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Department of Biological Sciences, Wright State University, Dayton, Ohio 45435
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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Ulkenia sp. zoospores are attracted to 492 nm wavelength light produced by the marine bacterium Vibrio fischeri. Zoospores are positively photoresponsive to wavelengths of 440, 460 and 480 nm and contain a pigment that absorbs blue light. The average velocity of the zoospores is 0.47 m h–1. Stimulatory intensities of these wavelengths ranged from 0.5 to 3.5 µEm–2 s–1 in both laboratory and field studies. The response of this protist to bioluminescence produced by Vibrio fischeri may direct zoospores to a nutrient rich environment colonized by these bacteria. In addition, the greatest responses were found at intensities associated with the light regime found near the bottom of naturally turbid estuaries or at greater depths of nonturbid, offshore waters. Positive phototaxis was not seen in zones of high light intensity either in field or laboratory studies, and there is some indication that zoospores may swim away from high light intensities.
Key words: action spectrum, bacteria, blue light, carotenoid, phototactic response, thraustochytrid, Vibrio fischeri
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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). In many cases the responses are to blue-light wavelengths and are associated with carotenoid pigments. Although the predominant source of light stimulating these responses is the sun, another source should be considered. Light from the sun normally penetrates the top 100 meters or fewer of the ocean. That area is the euphotic zone (Sverdrup et al 1942), but light generated by a wide variety of marine creatures is found at most depths including some of the deepest parts of the ocean. It is interesting to note that the wavelength of this bioluminescent light is nearly identical to the wavelength that stimulates the blue-light response. Although this light is abundant, its intensity is low and probably requires a keenly sensitive receptor to generate a photoresponse.
The ability of only a few marine fungi to respond to light is well documented. Labyrinthula sp. zoospores (Amon and Perkins 1968) and Rhizophydium littoreum Amon zoospores are positively phototactic (Muehlstein et al 1987). Both of these organisms contain yellow or orange pigments, but their action spectra have not been determined. Both were isolated in estuarine areas where a response to ambient light might confer an advantage. Note that Labyrinthula is not a true fungus but is grouped with Ulkenia spp. and other thraustochytrids within the stramenopiles (Leander and Porter 2001). Many recent papers refer to the thraustochytrids as protistans. Fan et al (2002) have described chemotactic responses of thraustochytrids, however, no in depth study of the response of this group to light can be found in the recent literature. Casual observations in our lab never gave a clear indication that Ulkenia sp. isolate SWU2 was attracted to light, but it possessed an orange pigment that looked like it might be a carotenoid, a pigment often associated with photoresponses (Hader 1979, p. 268–309). Simple tests such as those done with R. littoreum did not show a photoresponse, so we devised several experiments to look at a variety of illumination conditions. Our goal was to determine the potential role of the pigment in phototaxis and under what environmental conditions a phototactic response might occur.
Ulkenia sp. studied here belongs to a ubiquitous group of marine protists generally referred to as thraustochytrids. Thraustochytrids usually have a sessile stage that is attached to marine surfaces where they feed using an ectoplasmic net system (Perkins 1972). At maturation the cytoplasm undergoes multiple fissions, releasing motile biflagellate zoospores (Moss 1980). These organisms are important parts of the microplankton and can have biovolumes of up to 43% of the total microplankton (Ragukumar et al 2001). They consume bacteria (Ragukumar 1992) and may constitute an important part of the heterotrophic food web (Naganuma et al 1998). They seem to be particularly important on decaying vegetation (Bremer 1995), particulate organic carbon (Kimura et al 2001) and in sediments (Bonogiorni and Dini 2002). In addition, some thraustochytrids are associated with bioerosion of carbonate materials that are important parts of reef structure (Porter and Lingle 1992). Thraustochytrids recently have received much attention because they contain nutritionally important substances such as carotenes and omega-3 poly-unsaturated fatty acids (Lewis et al 1999, Raghukumar 2002, Aki et al 2003).
