Catalysts or contributors? Tracking bacterial mediation of early diagenesis in the marine water...

Pergamon Org. Geochem. Vol. 26, No. 9/10, pp. 531-544, 1997

© 1997 Elsevier Science Ltd. All rights reserved Printed in Great Britain

PII: S0146-6380(97)00033-8 o146-638o/97 $17.o0 + o.oo

Catalysts or contributors? Tracking bacterial mediation of early diagenesis in the marine water column

H. R O D G E R H A R V E Y t* and S T E P H E N A. M A C K O 2

'Chesapeake Biological Laboratory, The University of Maryland Center for Environmental Science, Box 38, Solomons, MD 20688, U.S.A. and 2Department of Environmental Sciences, Clark Hall, The

University of Virginia, Charlottesville, VA 22903, U.S.A.

Abstract--A series of oxic and anoxic incubations examined the changes in bacterial biomass and diagnostic fatty acids during degradation of two marine phytoplankton, the diatom Thalassiosira weissflogii and the coccoid cyanobacterium Synechococcus sp. Flow-through systems were used to simulate sedimentation, while macrozooplankton grazers were excluded. Bacterial abundance was strongly coupled to the decay sequence for algal carbon under both oxic and anoxic conditions, with loss of particular organic carbon (POC) accompanied by a rapid increase in bacterial biomass. Bacterial carbon increased over 3.7-fold during the period when maximal loss of diatom derived POC was observed. In the oxic cyanobacterial decay experiment bacterial biomass showed a different pattern, with a rapid increase prior to the time period of particulate carbon loss and decrease thereafter. The total concentration of "bacterial" fatty acids for all incubations increased from time zero, with highest concentrations (164/tg l -l) in oxic incubations and coinciding with periods of highest bacterial carbon. Bacterial carbon accounted for up to 32.6% of total POC during the early stages of anoxic diatom decay, with lower maximum amounts (21.4%) in parallel incubations under oxic conditions. Carbon isotopic signatures of individual fatty acids representing algal, bacterial and mixed sources were determined during oxic diatom decay, and showed a small enrichment (average of + 1.6%o) in 13C after the period of major POC loss. A comparison of total fatty acids attributed to bacteria and bacterial carbon calculated from measured bacterial cells showed no strong relationship, suggesting that fatty acids considered diagnostic of bacteria remain a qualitative tool for the estimation of bacterial contributions to sediments. © 1997 Elsevier Science Ltd

Key words--Bacteria, lipid, diagenesis, fatty acids, isotopes, preservation, anoxia

INTRODUCTION

The intimate association of bacteria with the degradation of particulate organic matter is well established. As consumers of both excreted dissolved constituents and colonizers of degrading particles, bacteria and other members of the microbial consortia are the primary recyclers of organic matter in the marine water column (Azam et al., 1983; Deming and Baross, 1993). The small fraction of organic matter which reaches underlying sediments in oceanic systems is testament to the efficiency of the process (for reviews see Emerson and Hedges, 1988; Wakeham and Lee, 1989). Bacteria also con- tribute to the particulate pool, transforming a fraction of dissolved substrates into new cells, which are then themselves subject to consumption by microeucaryotes or degradation. As a result, particulate organic carbon (POC) in marine sediments is a complex mixture of detrital material from water column transport and in situ synthesis during and following its descent.

In unraveling the myriad of organic sources to sediments and its preservation, an important ques-

*Corresponding author (Tel: 410-326-7206; Fax: 410-326- 7341; e-mail: [email protected]).

tion is the recalcitrance of the bacteria themselves, i.e. do they make a significant contribution to the

organic matter which is ultimately buried in marine

sediments? Many studies have documented the ac-

tivities of the bacterial assembledge during early diagenesis in the marine system during field (Newell

and Linley, 1984; Lee, 1992 and review by Deming

and Baross, 1993) and laboratory experiments (Spooner et al., 1994; Harvey et al., 1995), as well

as the presence of compounds of bacterial origin in

field collections (McCaffrey et al., 1989; Wakeham and Beier, 1991; Haddad et al., 1992; Fabiano and

Danovaro, 1994; Canuel and Martens, 1996), yet

the contribution of bacteria themselves as a source of organic matter to sediments remains uncertain.

A major problem is separating the contribution of

living bacteria from that of other sources of organic matter. The biochemical composition of bacteria is

generally similar to the organic matrix that sur- round them, making distinguishing each an impos- ing task. For example, the amino acids of phytoplankton, which comprise the major source of

organic matter, and amino acids of bacteria are very similar (Cowie, 1990), and only in rare cases does the distribution in sediments suggest distinct sources

531

532 H.R. Harvey and S. A. Macko

(Whelan and Emeis, 1992; Cowie and Hedges, 1994).

Attention to resolve this overlap has focused on lipids, including phospholipids or its components (Findlay et al., 1989), and in particular fatty acids which have served as surrogate measures of total biomass (Findlay and Dobbs, 1993 and references therein). Fatty acids have received widespread attention from the standpoint of specific processes (e.g. sulfate reducers; Taylor and Parkes, 1985) and as potential indices of changes in community structure or specific bacterial groups (Perry et al., 1979: Shiea et al., 1991; Goossens et al., 1989). The use of fatty acids as bacterial markers relies on specific structures rarely present in eukaryotic systems, typically odd and branched chain fatty acids (Ratledge and Wilkinson, 1989; Kaneda, 1991) or occasionally fatty acids in the trans-configuration (Gillan et al., 1981; Guckert et al., 1987).

