72 h at 29°C, were centrifuged at 12,000 × g for 10 min at 4°C. The supernatant fluid was then passed through a 0.2-μm-pore-size Millipore membrane filter. Ammonium sulfate was added with stirring at 0°C to the cell-free supernatant fluid to a final concentration of 80% saturation. After the preparation stood overnight at 4°C, the precipitate was collected by centrifugation and dissolved in 1/10th the initial volume of water. The concentrated crude toxin P was then extracted three times with an equal volume of ethyl acetate. The ethyl acetate extracts were combined and evaporated to dryness in vacuo at 30°C.
Three sequential columns were used to purify the peptide. The ethyl acetate-extracted material was dissolved in 1 ml of 50 mM Tris HCl buffer (pH 8.0) and applied to a Resource Q column (Pharmacia Bio Tech) with a bed volume of 1 ml and a height of 30 mm. The column was developed with a 0 to 1 M NaCl gradient at a flow rate of 1 ml/min. The active fractions (unconcentrated) were then run on a Superdex Peptide HR 10/30 column (bed volume, 24 ml; particle size, 13 μm; Pharmacia) and eluted with 50% ethanol at a flow rate of 0.25 ml/min. The active fractions were concentrated by evaporation in vacuo. The final purification was on an RP18 hydrophobic column (Merck) at a flow rate of 1 ml/min using increasing acetonitrile (ACN) concentrations.
Measurement of photosynthetic quantum yield of zooxanthellae.
A portable underwater mini pulse-amplitude-modulation fluorometer (Walz) was used to measure the quantum yield of zooxanthellae. This instrument allows direct noninvasive measurement of the effective quantum yield of photosystem II under ambient light conditions (
15,
23-25). Good correlations between measurements of quantum yield and photosynthetic rates (determined by O
2 evolution and CO
2uptake) have been reported for plants (
10) and cyanobacterial symbionts of lichens (
31).
In the experimental procedure used here, the quantum yield of 0.05 ml of zooxanthellae in seawater (5 × 106 algae per ml) was measured in an enzyme-linked immunosorbent assay plate (Y0). Measurements were obtained in the presence of a fluorescent lamp (light intensity, 16 μmol of photons m−2 s−1) at 25°C. Then 0.05 ml of sterile seawater, 0.05 ml of growth medium (controls), or 0.05 ml of an experimental sample was added to the algae, and the kinetics of the quantum yield (Yt) were measured with the mini pulse-amplitude-modulation fluorometer from 1 to 60 min. The percent quantum yields at different times were determined as follows:Yt/Y0 × 100.
Determination of toxin P activity.
In the initial growth experiments, toxin P activity was determined by removing a sample from a culture, centrifuging it to remove the cells, extracting the supernatant fluid with ethyl acetate three times, evaporating the combined ethyl acetate extracts to dryness in vacuo, and dissolving the residue in 50% ethanol. Dilutions of the solution (in sterile seawater) were then used to measure inhibition of photosynthesis of zooxanthellae in the presence of 12.5 mM NH4Cl (unless stated otherwise), as described above. Inhibition due to NH4Cl alone and ethyl acetate-extracted media (controls) was subtracted before toxin P activity was calculated. One unit of toxin P activity was defined as a 10% decrease in the quantum yield after 10 min of incubation. The same procedure was used for assaying purified toxin P except that the ethyl acetate step was omitted.
Micro sequence analysis.
Automated Edman degradation of the purified peptide was performed with pulse liquid automatic sequentor (Applied Biosystems model 473A) by Technion Protein Laboratory, Haifa, Israel.
Solid-phase peptide synthesis of toxin (PYPVYAPPPVVP).