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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To maintain cultures, 0.4 mL SWU2 zoospores were spread onto fresh 100 mm Y PDS plates, sealed with Parafilm®, inverted and incubated at 24 C in a normally lighted room. To reduce the possibility of bacterial contamination, cultures were cycled between Y PDS with streptomycin sulfate and ampicillin (100 mg/L each) and Y PDS agar plates without antibiotics. For long-term storage, cultures were maintained in slush agar with sterile pine pollen (Amon and Arthur 1979).
Vibrio fischeri (Beijerinck) Lehman and Neumann (Kreig 1984) was maintained in the dark at 24 C in bioluminescence medium containing 5 g Bacto Tryptone, 2.5 g Bacto Yeast Extract, 0.3 g NH4Cl, 0.3 g MgSO4•7H2O, 0.01 g FeCl3, 1 g CaCO3, 3 g KH2PO4, 10.4 g Na2HPO4, 13.1 g glycerol, 30 g NaCl and 15 g agar adjusted to pH 7 in 1 L of distilled water. The slight precipitate that formed after autoclaving did not affect the use of this media. Aluminum foil wrapped broth tubes with10–15 mL of the medium were inoculated heavily and shaken at 150 rpm to obtain bioluminescent cell suspensions at 18–24 h.
For each experiment, harvested zoospores were centrifuged at about 1000x g for 1 min to form a soft pellet and resuspended in a sterile buffer (CB) containing 6.89 g NaCl, 6.5 mg K2HPO4, 13.5 mg KH2PO4, 1.3 g MgSO4•7H2O, and 0.2625 g CaCl2•2H2O in 500 mL of distilled water. The pH was adjusted to 7.35 with 0.1N NaOH or 0.1N HCl. (Muehlstein 1985).
Laboratory light response experiments. –
We measured pho-totaxis to bioluminescence with a 20 mm diam x 30 mm long blackened Plexiglass® tube with two rubber septa as sampling ports (FIG. 1). At each end of the tube was an opening in which a glass vial could be inserted, one tube for V. fischeri the other for a control vial containing sterile seawater. The 20 mm plastic tube was filled with a zoospore suspension (approximately 2 x 107 zoospores per mL). After acclimation of zoospores to the dark for 5 min, a vial of luminescing V. fischeri and a control were inserted into the ends of the apparatus. All experiments were performed in a darkroom. Samples (0.2 mL) were taken at intervals of 5, 6, 7 and 8 min from both ends of the plastic tube, using a 1 cc syringe fitted with a 0.2 mL stop and an 18 gauge needle. The sampled zoospores were held briefly in capped microfuge tubes and counted at 100x magnification, using a phase-contrast microscope on darkfield setting. Bioluminescence experiments were performed in four replicates at 23–24 C.
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) and comparing the number to a homogeneous suspension of zoospores in a nonilluminated container. Only swimming zoospores were counted.
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Artifical light experiments. – Light was projected from a Kodak Carousel projector (125 V 500 W quartz halogen bulb) onto a front-silvered mirror (heat not reflected) and reflected through specific wavelength filters into the phototaxis block. In the phototaxis block, zoospores attracted to light concentrate where they can be sampled with a syringe and are compared to a nonlighted control. The wavelength of light was varied using Pomfret 10 nm narrowband wavelength filters of 400, 420, 440, 460, 480, 500, 520 nm (Pomfret Research Optics Inc., P.O. Box 1265, Orange, Virginia 22960) and Edmund Scientific 10 nm narrow-band filters of 540, 550, 577, 589, 600 nm (Edmund Scientific, 101 E. Gloucester Pike, Barrington, New Jersey 08007-1380). Light intensity was varied by altering the distance between light source and mirror and calibrated with a photometer (Li-Cor Model LI-185A).