For this study we focused on two major ques- tions. First, are bacteria a significant fraction of the organic carbon pool during early diagenesis? Using flow through systems to mimic the crash and sub- sequent sedimentation of blooms of two important phytoplanktons, a diatom and a cyanobacteria, we examined the contribution of bacteria and changes in bacterial biomass during mineralization. Epifluorescent microscopy was performed to quantify the importance of bacterial carbon during the degradative process. Oxic and anoxic conditions were examined as a major variable to consider the importance of depositional environment on bacterial dynamics. The second major question addresses the ability of diagnostic fatty acids attributed to bacteria to quantify inputs: how do they compare to direct cell observations? As conservative membrane components, fatty acids derived from bacteria might be expected to show a simple relationship with bacterial carbon content. To investi- gate this possibility, bacterial fatty acid concentrations were determined over time and compared with bacterial carbon, calculated from direct determination of bacterial biomass. Laboratory incubations have observed that bacterial phospholipids are rapidly hydrolyzed in sediments, suggesting that the presence of bacterial lipids may represent intact cells (Harvey et al., 1986). By tracking bacterial biomass and fatty acids considered to be derived solely from bacteria, our goal was to compare the changes in direct and chemically determined measures of bacterial biomass during the early diagenesis of diatom and cyanobacterial carbon under conditions of variable oxygen availability.

METHODS

Cultures

The diatom Thalassiosira weissflogii (CCMP 1336) was cultured in F/2 medium (Guillard, 1972)

prepared with 0.2/~m filtered and autoclaved estuar- ine water (salinity 20%o). The coccoid cyanobacterium, Synechococcus sp. (clone WH7803) was cultured similarly with salinity increased to 30%o with natural sea salts. Both cultures were maintained axenically under constant (19°C) temperature with a 12:12 h light:dark lighting regime provided by cool white fluorescent bulbs. Cell growth was monitored by cell counts using inverted microscopy.

Description o[ experimental sys tem

To simulate the fate of phytodetritus during sedimentation through the water column, the experimental system was designed to minimize handling and potential contamination while allowing exposure to the natural microbial community as described in detail previously (Harvey et al., 1995). Briefly, algae were first cultured axenically in 421 glass carboys under conditions similar to those described above. Cultures were continually aerated (500mlmin -I) during growth with hydrocarbon and particle-free filtered air. Growth was monitored by cell counts and the culture allowed to reach maximal density. Once cultures reached the early stationary phase of growth, decomposition experiments were started by enclosing the vessel in dual thickness black plastic bags to place it in darkness. Flow through the system was initiated at this time using a peristaltic pump which transferred filtered (5 ~m) sea water at a known rate (5% exchange day l) from a header vessel into the algal suspen- sion through PTFE lines with glass connectors. This procedure allowed dilution of any remaining nutrients or metabolites of cell growth by addition of screened sea water, minimized the potential effects of cell harvesting and resuspension and provided a constant input of the natural microbial community. As a consequence, these systems were not designed to allow a mass balance of total carbon, but rather the characterization of particulate material and its biochemical composition under controlled conditions during the decomposition process.

Identical designs were used for both oxic and anoxic experiments with the principal differences being the incoming water supply and purge gases. For oxic experiments, incoming water was aerated to a constant oxygen tension before entering the culture vessel and the vessel itself was gently aerated to maintain oxic conditions. For anoxic incubations, incoming water was initially purged with argon to remove oxygen and small amounts of neu- tralized NaS.9H20 added to scavenge any remaining oxygen. At time 0, anoxic cultures were also purged with argon, with a continual flow (60mlmin -~) across the headspace of the vessel used throughout the experimental period to exclude oxygen. Addition of anoxic Chesapeake Bay water to the header (5L) and culture (IL) vessel assured a

Catalysts or contributors 533

viable anaerobic microbial inoculum. Hydrogen sulfide in the experimental vessel and header tank were routinely monitored to assure anoxic conditions were maintained throughout anoxic decay experiments. Average sulfide concentrations were 17/~M and 47/~M over the course of the anoxic treatments of diatoms and cyanobacteria, respectively. For both experimental treatments, flow rates into culture vessels were kept at 2 1 day - l (equivalent to 5% exchange) to eliminate one possible variable in the two treatments. All parameters in both oxic and anoxic experiments were corrected for removal of particulate samples and resulting dilution as each incubation progressed.

Experimental sampling and measurements

At defined time points during each decomposition experiment, samples were withdrawn from each vessel for analysis. Before sampling enclosure bags were removed and the entire vessel gently stirred for 10 min to allow a homogenous sample to be withdrawn. While stirring continued, samples were aseptically removed through the sampling port (PTFE line) and processed immediately for analysis. We observed that large aggregates which formed infrequently during the incubations were disrupted by the mixing procedure, but the need for consist- ency in sampling outweighed the potential for significantly changing decomposition rates by altering particle interactions. Particulate material for CHN analysis and lipids were collected by filtration through combusted (450°C for 4 h) G F / F filters. Particulate organic carbon and particulate nitrogen was quantified by standard methods (Bratback, 1987). Dry weight was measured by filtering a known volume through tared 0.2/~m polycarbonate filters (Gelman), followed by drying to constant weight (50°C) and correction for salt retention. Samples for total bacterial numbers and biomass (15 ml) were collected in glass scintillation vials and preserved with particle-free 37% formaldehyde (final concentration 2%) and held at 4°C until slide preparation. Unfiltered material for microscopic analysis was fixed in buffered 0.2% glutaraldehyde. Additional samples were withdrawn for measurement of bacterial production (thymidine) and activity (glucose and amino acid uptake) as previously reported elsewhere (Harvey et al., 1995).

