Benthic Microbiological Biomass, Composition, Carbon-Utilization Rates, and Production: Summary of Methods for the U.S. JGOFS EqPac Program

Fred C. Dobbs
To my knowledge, there are no JGOFS protocols for sedimentary microbiology. What follows is a distillation of methods from my proposal.

General Sampling Scheme: Subcores will be taken from boxcores and multicores and immediately transferred to a refrigerator or ice tray. The upper 5 cm of sediment will be sampled at 0.5 cm intervals. Between 5 and 10 cm, samples will be collected at 1.0 cm intervals. Finally, a deep horizon, 14--15 cm, will be sampled. Measurements will be normalized to the dry weight of the sediment sample.


Direct Counts:
Sediment samples will be fixed in glutaraldehyde (EM grade, 2 % final conc.) and refrigerated for return to the laboratory at UH. The direct-counting procedure is that of Thistle and Eckman (1990), one that first involves disassociating the bacteria from the sediment, followed by standard acridine-orange staining and enumeration.

Sediment will be extracted in 1.47 M HPO. Samples will be centrifuged and the supernatant frozen for laboratory assay using a luciferin-luciferase bioluminescence reaction (Karl and Craven, 1980).

Phospholipid phosphate:
Sediment samples will be extracted in a chloroform-methanol solution. Following 24 hours of extraction, chloroform and water will be added to excess and the chloroform phase will be transferred to glass tubes having PTFE-lined screwcaps. The solvent-extractable lipids will be dried at 37 C under a stream of nitrogen gas. The extracts will be fractionated on a silicic-acid chromatography column into three general lipid classes: neutral lipids, moderately polar lipids, and polar lipids. An aliquot of the polar-lipid fraction will be analyzed for phospholipid phosphate according to the method of Findlay et al. (1989) to determine total microbial biomass.


I will use phospholipid, ester-linked fatty acids to assess community composition, as they have proven to be reproducible, capable indicators of a wide suite of microbes encountered in the environment. The fatty acids esterified to the phospholipid will be methylated by mild alkaline transesterification of the polar lipid fraction (Guckert et al., 1985). The resulting fatty acid methyl esters will be separated and quantified (Guckert et al., 1985). Structural verification will be done using a HP5995A capillary gas chromatograph/mass spectrometer. Fatty acid profiles will be expressed in absolute values (pmoles/gram dry weight of sediment) and as percentages of the total molar recovery.

Carbon-Utilization Rates:

I will assess microbial utilization of carbon at three stations: 0, 5, and 9N. These stations encompass the range of organic-carbon degradation rates reported by Martin et al. (1990). Sediment will be homogenized, then diluted with filtered (0.2 µm) seawater from the box core or multicore. Aliquots of the slurry will be withdrawn into 3-ml syringes containing a C-glutamic acid/seawater solution, the needles stuck into rubber stoppers, and the syringes placed in cylinders. The cylinders will be pressurized commensurate with the depth at which the sediment was collected. Time-course measurements will be made; samples will be decompressed at 24, 48, 96, and 144 hours and their contents processed as follows. The syringes' contents will be injected into serum bottles containing acid and CO will be collected. Macromolecules will be precipitated with cold acid and collected on membrane filters, which will be dried, then oxidized prior to radioassay (Novitsky and Karl, 1986). Results of the CO assay will represent respiration, those of the macromolecule assay, assimilation. A battery of control samples will be tested for effects of depressurization, abiotic uptake of radioactivity, stimulation of activity by the glutamic acid, and effects of slurrying on microbial activity.


I will use radiolabeled nucleic-acid precursors to estimate growth and production of the microbial community. Growth will be evaluated from the turnover of the adenine nucleotide pool. Production will be estimated based on the incorporation of radiolabel from H-adenine into microbial RNA and DNA. These measurements are interrelated when H-adenine is the precursor; measurement of biomass and production can be made contemporaneously. A bonus to using H-adenine to label DNA is that RNA also is labeled; the ratio of labeled DNA to labeled RNA will be used to monitor the growth stage of the microbial populations.

Sediments will be processed as described above for measurements of carbon-utilization rates, except that the added radioactive material will be 2-H-adenine. Once the samples are pressurized, time-course measurements will be made; samples will be decompressed at 24, 48, 96, and 144 hours and H-RNA, H-DNA, H-ATP and ATP will be extracted and radioassayed. Macromolecules will be collected by filtering the contents of a syringe, then washing the filter with cold acid. Radioactive RNA and DNA will be separated and measured using the technique of Craven and Karl (1984) for marine sediments. Both labeled and unlabeled ATP will be extracted as described above in the ``Biomass'' section. Thus, the specific activity of the ATP pool at each time point will be known and the turnover of the adenine nucleotide pool can be determined (Karl and Bossard, 1985; Karl et al., 1987). Rates of RNA and DNA synthesis will be calculated from the average rate of isotope incorporation divided by the integral of the ATP pool specific activity (Winn and Karl, 1984). Controls analogous to those described for glutamic acid will be run and HO produced through catabolism of radiolabeled adenine will be measured.

There is reason to consider that the disturbance caused by slurrying sediment may yield artifactual data. To evaluate this possibility, I will make a series of comparisons, comparable to those described in Dobbs et al. (1989), between results obtained from slurries and from cores.

Literature Cited

Craven, D.B. and D.M. Karl (1984).
Microbial RNA and DNA synthesis in marine sediments. Marine Biology, 83: 129--139.

Dobbs, F.C., J.B. Guckert and K.R. Carman (1989).
Comparison of three techniques for administering radiolabeled substrates to sediment for trophic studies: Incorporation by microbes. Microbial Ecolology, {\bf 17}: 237--250.

Findlay, R.H., G.M. King, and L. Watling (1989).
Efficacy of phospholipid analysis in determining microbial biomass in sediments. Applied and Environmental Microbiology, 55: 2888--2893.

Guckert, J.B., C.P. Antworth, P.D. Nichols and D.C. White (1985).
Phospholipid, ester-linked fatty acid profiles as reproducible assays for changes in prokaryotic community structure of estuarine sediments. Federation of European Microbiological Societies: Microbiology Ecology, 31: 147--158.

Karl, D.M. and P. Bossard (1985).
Measurement and significance of ATP and adenine nucleotide pool turnover in microbial cells and environmental samples. Journal of Microbiological Methods, 3: 125--139.

Karl, D.M. and D.B. Craven (1980).
Effects of aklaline phosphatase activity on nucleotide measurements in aquatic microbial communities. Applied and Environmental Microbiology, 40: 549--561.

Karl, D.M., D.R. Jones, J.A. Novitsky, C.D. Winn and P. Bossard (1987).
Specific growth rates of natural microbial communities measured by adenine nucleotide pool turnover. Journal of Microbiological Methods, {\bf 6}: 221--235.

Martin, W., M. Leinen, M. Bender, A. Isern, and J. Orchardo (1990).
The relationships between the sedimentary concentrations and rain rates of organic carbon and opal in the equatorial Pacific at 135W. EOS, 71: 124.

Novitsky, J.A. and D.M. Karl (1986).
Characterization of microbial activity in the surface layers of a coastal sub-tropical sediment. Marine Ecology: Progress Series, 28: 49--55.

Thistle, D. and J.E. Eckman (1990).
The effect of a biologically produced structure on the benthic copepods of a deep-sea site. Deep-Sea Research, 37: 541--554.

Winn, C.D. and D.M. Karl (1984).
Laboratory calibrations of the H-adenine technique for measuring rates of RNA and DNA synthesis in marine microorganisms. Applied and Environmental Microbiology, 47:835--842.