Zoospore velocity. – Zoospore suspensions on a counting chamber with 0.05 mm grids were videotaped on a phase-contrast microscope. The motion of the zoospores relative to the grids was determined by frame-to-frame analysis of numerous straight-line movements to obtain velocity. No attempt was made to provide directional illumination.
Field experiments. – The response of zoospores to natural light intensities was performed in a turbid estuarine channel off Pivers Island at the Duke University Marine Laboratory pier. The apparatus consisted of three sampling units fixed at 2 m intervals along a 3.8 cm x 6 m long PVC pipe. Each unit was equipped with three half-blackened sampling vials containing zoospore suspensions to test migration of zoospores toward the lighted end. Each vial was sampled independently by spring-loaded syringes activated from the surface. Simultaneous sampling at three depths (2, 4, 6 m) was achieved by pulling a cord that removed the actuator pins activating the spring-loaded syringes removing a 0.2 mL zoospore sample from each sampling vial. Vials containing zoospore suspensions were protected from light with aluminum foil until the assembly was immersed. Samples were taken at 0, 2 and 4 min after positioning. Tests were performed on 3 consecutive d within 1 h of noon starting in late June to maximize light availability. All tests were performed at or near high slack tide.
After sampling, the apparatus was removed from the water and the contents of each syringe (0.2 mL) were examined with a phase-contrast microscope at 400x magnification in a Petroff-Hausser bacteria-counting chamber to determine zoospore density in the lighted and unlighted portions of the sample units. Redistribution of zoospores in response to light lead to a higher concentration in one end of the chamber and a corresponding decreased concentration in the opposite end of the chamber. Light intensity readings were taken on the surface and at each sample depth before and after each experiment.
Pigment extraction. – SWU2 was maintained in Y PM (1 g yeast, 1 g peptone and 5 g maltose per L of Instant Ocean Seawater® at 20 ppt salinity) broth. Y PM was chosen because we noted its pigment-enhancing characteristics with SWU2. Cultures were grown at 23–24 C for 7 d on a gyro-rotary shaker at 150 rpm. After incubation, the cells were centrifuged to form a pellet. Liquid was decanted and the pellet was resuspended in 95% methanol, packed in ice and sonicated (Sonicator cell disrupter Model W200R) for 15 s at 25 000 Hz. Tubes of homogenate then were wrapped in aluminum foil and placed on a gyrorotary shaker at 150 rpm for 24 h at room temperature to complete the extraction. Cell debris was removed by centrifugation, and the liquid was concentrated with a nitrogen gas stream. Analysis was with a Beckman DU-40, spectrophotometer (Beckman Instruments Inc., Irvine, California).
Statistical analysis. – Because experiments over a range of intensities or wavelengths required long periods of time and several individual batches of zoospores, the number of zoospores varied from one experiment to the other. Each experiment was repeated 3 or 4 times, and the data from each were normalized to reflect percent change from a nonilluminated control. Analysis of variance, paired student t-testing and Tukey’s studentized range tests were performed on data collected from all phototaxis experiments. Since we could not entirely control mixing associated with sampling, we chose to consider probabilities at the P = 0.1 as significant differences from the controls.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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). Zoospore density at the illuminated end increased by 203% while decreasing by 181% at the non-illuminated end. Thus, control and illuminated zoospore populations were significantly different (P = 0.01). Light intensity estimated from bacteria spread on a flat surface was approximately 0.4–2.6 µEm–2 s–1.
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). High intensities of white and blue light gave no response and zoospores responded negatively to blue light at 250 µEm–2 s–1 and above (FIG. 4). Positive white light responses were observed at intensities between 0.5 and 25 µEm–2 s–1 and positive blue-light responses were observed for intensities between 0.5 and 3.5 µEm–2 s–1. Individual wavelength filters gave positive responses at 420, 440, 460, 480 and 500 nm at intensities of 3.5 µEm–2 s–1, but no responses were observed at wavelengths of 400 nm or from 520 through 600 nm (FIG. 5). The strongest response was at 480 nm, coincidentally near the single 470 nm absorption peak of pigment extracted from zoospores (FIG. 5). In low intensity white light from the microscope and without a confining cover glass, zoospores moved at a velocity of 0.47 m h–1 (131 µm s–1) while traveling in straight lines.