Fatty acids

For individual component analysis, an internal standard of nonadecanoic acid was added to one quarter of the lipid extract and the fraction sub- jected to mild alkaline hydrolysis using 0 .2N methanolic KOH with gentle heating (70°C, 1 h). After cooling and addition of water, neutral lipids were partitioned into hexane:diethyl ether (9:1). Fatty acids were partitioned following acidification to pH 2 and converted to the fatty acid methyl

esters (FAMES) using BF3 in methanol (70°C, 1 h). FAMES were further purified by partitioning into hexane:diethyl ether (9:1). FAMES at each time point were quantified by capillary gas chromatog- raphy (GC) using a Hewlett Packard 5890A instru- ment with flame ionization detection. Separations were performed with a 5% cross-linked methyl sili- cone column (DB-5, 30m length, 0.32mm i.d., 0.25 pm film thickness) with hydrogen as the carrier gas. For the polar fraction, the temperature program was 50-120°C at a rate of 10°Cmin -~ followed by a 4°C min -1 to 300°C. Chromatographic data were collected using a dedicated data system (Chemstation-Hewlett Packard) and quantified by comparison of peak areas with the internal standard. Structural identifications were performed using two instruments. Electron impact (EI) mass spectra were obtained by interfacing an identical capillary GC with a Hewlett Packard 5970 mass selective detector operating at 70 eV with acqui- sition over the 50-600 amu range. Structural assignments were made using a combination of comparison with authentic standards and mass spectral interpretation. Confirmation of molecular mass assignments obtained in the EI mode was made using chemical ionization GC/MS with a HP 5889a and CH4 as the reagent gas (1.7 torr). Similar operating conditions to those described for GC were used for both instruments with the exception that helium was the carrier gas and a 0.2 mm i.d. column was employed.

Isotopic analysis of individual fatty acids

A portion of the total lipid extract was dried and converted to CO2 for isotope analysis by the Dumas sealed-tube method as described by Macko et al. (1987). The sample was mixed with high- purity copper (Alpha Resources) and precombusted copper oxide in a quartz tube, the solvent was evap- orated, and the tube was then evacuated, sealed and combusted at 850°C for 1 h. The COz was isolated cryogenically and analyzed with a PRISM isotope ratio mass spectrometer (V.G. Isogas). The stable isotopic ratio is reported as follows:

613C = [ R sample ] L R ~ d J - 1 x 1000%o

where R = '3C/12C. The standard for carbon is the Peedee Belemnite limestone that has been assigned a ~3C value of 0.0%0. Precision in the measurement of replicate samples is typically _+0.1%o. In the laboratory, the samples are commonly measured against a tank of carbon dioxide which has been calibrated against NBS 22.

The isotopic abundances of individual compounds from the fatty acid mixture were obtained using a Hewlett Packard 5890 Series II gas chro- matograph (GC) coupled through a combustion furnace and water trap to the VG PRISM isotope


ratio mass spectrometer (IRMS). The GC was equipped with the 50 m HP1 column, with operating conditions being: He flow 1 ml min- l ; injector temperature 275°C; detector 300°C; ramp from 100 to 280°C at 6 ° min -z. A heart split valve was used to direct the flow of the column effluent to the F ID for peak detection or through the combustion furnace to the IRMS for isotopic analysis. The combustion furnace consisted of a quartz tube packed with CuO granules and was maintained at 800°C. The effluent from the furnace passed through a cryogenically cooled trap to remove water vapor before entering the mass spectrometer, and the sep- arated compounds were analyzed individually as pulses of CO2. A naphthalene-d8 standard (99%, Cambridge Isotope Laboratories) of known isotopic composit ion was co-injected with each sample so that instrumental performance could be monitored. Three replicate analyses were performed for most samples. Instrumental precision varied with each peak and depended on peak size and resolution from adjacent peaks. Typical precisions (l SD) for sample components ranged from 0.1 to 0.5%0. The fatty acids were corrected for the contribution of the methyl group addition to make the respective methyl esters (FAMES) injected on the GC as described elsewhere (Ballentine et al., 1996). This contribution typically results in a 0.5%o addition to the value actually derived on the GC/C / IRMS.

Bacterial biomass

Bacterial biomass was determined at each time point from microscopic observations of acridine orange-stained cells collected on irgalan black stained Nuclepore 0.2/~m membrane filters (Watson et al., 1997). Cellular biovolume was calculated based on three categories: 1/~m (average size 0.75tma diameter spheres), 1 -2 t im rods (average 1.5/~m length by 0.5 :tm width), and >2/~m rods (average size 2 .5#m length by 0 .5#m width). Although a range of values are discussed in the lit- erature (e.g. Lee and Furham, 1987; Nagata and Watanabe, 1990), total bacterial abundances were converted to carbon biomass by multiplying the total biovolume by the conversion factor 0.310 pgC ltm -3 recommended by Fry (1990). Weighted averages were calculated to estimate bacterial carbon contribution to the total particulate material.

RESULTS

Bacterial carbon during degradation

Bacterial abundance appears strongly coupled to the decay sequence for algal carbon during the simulated sedimentation (Fig. 1). The relationship was most clearly seen during oxic decay of diatom cells, in which the rapid rate of POC loss was accompanied by a rapid increase in bacterial biomass (Fig. l). Bacterial carbon increased nearly 4-fold

t : t ,

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Time (Days)

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Fig. 1. Time course of concentrations for total particulate carbon and bacterial carbon derived from direct counts during oxic and anoxic decay of the diatom (T. weiss[togii) and cyanobacterium

(Synechococcus sp.) cells. Note different scales for various incubations.


during the initial 20 days when maximal loss of diatom POC was observed, from 4.7x 106cellsml - l (equivalent to 412pgca rbon l -~) at day 1 to 1.8 x 107 cells (1455 #g C 1 - t ) at day 15. In contrast, during oxic cyanobacterial decay bacterial biomass showed a different pattern, with a rapid increase prior to the period of particulate carbon loss. Bacterial abundances increased from 2.8 x 105 cells ml -I (25/~g C l -I) at day 0 to a maximum range of 8.6 x 106 cells ml -~ to 1.3 x 107 over days 1-5 (756-1086/~gC1-1), but declined thereafter. The decline in bacterial abundance during maximal loss of particulate material suggests that labile carbon pools present at the start of the cyanobacterial experiment rapidly decreased over time compared to in diatom incubations. Alternatively, the consumption of bacteria by grazing microzooplankton such as ciliates and flagellates, which would not be excluded by the filtration steps used, could be responsible for the rapid decline in bacterial numbers seen during the incubations (e.g. Neweli et al., 1981). Bacterial predation by flagellates or other protists appears particularly likely for the diatom incubations where the rapid reduction in bacterial numbers appears much greater than could be accounted for by dilution with incoming waters which contained significantly fewer bacteria.