Val-to-Val couplings are often sluggish, and Pro at the carboxyl terminus of a peptide has been associated with high levels of diketopiperazine formation during solid-phase peptide synthesis (
28). Given the fact that the peptide toxin had a Val-Val-Pro sequence at the carboxyl terminus, a synthetic strategy that would minimize diketopiperazine formation during the Val-Val coupling step was developed. Accordingly, synthesis was carried out on a 2-chlorotrityl chloride resin because the bulk of the trityl handle minimized diketopiperazine formation during tripeptide formation. The tripeptide 9-fluorenylmethoxy-carbonyl (Fmoc)–Val–Val–Pro–Resin was synthesized manually, and after evaluation of its quality, chain assembly was completed by using an Applied Biosystems synthesizer.
The protected tripeptide was assembled by manual solid-phase synthesis, starting with H–Pro–2-chlorotrityl chloride resin (0.52 mmol/g). Coupling of Fmoc-Val was accomplished by using the bromo-tris-pyrrolidino-phosphonium hexafluorophosphate–diisopropylethyl amine procedure (
8). Double coupling was carried out at both the di- and tripeptide stages to avoid deletion sequences. Complete coupling was confirmed by using the Kaiser test (
14). A small portion of Fmoc-Val-Val-Pro-resin was cleaved by using a trifluoroacetic acid (TFA)-triisopropylsilane-H
2O cocktail (95:2.5:2.5, vol/vol/vol) to assess the purity of the peptide. High-performance liquid chromatography (HPLC) analysis indicated that the purity of the crude tripeptide was more than 95%. This product was judged acceptable for completion of synthesis.
Chain assembly from Fmoc-Val-Val-Pro-trityl resin was continued in a stepwise manner on a 0.1-mmol scale by using an Applied Biosystems, Inc., model 433A synthesizer. The coupling protocol used was the FastMoc chemistry protocol using 2-(1-H-benzotriazol-lyl)-1,1,3,3-tetramethyluronium hexafluorophosphate–1-hydroxytriazole activation. The Fmocgroup was used for protection of all N-α groups except the N-terminal group, where Boc-Pro was used. Tert-butyl was employed for protection of the Tyr side chain. All Fmoc groups were deprotected in 20% piperidine inN-methylpyrrolidine. After chain assembly was completed, the protected peptidyl resin was treated with TFA-triisopropylsilane-water (95:2.5:2.5, vol/vol/vol) at room temperature for 1.5 h. The reaction mixture was filtered to remove the resin, the resin was washed with TFA, and the combined filtrates were concentrated to a small volume with a rotary evaporator. The crude peptide obtained in this way was precipitated with cold diethyl ether.
The crude peptide was purified on a Waters μBondpack semipreparative C18 column (19 by 300 mm). A water-ACN-TFA gradient was used. The purity of the final product was assessed by using analytical reversed-phase HPLC (Hewlett-Packard series 1050 or 1090 HPLC) with a Waters μBondpack C18 column (3.9 by 300 mm), water-ACN-TFA and water-methanol-TFA elution systems, and detection at 220 and 260 nm. Homogeneity was also assessed by silica gel thin-layer chromatography (Silica Gel 60 precoated aluminum sheet from E. Merck, Darmstadt, Germany) with a gel developed inn-butanol–acetic acid– water–pyridene(9:2:4:3) andn-butanol–acetic acid–water (2:1:1); iodine was used as the developer. Electron spray ionization mass spectrometry was performed at PeptidoGenic Research Company, Livermore, Calif.
DISCUSSION
The data presented here demonstrate that toxin P is a linear, proline-rich dodecapeptide. It is likely that toxin P is produced from a larger peptide by proteolysis. The amino acid sequence of toxin P shows strong similarity (10 of the 12 amino acids are identical) to the amino acid sequence of an internal peptide in the
vrg-6 gene product of
Bordetella pertussis (GenBank accession numberM77374 ). It has been suggested that the
vrg gene products facilitate intracellular survival and the persistence of the bacterium in the host (
3). In this regard it is interesting that
V. shiloi is also an intracellular pathogen (
2).