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), which is very similar to the white light response seen in the laboratory where no positive responses were noted above 25 µEm–2 s–1. The data for these experiments, gathered over three days, show suggestive trends, but variation was high due to vibrations induced by ambient water currents outside the chambers, and because only one of the three days produced statistically significant results. In spite of that, the trends on each day were similar. In each experiment the maximum response was at the 6 m depth, where light intensity was lowest (0.5–4.8 µEm–2 s–1). The greatest response (P = 0.05) was when the light intensity was 0.5 µEm–2 s–1, the lowest light measured in the three day experiment.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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Bioluminescence is common in the sea (Widder et al 1983), but the light produced by mobile organisms may be inconsequential if those organisms move faster than the zoospore’s velocity of only 0.47 m h–1. A more likely situation would be attraction to stationary targets. Numerous sessile organisms display bioluminescence, and attraction to these organisms could enable Ulkenia sp. to parasitize the organism or to live saprophytically on its byproducts. Another possibility is that bioluminescence may be produced by bacteria involved in biodegradation. Bioluminescent bacteria such as Vibrio fischeri are well known from live and decaying fish (Madigan et al 2003, 380) and may be involved in decay of nearly any organismal matter. Decaying materials may provide a rich source of food and could promote rapid and robust increases in the local population of Ulkenia sp. Raghukumar et al (2001) found thraustochytrids in the open ocean at depths of 150 m, probably associated with decay of phytoplankton, in areas of high bacterial concentrations. They also noted thraustochytrids in dense populations at depths up to 2000 m with a probable correlation to decaying organic matter.
The specific light responses of this organism also may help it avoid competition with other ecologically similar organisms. Rhizophydium littoreum Amon (Muehlstein et al 1987) also responds to blue light but is less photophobic to the higher intensities. R. littoreum responds positively to blue light at intensities of up to 300 µEm–2 s–1, which may cause its population to dominate higher in the water column, as opposed to Ulkenia sp. SWU2. Other organisms have been found to be photoresponsive at low intensities of light. Euglena gracilis is photoresponsive at 3–60 µEm–2 s–1 intensities and Dictyostelium purpureum is photoresponsive at intensities of 0.–0.3 µEm–2 s–1 (Carlile 1970, 309–344), so responses shown by SWU2 are in the same light intensities as several other microbes.
Night and daytime responses may differ among phototactic microbes. Attraction to bioluminescence enables responses at anytime in the aphotic zone, whereas attraction to light produced by the sun is intermittent. Zoospores moving upward from the dark depths in the daytime may be attracted downward to sessile sources of biologic light at night. Sunlight may cause a negative response during daytime, causing zoospores in some estuaries to remain below a certain depth by photophobic responses. With photoresponses constrained below intensities of 25 µEm–2 s–1 and wavelengths of 440–500 nm, results indicate a form of light stratification that could play an important role in the distribution of SWU2 in day and night waters.
Chemotactic responses in thraustochytrids recently have been shown by Fan et al (2002), and we have seen (unpubl observations) strong chemotactic responses in SWU2 as well. A combination of phototactic and chemotactic responses may be useful to the organism searching for nutrient. Phototactic clues are not affected greatly by currents, but currents may dilute or displace chemical signals. A positive phototactic response may position a zoospore close enough to food to allow chemoresponses to become significant for further positioning. The use of blue light as stimulus is useful because that wavelength has a greater ability to penetrate seawater than the longer wavelengths of visible light. Blue light from sunlight is present at sufficient intensities to stimulate phototaxis in the photic zone (Clarke and Denton 1966) and also, as we have shown, sufficient light as bioluminescence is available at any depth.