In contrast to the variable response of the bacterial community in oxic waters, bacterial populations during anoxic phytoplankton decay responded similarly to both diatom and cyanobacterial carbon as substrates. After introduction via inflow waters, bacterial biomass increased dramatically throughout the first 13 days of the diatom incubation, with total numbers reaching more than 17-fold higher than those in incoming waters (Fig. 1). This response mir- rors that observed for oxic diatom incubations except that the peak in total bacterial numbers was over twice that observed in oxic waters under identical conditions. Bacteria responded similarly during anoxic incubations of cyanobacterial cells with over 6-fold higher bacterial biomass present at day 16 than introduced through incoming anoxic waters. In both anoxic incubations, bacterial abundance declined significantly after day 20, with a secondary peak observed in the cyanobacterial incubation at day 30. Bacterial abundance showed little change in the anoxic diatom incubation from day 70 onwards where numbers were similar to that present in incoming waters (7.2 x 105 ml). In anoxic cyanobacterial incubations bacteria declined from day 30 to day 153.

Using the directly counted bacteria and their calculated carbon content, a conservative estimate can be made of the bacterial carbon contribution to the total particulate organic carbon (POC) pool during decay (Fig. 2). Using this approach, bacterial carbon is considered as a percentage of the total particulate pool collected by filtration. Examination of filtrates after passage through the glass fiber (GF/F)

30

25

o

~ 0

Ox/c decay

0 15 30 45 6O 75

Time (Days)

"•30 Anoxie decay

~ 2o

O J , 0 30 60 90 120 150 180

Time (Days)

Fig. 2. The contribution of bacterial carbon as a percentage of the total particulate organic carbon oxic and anoxic decay of diatom (s) and cyanobacterial (C)) cells.

Note longer incubation time for anoxic incubations.

filters found that greater than 96% of bacteria were retained (Harvey, unpubl, data). The low numbers of bacteria which did pass through the filter were generally smaller in size. This passage was highest in number near the end of the experimental periods when overall bacterial abundance was reduced. In both oxic and anoxic decay series, the contribution of bacterial carbon as intact cells varied over time (Fig. 2). Such variations were due to the substantial changes in bacterial biomass during the degradation sequence as well as the decrease in particulate carbon derived from algal sources (Fig. 1). For oxic incubations of diatoms, bacterial carbon showed two peaks (Fig. 2). The first peak occurred during the time of greatest bacterial numbers (days 15 and 20) where they accounted for 17.2 and 21.4% of total particulate carbon. The second peak occurred during the later stages of the incubation after algal derived POC was greatly reduced, with bacterial carbon accounting for 25.2% of total POC at day 77. The bacterial contribution during cyanobaeterial decay under identical conditions showed a different pattern, with the contribution of bacterial cells increasing early in the incubation to 12.1%, slowly decreasing to day 20 and increasing at day 30 of 10.7% of POC (Fig. 2).


(D

50

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1 3 5 7 10 15 20 30 41 51 62 77 1 7 13 21 28 42 56 70 92 153 181

75

Anoxic

50

25

3 5 7 9 12 15 20 30 41 51 0 2 9 16 23 30 38 52 127 153

Time (Days) Time (Days)

Fig. 3. Changes in the relative abundance of fatty acid classes during oxic and anoxic decay of diatom (A and C) or cyanobacteria (B and D). In all incubations, odd and branched chain (odd and br) and hydroxy fatty acids (OH) attributed to bacteria increased with loss of polyunsaturated (PUFA) acids in

diatom incubations and shifts in monounsaturated (mono) and saturated (sat) acids.

During anoxic degradation experiments, bacterial carbon as a percentage of total POC displayed a somewhat different pattern (Fig. 2). Bacteria present in diatom incubations increased to a maximum for all incubations at day 13 (32.6% of POC) which coincided with the time of greatest abundance seen in the oxic diatom decay experiment. Bacterial biomass decreased thereafter, particularly after day 56 to the end of the experiment (Fig. 2). Bacterial biomass during anoxic cyanobacterial decay showed an overall similar pattern, increasing to a maximum of 21.6% of total POC at day 16 and slowly decreasing as the incubation progressed. Although total bacterial carbon was greater in both anoxic incubations compared to their oxic counterparts when incubations were halted (Fig. 1), the dilution by re- sidual algal organic matter retained under anoxic conditions resulted in a smaller contribution of bacterial to the total POC pool compared with oxic degradations.

Bacterial fatty acids during decay

The distribution of total fatty acids associated with particles during oxic and anoxic decay of the two phytoplankton showed marked changes over

time (Tables 1-4). Changes in concentration and abundance include the loss of algal fatty acids as well as shifts over time in the concentration and abundance of fatty acids derived from the microbial consortia. Degradation rates for algal lipids under oxic and anoxic conditions are presented elsewhere (Harvey and Macko, in press) and here we focus on fatty acids attributed to the microbial community over time. Bacterial fatty acids were defined as those absent from axenic cultures and which are documented to originate principally from bacteria (for thorough review see Ratledge and Wilkinson, 1989). They include the C I 2 and Cl3 saturated acids, odd and branched chain acids, especially the Cl5 and CI7 acids (Kaneda, 1991), the 2- and 3- hydroxy acids and several fatty acids containing unsaturation in the trans configuration such as the monounsaturated 16:lt and 18:lt acids. Although many fatty acids were present which are common to both algae and bacteria (e.g. Dunstan et al., 1994), the dominance of algal derived acids did not allow potential bacterial contributions to be assessed by concentration alone.

