SAMUELSSON, G., AND G. OQUIST. 1980. Effects of copper chloride on photosynthetic electron transport and chlorophyll-protein complexes of *Spiacia oleracea*. Plant Cell Physiol. **21**: 445-454.

SELLNER, K. G., L. LYONS, E. S. PERRY, AND D. B. HEIMARK. 1982. Assessing physiological stress in *Thalassiosira fluviatilis* (Bacillariophyta) and *Dunaliella tertiolecta* (Chlorophyta) with DCMU-enhanced fluorescence. J. Phycol. **18**: 142-148.

SHARP, J. H., M. J. PERRY, E. H. RENGER, AND R. W. EPPLEY. 1980. Phytoplankton rate processes in the oligotrophic waters of the central North Pacific. J. Plankton Res. **2**: 335-353.

SHELDON, R. W. 1984. Phytoplankton growth rates in the tropical ocean. Limnol. Oceanogr. **29**: 1342-1346.

SHIOI, Y., H. TAMAI, AND T. SASA. 1978. Inhibition of photosystem II in the green alga *Ankistrodesmus falcatus* by copper. Physiol. Plant. **44**: 434-438.

SHULENBERGER, E., AND J. L. REID. 1981. The Pacific shallow oxygen maximum, deep chlorophyll maximum, and primary productivity, reconsidered. Deep-Sea Res. **28**: 901-919.

STEEMANN NIELSEN, E., L. KAMP-NIELSEN, AND S. WIUM-ANDERSEN. 1969. The effect of deleterious concentrations of copper on the photosynthesis of *Chlorella pyrenoidosa*. Physiol. Plant. **22**: 1121-1133.

—, AND S. WIUM-ANDERSEN. 1970. Copper ions as poison in the sea and freshwater. Mar. Biol. **6**: 93-97.

STRICKLAND, J. D., AND T. R. PARSONS. 1972. A practical handbook for seawater analysis, 2nd ed. Bull. Fish. Res. Bd. Can. 167.

TUSSEN, S. B. 1979. Diurnal oxygen rhythm and primary production in the mixed layer of the Atlantic Ocean at 20°N. Neth. J. Sea Res. **13**: 79-94.

TRIATHY, B. C., AND P. MOHANTY. 1980. Zinc-inhibited electron transport of photosynthesis in isolated barley chloroplasts. Plant Physiol. **66**: 1174-1178.

VENRICK, E. L., J. R. BEERS, AND J. H. HEINBOKEL. 1977. Possible consequences of containing microplankton for physiological rate measurements. J. Exp. Mar. Biol. Ecol. **26**: 55-76.

VINCENT, W. F. 1980. Mechanisms of rapid photosynthetic adaptation in natural phytoplankton communities. 2. Changes in photochemical capacity as measured by DCMU-induced chlorophyll fluorescence. J. Phycol. **16**: 568-577.

—, P. J. NEALE, AND P. J. RICHERSON. 1984. Photoinhibition: Algal responses to bright light during diel stratification and mixing in a tropical alpine lake. J. Phycol. **20**: 201-211.

WELSCHMEYER, N. A., AND C. J. LORENZEN. 1985. Chlorophyll budgets: Zooplankton grazing and phytoplankton growth in a temperate fjord and the Central Pacific Gyres. Limnol. Oceanogr. **30**: 1-21.

WOOD, A. M. 1983. Available copper ligands and the apparent bioavailability of copper to natural phytoplankton assemblages. Sci. Total Environ. **28**: 51-64.

Submitted: 11 April 1985

Accepted: 10 April 1986

Limnol. Oceanogr., 31(6), 1986, 1373-1376
© 1986, by the American Society of Limnology and Oceanography, Inc.

## A simple method to detect bacterial associations in bivalve gills

*Abstract*—Recent discoveries have shown that endosymbiotic bacteria may be widespread among bivalve species in different habitats. A simple method is described which allows the detection of bacterial-gill association with an ordinary light microscope.