Pigment extractions from Ulkenia sp. SWU2 revealed an absorption peak at 470 nm with a range of 440–550 nm, which correlates with phototaxis responses of zoospores at 460 and 480 nm and is indicative of a blue-light response. This result may indicate that the photoreceptive pigment in SWU2 zoospores and the pigment extracted from SWU2 are the same. The response is characteristic of carotenoid pigments found in other organisms exhibiting a blue-light response (Ninneman 1980). Additionally, SWU2 zoospores are attracted to bioluminescent light produced by Vibrio fischeri (492 nm). Although results show an absorption peak of SWU2 pigment is at 470 nm, the action spectrum shows a peak at 480. Skewing of the absorption spectrum and the action spectrum may be caused by a protein-pigment interaction. Proteins, which are associated with the pigment but are not an integral part of the pigment, may cause a shift in the wavelength of absorption. Because we used fixed wavelength filters, no measurements between 441–460 nm or 480–499 nm could be tested, so the peaks of the photoresponses are estimates. A photoresponse to the 492 nm light produced by Vibrio fischeri could not be tested precisely in our experiments. Photoresponses of SWU2 do fall within the scope light produced by most (FIG. 5
) bioluminescent organisms (Kelly and Tett 1978).
Vibrio spp. is known to degrade cellulose (Perkins 1974) and chitin (Sera and Ishida 1972) into smaller glucose units of which a small portion is metabolized and the remainder is released into the surrounding aquatic environment (Seki 1982). SWU2’s ability to respond to bioluminescent light produced by Vibrio fischeri may direct zoospores to a nutrient rich environment. Thraustochytrids are known to parasitize brown algae (Miller and Jones 1983) and are found in both corals and coral mucus (Ragukumar and Balasubramian 1991). Thraustochytrid zoospores are grazed upon by protozoa within the oceanic food chain (Ragukumarumar and Balasubramian 1991). Evidence shows that thraustochytrids are associated with bottom sediments, particulate organics and detritus (Bremer 1995, Kimura et al 2001, Bonogiorni and Dini 2002) where bioluminescent sources such as V. fischeri may grow. In addition, thraustochytrids may consume the bacteria that attract them (Ragukumar 1992). By combining phototaxis and chemotaxis (Fan et al 2002), zoospores of thraustochytrids increase their chances of finding optimal nutritional substrates for growth and reproduction.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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1 Corresponding author. E-mail: james.amon{at}wright.edu
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LITERATURE CITED |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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Amon JP, Perkins, FO. 1968. Structure of Labyrinthula sp. zoospores. Jour Protozool 15:543–546.
———, Arthur RD. 1979. Agar slush as a method to insure homogeneity of monocentric fungi in continuous culture. Appl and Environ Microbiol 30:763–764.
Bongiorni L, Dini F. 2002. Distribution and abundance of thraustochytrids in different Mediterranean coastal habitats. Aquatic Microbial Ecology 30:49–56.
Bremer GB. 1995. Lower marine fungi (Labyrinthulomycetes) and the decay of mangrove leaf-litter. Hydrobiologia 295:89–95.
Carlile MJ. 1970. The photoresponses of fungi. Photobiology of microorganisms. New York: John Wiley & Son. 479 p.
Clarke GL, Denton EJ. 1966. Light and animal life. In: Hill MN, ed. The sea. Vol. 1. Physical oceanography, ideas and observations on progress in the study of the seas. New York: Interscience Publishers. p 456–468.
Fan KW, Vrijmoed LLP, Jones EBG. 2002. Zoospore chemotaxis of mangrove thraustochytrids from Hong Kong. Mycologia 94:569–578.
Hader DP. 1979. Physiology of movements. In: Haupt W, Fernlab W, eds. Control of locomotion, photomovement. New York: Springer-Verlag. p 268–309.
Kelly MG, Tett P. 1978. Bioluminescence in the ocean. In: Herring P, ed. Bioluminescence in action. New York: Academic Press. p 399–418.