The total concentration of bacterial fatty acids during oxic diatom decay increased rapidly after

Catalys ts or con t r ibu tors

Table 1. Relative abundance of free and esterified fatty acids during oxic diatom decay*

537

Time ~omonse t ofincubation (days)

Fame 0 1 3 5t 7 10 15 20 30 41 51 62 77

13:0 0.0 0.0 0.0 0.0 0.1 0.0 0.1 0.5 0.0 0.0 0.0 0.0 0.0 12:2-2OH 0.0 0.0 0.0 0.0 0.0 0.1 1.0 1.7 1.0 2.2 0.0 0.0 3.3 14:1 0.4 0.4 0.3 0.2 0.3 0.1 0.2 0.0 0.0 0.0 0.0 0.0 0.0 14:0 13.6 15.9 23.3 15.7 16.7 15.3 9.3 12.3 13.8 11.8 6.7 2.9 17.7 13:0-OH 0.0 0.0 0.1 0.2 0.5 0.4 0.3 0.0 0.0 0.0 0.0 0.0 0.0 15:0i 0.2 0.3 0.7 0.7 1.1 1.3 10.4 16.5 7.2 11.1 7.0 2.7 4.6 15:0a 0.1 0.2 0.4 0.4 0.8 0.6 2.2 4.8 3.2 4.6 3.4 1.3 3.5 15:la 0.1 0.1 0.2 0.2 0.0 0.1 0.1 0.0 0.0 0.0 0.0 0.0 0.0 15:1 0.5 0.5 0.6 0.6 0.6 0.4 0.2 0.0 0.0 0.0 0.0 0.0 0.0 15:0 1.4 2.0 4.2 2.7 2.2 2.8 2.4 3.9 3.1 2.8 3.2 1.5 2.7 14:0-2OH 0.0 0.0 0.0 0.0 0.0 0.0 0.1 0.5 0.4 0.0 0.0 0.0 0.0 16:0-i 0.0 0.1 0.0 0.0 0.3 0.1 0.0 1.1 0.0 0.0 0.0 0.0 0.0 16:3 0.3 0.3 0.2 0.2 0.3 0.2 0.0 1.6 0.0 0.0 0.0 1.6 0.0 16:2 11.8 10.5 6.2 8.7 8.1 8.5 4.8 0.0 2.8 6.7 4.2 0.0 2.3 16:1(9) 30.2 27.5 21.0 25.5 31.1 25.7 19.6 0.2 1.2 0.0 0.0 0.0 4.0 16:1(11) 11.8 11.4 5.9 10.1 7.9 10.3 0.0 0.0 0.0 0.0 0.0 0.0 0.0 16 0.7 0.6 0.4 0.9 0.6 0.3 2.6 0.0 0.8 0.0 0.0 0.0 0.6 16:0 2.2 1.7 3.0 3.4 2.9 3.3 1.2 0.9 1.2 0.0 0.0 0.0 0.0 16:0n 13.5 14.4 19.9 17.8 14.4 21.2 20.5 38.4 40.9 32.6 42.0 32.0 42.6 15:0-OH 0.1 0.3 0.2 0.2 0.2 0.1 0.1 0.5 0.0 0.0 0.0 0.0 0.0 17:0i 0.0 0.0 0.1 0.2 0.2 0.2 0.4 2.3 2.9 4.2 3.4 2.1 1.5 17:0a 0.0 0.0 0.0 0.3 0.3 0.1 0.3 0.0 0.0 2.4 2.2 0.0 0.8 17:0cyc 0.0 0.0 0.0 0.0 0.1 0.0 0.2 0.0 0.0 0.6 0.0 0.0 0.7 17:0n 0.1 0.0 0.0 0.1 0.1 0.1 0.3 1.1 1.2 1.3 2.0 0.0 0.9 16:2-OH 0.0 0.0 0.2 0.0 0.1 0.1 0.1 0.0 0.0 0.0 0.0 2.1 0.0 18:4 1.1 1.2 0.4 0.8 1.3 0.8 0.0 0.0 0.0 0.0 0.0 0.0 0.0 18:3 0.2 0.3 0.3 0.1 0.3 0.2 0.8 0.0 0.0 0.0 0.0 0.0 0.0 18:2 0.5 0.5 0.8 0.4 0.5 0.5 0.9 1.2 1.2 0.0 0.0 0.0 0.0 18:1(9) 0.6 0.6 0.5 0.5 0.6 0.7 3.6 0.8 2.5 1.9 0.0 9.7 1.4 18:1(11) 0.7 0.7 0.4 1.2 1.1 1.1 7.5 0.9 3.7 0.0 0.0 10.8 1.7 18:0 0.2 0.4 0.5 0.5 0.5 0.8 2.4 7.2 6.5 5.2 13.2 24.2 5.2 Unknown 0.1 0.1 0.4 0.5 0.5 0.4 0.5 1.2 1.5 0.0 0.0 2.2 0.4 19:~yc 0.1 0.2 0.2 0.3 0.7 0.2 0.l 0.0 0.0 0.7 0.0 0.0 0.0 20:5 8.0 7.7 7.4 6.0 2.7 2.9 3.1 0.0 0.3 0.0 0.0 0.0 0.0 20:3 0.0 0.2 0.3 0.0 0.2 0.1 2.8 1.0 3.1 5.9 6.7 2.8 3.9 20:0 0.0 0.1 0.0 0.0 0.2 0.0 0.1 0.5 0.4 0.0 0.0 1.2 0.0 22:1 1.4 1.6 0.0 1.1 1.6 0.7 0.8 0.0 0.0 0.0 0.0 0.0 0.0 22:0 0.0 0.0 0.0 0.0 0.0 0.0 0.0 0.3 1.1 4.1 5.1 1.9 1.4 24:0 0.0 0.0 0.0 0.0 0.1 0.0 0.1 0.4 0.0 1.4 1.0 0.9 0.2

*Individual fatty acids which were < 0.2% of total acids at all time points are excluded for brevity. tTime of cell death as defined by point where algal protein synthesis ceased (see Harvey et al., 1995).