The high hydrophobicity predicted from the amino acid sequence of toxin P may help explain the observed binding of the toxin to zooxanthellae. Bound toxin P by itself did not affect algal photosynthesis. However, addition of NH
4Cl to algae containing the bound toxin resulted in rapid inhibition of photosynthesis and a concurrent decrease in the pH of the bulk liquid. Inhibition of photosynthesis by ammonia is well established (
1,
7,
35). Ammonia acts as an uncoupler of photosynthesis by passing across membranes, thereby destroying the pH gradient across the thylakoid membrane (
27,
34). Many membrane-penetrating peptides, including melittin (
33), gaegurin (
29), cecropin (
22), and buforin (
21), contain prolyl residues. Although the exact role of prolyl in the function of these peptides is unknown, proline causes kinks in helical polypeptides and increases the flexibility of peptide chains in its immediate environment. Some of these peptides cause lysis of cell membranes, while others act as ion channel formers (
30). In addition, some membrane-active peptides orient parallel to the bilayer, whereas others orient perpendicular to the plane of the membrane (
26). Very recently, a hydrophilic proline-rich domain from the C terminus of L-type calcium channels was shown to remain membrane associated (
11). Cyclic peptides rich in proline have been found to bind alkali metals (
9,
19). Since toxin P is a dodecapeptide, it cannot traverse the membrane bilayer. However, it could form aggregates of the head-to-head or head-to-tail type, as found for gramicidin A. Furthermore, toxin P aggregates in aqueous solution (Fig.
3), supporting the hypothesis that it can enter the algal membrane and act as an ammonia channel. However, the possibility that ammonia activates toxin P has not been eliminated.
Proline-rich antibacterial peptides are also produced by mammalian neutrophils (
20). These peptides are mainly active against gram-negative bacteria. Investigation of the secondary structure of one of these peptides (PR-39), which contains 39 amino acids (19 Pro residues), suggests that it exists in a polyproline II conformation in water. After interacting with the membrane, PR-39 rapidly enters human microvascular endothelial cells and binds to a number of cytoplasmic proteins (
6). Now that pure chemically synthesized toxin P is available, it should be possible to examine its structure and mode of action in more detail. It will also be interesting to examine synthetic peptides with defined amino acid replacements in order to determine their ability to inhibit photosynthesis.
In considering the natural role of toxin P in pathogenesis (coral bleaching), we should mention that high levels of toxin P were found in coral tissues shortly after infection with
V. shiloi(
8). The toxin is produced only at the elevated seawater temperatures (25 to 30°C) necessary for bacterial bleaching of corals (
16). Presumably, once
V. shiloi penetrates into the coral tissue, it produces toxin P and ammonia (from metabolism of coral cytoplasmic protein). The resulting inhibition of photosynthesis in the intracellular zooxanthellae would damage the algae and contribute to coral bleaching (loss of the algae). It should be noted that the equilibrium of NH
4+dissociation to NH
3 is shifted towards increased NH
3 formation with increasing temperature. For example, three times more NH
3 is produced at 25°C than at 10°C at a constant pH and a constant total NH
3-NH
4+ concentration. It should be noted that
O. patagonica in the eastern Mediterranean Sea experiences a shift in temperature from 16°C (winter) to 29 to 30°C (summer, when bleaching occurs) (
16). This may contribute to magnification of the toxin P-enhanced toxicity of ammonia for zooxanthellae in infected coral during the summer.
ACKNOWLEDGMENTS
This work was supported by United States-Israel Binational Science Foundation grant 95-00177, by the Pasha Gol Chair for Applied Microbiology, by the Israel Center for the Study of Emerging Diseases, and by National Institutes of Health grant GM 22086 to F. Naider.
F. Naider is currently a Varon Visiting Professor at the Weizmann Institute of Science, Rehovot, Israel. We thank G. Fleminger for help with the HPLC analysis.