The discovery of a rich fauna in the vicinity of hydrothermal vents raised several important questions about food chain relationships at great depths. A symbiotic association was first shown between chemolithotrophic sulfur-oxidizing bacteria and the gutless worm *Riftia pachyptila* Jones, one of the most abundant species of this

vent fauna (Cavanaugh et al. 1981; Felbeck 1981; Southward et al. 1981). Later, two large species of bivalve Mytilidae were found to carry similar bacteria in their gill tissue (Cavanaugh 1983; Fiala-Medioni 1984; Le Pennec and Hily 1984; Le Pennec and Prieur 1984). These chemolithotrophic bacteria are thought to constitute a major source of the energy and nutrition of their hosts (cf. Janasch 1985). Symbiotic bacteria and possible chemosynthetic capabilities have also been described for animals living in shallow sulfide-rich environments. These include groups such as oligochaetes (*Phallodrilus leukodermatus* and *Phallodrilus planus*:
<figure><img src="image_1.png" /><figcaption>Fig. 1. Results of the method using a gill tissue from *Spisula subtruncata*: photomicrographs of cell suspension, Giemsa staining. A. Dispersed bacteria from packed assemblages (arrow); b—bacteria; n—nuclei of gill tissue. B and C. Two different types of bacteria.</figcaption></figure>

Giere 1981; Felbeck et al. 1983), nematodes (Astomonema jenneri) and turbellarian Parrecatenula sp. (Ott et al. 1982), and bivalves of the family Solemyidae (Cavanaugh 1983; Felbeck 1983; Powell and Somero 1985) and Lucinidae (Cavanaugh 1983; Berg and Alatolo 1984; Fisher and Hand 1984; Vetter 1985). Finally, Dando et al. (1985) have described endosymbiotic bacteria in the gill tissue of the bivalve lucinid Myrtea spinifera, a species which inhabits shallow sea sediments with a low concentration of free sulfides. They also suggested that the bacterial-gill association might be more widespread than hitherto suspected and might contribute in general to the energy needs of the host animals.

The intracellular symbionts in bivalve gill tissue have until now been demonstrated by TEM observations that involve elaborate fixations and special techniques not always available in the field. We describe a simple method to detect bacterial associations in bivalve gill tissue with a transmitted light microscope. Using this method, we have found two different types of bacteria in gill

tissue of the surfclam *Spisula subtruncata* (Da Costa) (Mactridae) and verified the results by TEM observations that showed their intracellular and endosymbiotic character (Bouvy et al. 1986).

The steps of our method follow.

1. Dissect gills from live specimens.

2. Wash in filtered seawater (0.22 µm).

3. Transfer to distilled water or 25% seawater for 30 min (the latter gives better results on photomicrographs).

4. Remove supernatant.

5. Fix in freshly prepared solution: three parts absolute ethanol to one part acetic acid, 4 × 10 min (can be stored at 4°C before further steps).

6. Transfer gill tissue to a depression slide containing 50% acetic acid and macerate with forceps to obtain a cell suspension.

7. Release one drop of cell suspension with a Pasteur pipette from a height of about 10 cm onto a cleaned microscope slide that has been warmed to 44°C (±2°C) (the temperature necessary to split cell membranes).

8. Quickly aspirate excess liquid.

9. Air-dry.
10. Stain for 10 min in freshly prepared Giemsa solution: 4 ml of Giemsa (Merck, Art. 9204); 4 ml of phosphate buffer 0.2 M; pH 6.8; 92 ml of distilled water.

11. Wash in water for 5 min and air-dry.

The preparation can be viewed immediately with the light microscope. For high magnification (×100 optics), immersion oil is placed directly on the dried droplet without need of a cover glass. If wanted, the slide can be made permanent: keep 10 min in xylol and mount in Euparal (or another resin).