flow through the system was started, increasing over 4-fold during the first 10 days (Fig. 4A). Highest concentrations (164 #g l - l ) appeared during the 15-20-day period, which coincided with periods of highest bacterial carbon (Fig. 1), and decreasing rapidly thereafter. During the first 10 days of oxic decay of cyanobacterial cells, bacterial fatty acids showed a slow increase to 76 pg l - l , followed by a decline throughout the remainder of the experimental period (Fig. 4). Although total bacterial carbon over the initial 10 days of the incubation period was substantial higher in the oxic cyanobacterial incubation than with using diatom cells (Fig. 1), average bacterial acid concentrations were less (50 + 19 #g l-l), than for the incubation with atom cells as the substrate (70 + 60/~g l-l). The pattern of summed bacterial fatty acids during anoxic decay of diatom and cyanobacteria showed similar trends early in the incubation period (Fig. 4). Both showed rapid increases in concentration from the time of exposure to the microbial assembledge present in incoming waters, with almost 6-fold change over the first 20 days compared with initial values. The increases also corresponded to the increase in bac-

terial carbon early in the experimental period (Fig. 1). As the incubations progressed, concentrations of bacterial acids in anoxic diatom incubations decreased rapidly to 9.6 pg 1 - j , following a similar pattern to that observed for total bacterial carbon. In contrast, fatty acids diagnostic of bacteria during anoxic cyanobacterial decay declined from day 20 to day 60, but remained significantly higher (averaging 230 + 29 pg 1 - l over the final 90 days) than in all other incubations for the rest of the experimental period.

Isotopes of individual fatty acids

In a sub-set of samples from the oxic diatom decay experiments, carbon isotopic signatures of individual fatty acids representing algal, bacterial and mixed sources were determined over the time period of greatest bacterial abundance (Fig. 5). The acids examined included saturated 15:0 representing the major bacterial acid, oleic acid (18:l(nll)) of predo- minantly diatom origin together with other acids known to occur in both algae and bacteria including the saturated 14:0 and 16:0 acids and the monounsaturated 16:1. After initiation of flow, isotopic

538 H. R. Harvey and S. A. Macko

Table 2. Relative abundance of free and esterified fatty acids during anoxic diatom decay*

Time from onset o f incubation (days)

Fame 0 t 8 13 21 28 42 56 70 92 153 181

12:0 0.0 0.3 0.2 0.2 0.0 0.3 0.3 2.4 0.5 0.5 0.0 13:0 0.0 0.4 0.4 0.3 0.0 0.4 0,4 0.0 0.0 0.0 0.0 12:0-OH 0.0 0.2 0.4 0.3 0.0 0.3 0.3 4.2 0.0 5.6 0.0 14:1 0.2 0.3 0.3 0.2 0.0 0.2 0.4 0.0 0.0 0.0 0.0 14:0 8.1 14.7 16.9 13.4 11.5 15.6 15.5 8.2 8.8 5.0 6.4 15:0i 0.3 0.7 3.7 3.5 4.0 5.1 4.0 4.5 7,0 4.1 6.3 15:0a 0.2 0.3 1.3 1.5 1.7 2.4 2.0 1.8 2.6 1.6 2.4 14:1-OH 0.1 0.0 0.7 0.5 0.4 0.5 0.4 0.0 0.0 0.0 0.0 15:1 0.4 1.2 1.0 0.7 0.6 0.7 0.6 0.0 0.3 0.0 0.0 15:0n 1.0 5.8 5.2 4.1 4.3 4.6 4.0 5.5 5.1 2.9 4.2 16:3 0.4 0.3 0. I 0.0 0.0 0.0 0.0 0.0 0.0 0.0 0.0 16:2 15.4 13.2 6.9 5.0 4.1 3.6 3.1 3.2 0.5 0.0 0.0 16:1(9) 29.2 26.7 28.8 33.8 34.5 33.3 32.7 2.3 20.1 12.2 17.8 16:1(1 I) I 1.1 0.9 2.1 1.3 1.4 1.2 2.2 6.0 1.1 0.6 0.0 16:0 13.0 18.1 20.8 18.3 22.3 19.8 19.7 37.1 29.2 17.5 29.5 15:0-OH 0.0 0.0 0.4 0.3 0.3 0.2 0.2 3. I 0.4 6.5 0.0 Unknown 0.0 0.0 0.0 0.0 0.0 0.0 0.0 1. I 1.8 1.3 2.1 Unknown 0.0 0.0 0.0 0.0 0.0 0.0 0.0 2.4 0.0 5.6 0.0 17:0i 0.0 0.3 0.2 0.1 0.0 0.0 0.0 3.0 0.6 5.5 0.6 17:0a 0.0 0.1 0.2 0.2 0.0 0.3 0.4 0.0 0.8 0.6 1.1 17:0n 0.0 0.1 0.2 0.2 0.0 0.0 0.2 2.9 0.7 5.0 0.8 Unknown 0.3 0.2 0.0 0.0 0.0 0.0 0.0 0.0 0.0 0.0 0.0 18:3 1.4 1.1 0.4 0.4 0.4 0.2 0.2 0.0 0.0 0.0 0.0 18:2 0.7 0.3 0.0 0.0 0.0 0.0 0.0 0.0 0.0 0.4 0.0 18:2 0.5 0.5 0.1 0. I 0.0 0.0 0.0 1.5 I).0 0.0 0.0 18:1(9) 0,6 0.6 0.6 2.1 0.7 0.3 0.4 1.6 3.0 3.5 4.9 18:1(11) 0.5 0.9 3.8 6.6 8.6 7.4 9.2 3.5 11.1 7.4 13.0 18:0 0.3 0.4 0.4 0.5 0.6 0.3 0.5 1.7 2.5 2,0 3.9 U n k n o w n - O H 0.1 0.1 0.2 0.3 0.3 0.3 0,3 0.0 1.0 0.7 1.3 Unknown 0.0 0.4 0.1 0.2 0.0 0.0 0.0 0.0 0.4 0.9 1.9 20:5 13.7 10.3 3.3 4.7 3.6 2.3 2,5 1.0 1.4 1.7 2.2 Unknown 0.0 0.0 0.0 0.0 0.0 0.0 0.0 2.0 0.0 5.4 0.0 20:0 0.0 0.0 0.6 0.3 0.0 0.0 0.0 0.0 0.0 0,0 0.0 22:1 2.6 1.5 0.6 1.0 0.7 0,5 0.5 0.0 0.8 0,8 0.9 22:0 0.0 0.0 0.0 0.0 0.0 0.0 0.0 1.0 0.5 2.6 0.7