Kimura H, Sato M, Sugiyama C, Naganuma T. 2001. Coupling of thraustochytrids and POM, and of bacterio-and phytoplankton in a semi-enclosed coastal area: implication for different substrate preference by the planktonic decomposers. Aquatic Microbial Ecology 25:293–300.
Krieg NR. 1984. Bergey’s manual of systematic bacteriology. Vol. 1. In: Holt JG, ed. Baltimore: Williams and Wilkins. 964 p.
Latz MI, Frank TM, Case JF. 1988. Spectral composition of epipelagic organisms from the Sargasso Sea. Marine Biology 98:441–446.
Leander CA, Porter D. 2001. The Labyrinthulomycota is comprised of three distinct lineages. Mycologia 93:459–464.
Lewis TE, Nichols PD, McMeekin TA. 1999, The biotechnological potential of thraustochytrids. Marine Biotechnology 1:580–587.
Madigan MT, Martinko JM, Parker J. 2003. Brock biology of microorganisms. 10th ed. Upper Saddle River, New Jersey: Prentice Hall. 1019 p.
Miller JD, Jones EBG. 1983. Observations on the association of thraustochytrid marine fungi with decaying seaweed. Botanica Marina 12:345–351.
Moss ST. 1980. Ultrastructure of the endomembrane—sagenogenetosome—ectoplasmic net complex in Ulkenia visurgensis (Thraustochytriales). Botanica Marina 23: 73–94.
Muehlstein LK. 1985. Chemotaxis and phototaxis in the marine fungus Rhizophydium littoreum [Master’s thesis]. Dayton, Ohio: Wright State University. 81 p.
———, Amon JP, Leffler DL. 1987. Phototaxis in the marine fungus Rhizophydium littoreum. Appl Env Micro 53:1819–1821.
Naganuma T, Takasugi H, Kimura H. 1998. Abundance of thraustochytrids in coastal plankton. Marine Ecology—Progress Series 162:105–110.
Ninneman H. 1980. Blue light photoreceptors. Bioscience 30:69–93.
Perkins FO. 1972. Ultrastructure of holdfasts, ‘‘rhizoids’’, and ‘‘slime tracks’’ in thraustochytriaceous fungi and Labyrinthula spp. Arch Mikrobiol 90:1–17.
Perkins EJ. 1974. The biology of estuarines and coastal waters. New York: Academic Press. 678 p.
Porter D, Lingle WL. 1992. Endolithic thraustochytrid marine fungi from planted shell fragments. Mycologia 84: 289–299.
Ragukumar S. 1992. Bacterivory—a novel dual role for thraustochytrids in the sea. Marine Biology 113:165–169.
———. 2002. Ecology of the marine protists, the Labyrinthulomycetes (thraustochytrids and labyrinthulids). European Jour of Protistology 38:127–145.
———, Balasubramian R. 1991. Occurrence of thraustochytrid fungi in corals and coral mucus. Indian Journal of Marine Sciences 20:176–181.
———, Ramaiah N, Ragukumar C. 2001. Dynamics of thraustochytrid protists in the water column of the Arabian Sea. Aquatic Microbial Ecology 24:175–186.
Seki H. 1982. Organic materials in aquatic ecosystems. Boca Raton, Florida: CRC Press. 201 p.
Sera H, Ishida Y. 1972. Bacterial flora in the digestive tracts of marine fish. Classification of isolated bacteria. Bull Jap Soc Sci Fish 38:633–859.
Sverdrup HU, Johnson MW, Fleming RH. 1942. The oceans, their physics, chemistry and general biology. New York: Prentice Hall. 1060 p.
Widder EA, Latz MI, Case JF. 1983. Marine bioluminescence spectra measured with an optical multichannel detection system. Biol Bull 165:791–810.
Wolken JJ. 1975. Photoprocesses, photoreceptors, and evolution. New York: Academic Press. 317 p.
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