This procedure yields preparations of disrupted cells at the periphery of the drop sites. Giemsa stains DNA of nuclei and bacteria. Results with *S. subtruncata* show in each droplet of gill cell suspension many assemblages of bacteria-shaped organelles. We have used a minimum of 10 animals (each gill gave one droplet of cell suspension). We also tested mantle and foot tissue from *S. subtruncata*, and gill tissues from other bivalves (*Mytilus edulis*, *Mytilus galloprovincialis*, *Ostrea edulis*, *Crassostrea gigas*, *Ruditapes philippinarum*, *Venus gallina*); bacteria-shaped organelles were not observed. The results with *S. subtruncata* are shown in Fig. 1. Large numbers of bacteria-shaped organelles were confirmed by TEM to be Gram-negative bacteria (Bouvy et al. 1986) which had been released from their packed assemblages (Fig. 1A). Two different types of bacteria are present, one rod-shaped (about 3 × 0.6 µm) (Fig. 1B) and the other shorter and wider (about 2 × 1 µm) (Fig. 1C).

It is also possible to use the Gram method directly with fresh tissue. The procedure used is as above except that step 5 is omitted and after air-drying, the preparation is Gram-stained. Gram-negative bacteria were found in *S. subtruncata* (Bouvy et al. 1986).

The method can be useful in a systematic search for bacterial associations among bivalves and could also be used to detect surface bacteria as well as bacteria in other tissues of other species. However, it should be considered as a first step to detect bacteria-shaped organelles; further TEM techniques should be used to confirm the characteristics of procaryotic cells. After that, enzymatic assays, chemical analyses, carbon

dioxide fixing, nitrogen fixing, etc. are necessary to determine metabolic pathways and actual symbiotic relationships.

C. Thiriot-Quiévreux

Station Zoologique
06230 Villefranche-sur-Mer
France

J. Soyer

Laboratoire Arago
66650 Banyuls-sur-Mer
France

References

BERG, C. J., AND P. ALATOLO. 1984. Potential chemo-synthesis in molluscan mariculture. Aquaculture 39: 165-179.

BOUVY, M., AND OTHERS. 1986. Sur la présence de bactéries dans la branchie d'un Mollusque Bivalve littoral *Spisula subtruncata* (Da Costa). C.R. Acad. Sci. Paris. 303: 257-262.

CAVANAUGH, C. M. 1983. Symbiotic chemoautotrophic bacteria in marine invertebrates from sulphide habitats. Nature 302: 58-61.

—, S. L. GARDINER, M. L. JONES, H. W. JANNASCH, AND J. B. WATERBURY. 1981. Prokaryotic cells in the hydrothermal vent tube worm *Riftia pachyptila* Jones: Possible chemoautotrophic symbionts. Science 213: 340-342.

DANDO, P. R., E. C. SOUTHWARD, N. B. SOUTHWARD, N. B. TERWILLIGER, AND R. C. TERWILLIGER. 1985. Sulphur-oxidizing bacteria and haemoglobin in gills of the bivalve mollusc *Myrtea spinifera*. Mar. Ecol. Progr. Ser. 23: 85-98.

FELBECK, H. 1981. Chemoautotrophic potential of the hydrothermal vent tube worm *Riftia pachyptila* Jones (Vestimentifera). Science 213: 336-338.

—. 1983. Sulfide oxidation and carbon fixation by the gutless clam *Solemya reidi*: An animal-bacteria symbiosis. J. Comp. Physiol. 152: 3-11.

—, G. LIEBEZEIT, R. DAWSON, AND O. GIERE. 1983. CO₂ fixation in tissues of marine oligochaetes (*Phallodrilus leukodermatus* and *P. planus*) containing symbiotic chemoautotrophic bacteria. Mar. Biol. 75: 187-189.

FIALA-MEDIONI, A. 1984. Mise en évidence par microscopie électronique à transmission de l'abondance de bactéries symbiotiques dans la branchie de Mollusques Bivalves de sources hydrothermales profondes. C.R. Acad. Sci. Paris 298(3): 487-492.

FISHER, R. M., AND S. C. HAND. 1984. Chemoautotrophic symbionts in the bivalve *Lucina floridana* from sea grass beds. Biol. Bull. 167: 445-459.

GIERE, O. 1981. The gutless marine oligochaete *Phalodrilus leukodermatus*. Structural studies on an aberrant tubificid associated with bacteria. Mar. Ecol. Progr. Ser. 5: 353-357.
JANNASCH, H. W. 1985. The chemosynthetic support of life and the microbial diversity at deep-sea hydrothermal vents. Proc. R. Soc. Lond. Ser. B **225**: 277-297.