*Individual fatty acids which were < 0.2% of total acids at all t ime points are excluded for brevity. t T i m e of cell death as defined by point where algal protein synthesis ceased (see Harvey et al., 1995).

signatures among the fatty acids ranged from 18.4%o from bacterial sources (15:0) to -22.2%0 (18:1 and 14:0) from algae (Fig. 5). During the incubation period all five acids showed changes in isotopic composition, displaying a small enrichment (average of + 1.6%o) in 13C after the period of major POC loss. Myristic acid (14:0) was the only fatty acid to show a depletion over the first 15 days, shifting 3.2%o whereas other acids either showed little change (e.g. 15:0) or a shift to more enriched values as degradation proceeded (Fig. 5). Oleic acid shows the largest change at any one time with a 4.2%o enrichment over the first 15 days of degradation, returning to more depleted values as the incubations progressed.

D I S C U S S I O N

As these laboratory results support, bacteria are key catalytic agents during the early diagenesis of phytoplankton derived organic matter in the water column. They show a strong correspondence with the pattern of degradation for all but oxic cyanobacterial decay, with greatest abundance during times of most rapid POC loss (Fig. I). Measures of metabolic activity conducted in parallel also support a tight linkage, with observations of elevated

uptake rates during greatest POC loss and much reduced rates in incubations after the bulk of organic matter is removed (Harvey et al., 1995). As a contributor of carbon to detrital organic matter, however, bacteria appear to play a variable role which is dependent on the source material and the fraction which is preserved (Fig. 2). The contribution of bacterial biomass to POC during oxic and anoxic diatom degradation sequences peaked during the early portion of the incubation period when POC loss was maximal, with turnover times for POC equal to 18.6 and 68.4 days for oxic and anoxic, respectively. A second increase in the bacterial biomass contribution appeared late in oxic diatom decay, but only when little ( < 4% of initial) POC remained. This increase was unexpected given the presumed low amounts of labile carbon remaining and appears an artifact of the flow-through system used, which continually introduces significant bacterial numbers in incoming water. Our earlier results found that bacterial community metabolism of major soluble organic pools (dissolved amino acids and monosaccharides) corresponded with changes in bacterial numbers over time (Harvey et al., 1995). More importantly, metabolism of both pools declined in the later stages of all incubations. During these continual flow experiments, bacterial

Table 3. Relative abundance of free and esterified fatty acids during oxic cyanobacterial decay*

Time from onset of incubation (days)

Fame 0 1 3 5 7 9t 12 15 20 30 41 51

13:0 0.0 0.0 0.1 0.1 0.0 0.1 0.0 1.7 1.2 1.7 1.4 1.7 12:0-OH 0.1 2.6 1.3 1.3 1.5 1.6 0.4 0.0 0.2 0.0 0.0 0.0 14:0 49.2 33.3 33.5 30.0 26.6 28.6 19.8 25.3 11.3 6.6 5.9 5.6 13:0-OH 0.0 0.0 0.1 0.1 0.1 0.3 0.5 0.0 2.0 0.6 0.0 0.4 15:0i 0.8 1.2 2.6 2.9 3.0 3.6 2.7 9.6 9.2 8.1 11.5 9.3 15:0a 0.2 0.2 0.4 0.4 0.3 0.5 0.6 8.8 4.7 4.2 4.8 4.2 15:1 0.0 0.6 0.3 0.3 0.4 0.4 0.2 0.0 0.3 0.8 1.6 2.0 15:0n 1.4 1.2 1.3 1.4 1.2 1.3 2.0 3.7 2.3 2.3 2.7 2.1 14:0-2OH 0.0 0.0 0.2 0.3 0.2 0.3 0.3 1.0 2.2 2.1 3.5 2.6 16:1(9) 1.3 35.3 24.0 24.7 31.2 30.9 20.9 0.0 14.2 2.5 2.7 14.8 16:0i 0.0 0.7 0.4 0.5 0.6 0.7 0.4 0.0 0.4 0.0 0.4 1.3 16:0 16.7 13.8 15.3 15.2 13.8 13.0 19.4 21.7 13.8 23.7 28.5 22.7 Unknown 0.0 0.0 0.0 1.6 1.0 0.2 0.3 1.9 0.7 1.0 0.0 0.3 17:0i 0.1 0.3 0.4 0.8 1.0 1.0 0.5 0.0 0.5 0.0 0.0 0.3 Unknown 0.0 0.0 0.0 0.0 0.1 0.1 0.0 0.2 0.2 0.5 1.0 0.7 Unknown 0.0 0.0 0.1 0.1 0.2 0.4 1.1 0.0 0.1 0.0 0.0 0.0 17:0a 0.1 0.1 0.4 0.5 0.7 0.9 0.9 1.0 1.1 1.2 2.4 1.7 UnknownOH 0.0 0.0 0.1 0.1 0.1 0.2 0.5 0.6 2.0 2.0 2.4 1.8 17:1 0.3 0.2 0.4 0.5 0.3 0.4 1.3 1.9 1.1 1.2 1.7 1.0 17:0 2.6 0.2 0.6 0.4 0.1 0.1 0.2 0.7 0.0 0.0 0.0 0.0 16:0-OH 0.4 0.1 0.2 0.1 0.2 0.2 1.9 0.4 0.9 0.6 0.3 0.5 18:1(9) 0.1 0.3 0.5 0.6 0.7 0.7 2.9 0.4 2.7 8.2 1.8 3.0 18:1(11) 0.2 5.1 5.6 5.6 8.4 4.0 5.3 0.0 12.1 11.3 4.4 9.5 18:0 0.4 0.3 0.4 0.9 0.8 0.6 2.5 3.6 2.9 7.4 8.1 3.4 17:0-OH 0.2 1.3 1.8 2.0 2.2 1.1 0.7 0.0 2.6 1.5 3.6 3.5 UnknownOH 0.8 0.1 0.1 0.2 0.1 0.2 0.3 0.4 0.5 0.4 1.0 0.8 UnknownOH 0.0 0.0 0.0 0.2 0.2 0.1 0.0 0.8 0,4 1.8 0.2 0.1 19:0cyc 0.1 0.0 0.0 0.0 0.1 0.1 0.0 1.2 0.6 1.3 0.8 1.0 20:1 19.0 1.3 4.7 3.9 0.7 0.5 0.3 0.2 1.0 0.4 0.7 0.3 20:0 2.5 0.4 1.5 0.9 0.8 1.8 5.0 8.0 0.4 1.9 1.2 0.3 Unknown OH 0.7 0.6 1.4 1.2 1.3 2.0 1.7 0.2 0.7 0.7 1.4 0.7 Unknown br 0.0 0.0 0.0 0.1 0.0 0.1 0.8 0.8 0.5 0.6 1.2 0.6 Unknown-OH 0.4 0.4 1.0 0.9 0.8 1.5 1.2 0.1 0.5 0.4 1.0 0.6 22:u 1.8 0.1 0.6 0.6 0.0 0.0 0.0 0.3 0.3 0.5 0.0 0.4 22:1 0.2 0.0 0.2 0.1 0.2 0.2 0.8 2.4 0.3 2.2 1.1 0.2 22:0 0.0 0.0 0.0 0.4 0.0 0.1 0.3 0.8 0.5 0.6 0.5 0.0 21:0-OH 0.1 0.0 0.2 0.2 0.1 0.2 0.3 0.3 0.4 1.0 1.2 0.1 24:0 0.0 0.0 0.0 0.0 0.0 0.0 0.2 0.2 0.1 0.0 0.3 0.2