LE PENNEC, M., AND A. HILY. 1984. Anatomie, structure et ultrastructure de la branchie d'un Mytilidae des sites hydrothermaux du Pacifique oriental. Oceanol. Acta **7**: 517-524.

—, AND D. PRIEUR. 1984. Observations sur la nutrition d'un Mytilidae d'un site hydrothermal actif de la dorsale du Pacifique oriental. C.R. Acad. Sci. Paris **298**(3): 493-498.

OTT, J., G. RIEGER, R. RIEGER, AND F. ENDERES. 1982. New mouthless interstitial worms from the sulfide system: Symbiosis with prokaryotes. Mar. Ecol. (Pubbl. Sta. Zool. Napoli 1) **3**: 313-333.

POWELL, M. A., AND G. N. SOMERO. 1985. Sulfide oxidation occurs in the animal tissue of the gutless clam, *Solemya reidi*. Biol. Bull. **169**: 164-181.

SOUTHWARD, A. J., AND OTHERS. 1981. Bacterial symbionts and low ¹²C/¹³C ratios in tissues of *Pogo-nophora* indicate unusual nutrition and metabolism. Nature **293**: 616-620.

VETTER, R. D. 1985. Elemental sulfur in the gills of three species of clams containing chemoautotrophic symbiotic bacteria: Possible inorganic energy storage compound. Mar. Biol. **88**: 33-42.

Submitted: 11 February 1986

Accepted: 2 July 1986

Limnol. Oceanogr., 31(6), 1986, 1376-1383

© 1986, by the American Society of Limnology and Oceanography, Inc.

A simple fiber-optic microprobe for high resolution light measurements: Application in marine sediment<fnref n="1" />

*Abstract* — A fiber-optic microprobe is described which is inexpensive and simple to build and use. It consists of an 80-µm optical fiber which at the end is tapered down to a rounded sensing tip of 20-30-µm diameter. The detector is a hybrid photodiode/amplifier. The probe has a sensitivity of 0.01 µEinst m⁻² s⁻<fnref n="1" /> and a spectral range of 300-1,100 nm. Spectral light gradients were measured in fine-grained San Francisco Bay sediment that had an undisturbed diatom coating on the surface. The photic zone of the mud was only 0.4 mm deep. Measured in situ spectra showed extinction maxima at 430-520, 620-630, 670, and 825-850 nm due to absorption by chlorophyll *a*, carotenoids, phycocyanin, and bacterio-chlorophyll *a*. Maximum light penetration in the visible range was found in both the violet and the red at ≤400 and ≥700 nm.

Most of our knowledge about light distribution and microalgal photosynthesis in aquatic ecosystems is based on studies in the water column. In many lakes and coastal marine environments, however, benthic microalgae contribute significantly to primary production (e.g. Hunding 1971; Gar-

gas 1971; Cadée and Hegeman 1974; Hartwig 1978). The benthic systems differ from the plankton in certain respects which make the study of microbenthic photosynthesis and its regulation by light experimentally difficult: the photic zone is very narrow in sediments, light scattering is strong, and many benthic microalgae migrate vertically in response to changing environmental conditions such as light.

Earlier techniques for measuring light in sediments have not been able to resolve the spectral light gradients with a high spatial resolution. Light penetration has been studied by covering a large light sensor with millimeter-thin layers of sediment or benthic microalgae (e.g. Hoffmann 1949; Taylor 1964; Taylor and Gebelein 1966; Jørgensen et al. 1979; Haardt and Nielsen 1980) or by inserting a small light probe into the sediment (Fenchel and Straarup 1971). Only in the leaves of higher plants have light measurements recently been made on a very small scale by the use of fiber-optics (Vogelmann and Björn 1984).

Since the development of microelectrode techniques for the measurement of chemical gradients and of photosynthesis at ≤100-

<footnote n="1">¹ Supported by a research fellowship from the National Research Council to B.B.J.</footnote>