*Individual fatty acids which were <0.3% of total acids at all times are excluded for brevity.

Table 4. Relative abundance of free and esterified fatty acids during anoxic cyanobacterial decay*

Time from onset of incubation (days)

ID 0t 2 9 16 23 30 38 52 127 153

14:1 5.1 6.3 1.9 1.0 0.3 0.0 0.0 0.0 0,0 0.0 14:0 39.8 52.7 38.8 27.4 25.8 28.9 31.8 26.0 18,2 18.9 15:0i 0.3 0.7 0.5 1.7 2.7 4.0 5.3 8.3 10.2 11.3 15:0a 0.1 0.3 0.3 0.8 1.2 1.7 2.2 3.0 3,7 3.6 15:1 1.2 0.0 0.6 0.5 0.2 0.0 0.0 0.0 0.1 0.0 15:0 1.3 2.5 1.5 1.6 1.8 2.3 2.6 2.6 3.3 4.0 14:0-OH 0.0 0.0 0.0 0.0 0.0 0.0 0.0 0.0 0.2 0.3 16:0br 0.0 0.0 0.0 0.1 0.0 0.2 0.2 0.3 0.5 0.6 16:1 (9) 1.3 1.8 2.0 1.2 0.5 0.3 0.4 0.6 0.6 0.2 16:0 9.4 19.7 13.3 19.1 25.5 30.4 34.0 30.7 29.6 32.1 17:0br 0.0 0.0 0.0 0.0 0.0 0.0 0.0 0.3 1.8 2.4 15:0-OH 0.1 0.0 0.1 0.2 0.3 0.3 0.4 0.5 0.8 0.9 17:0a 0.0 0.0 0.1 0.1 0.1 0.2 0.2 0.4 0.6 0.7 17:1 0.1 0.0 0.2 0.2 0.2 0.0 0.2 0.0 0.3 0.3 17:0n 0.2 0.3 0.2 0.3 0.4 0.5 0.6 0.6 0.8 0.9 Unknown 0.0 0.0 0.0 0.0 0,5 1.0 0.0 0.0 0.0 0.0 16:0-OH 0.3 2.2 0.1 0.2 0.3 0.0 1.3 1.9 3.8 3.2 20:br 2.0 0.4 5.3 8.9 6.2 3.7 0.0 0.3 2.2 0.3 18:1 (9) 0.0 0.0 0.0 0.2 0.2 0.2 0.2 0.2 0.2 0.0 18:0 0.1 0.3 0.2 0.4 0.6 0.9 1.1 1.5 1.8 1.9 17:0-OH 0.5 0.2 0.5 0.9 0.7 0.6 0.0 0.0 0.5 0.1 18:0-2OH 0.1 0.7 0.0 0.0 0.0 0.0 0.3 0.0 0.0 0.0 Unknown OH 0.0 0.9 0.0 0.0 0,3 0.3 0.0 0.0 0.2 0.3 Unknown OH 0.2 0.2 0.1 0.4 0.5 0.6 0.1 0.2 1.0 0.8 18:0-OH 0.0 0.4 0.0 0.0 0.2 0.0 1.2 0.0 0.3 0.3 20:0 2. I 1.9 0,4 0.3 8.2 9.1 2.0 0.9 4.8 5.8 Unknown 0.1 0.9 0.0 0.0 0.3 0.3 0.0 0.0 0.2 0.3 Unknown OH 0.4 0.6 0.3 0.5 1.8 1.9 4.2 5.6 1.9 2.4 20:br 0.0 0.0 0.0 0.0 0.0 0.0 0.2 0.3 0.2 0.0 22:1 0.0 0.7 0.0 0.0 0.2 0.2 0,4 0.6 0.4 0.4 22:0 0.4 0.8 0.6 0.7 2.3 1.9 6.9 10.7 3.3 3.4 Unknown oh 0,0 0.0 0.0 0.0 0.0 0,0 0.5 0.0 0,1 0.0 Unknown sat 0.0 0.0 0.0 0.1 3,2 3.6 0.8 0.3 1.6 2.6 22:0-OH 0.0 0.0 0.0 0.1 0.5 0.4 1.7 3.2 0.6 0.5

*Individual fatty acids which were <0.3% of total acids at all times are excluded for brevity. tTime of cell death (Harvey et al., 1995).

540

180

160

140

120 I00 SO ~ 60

.~ 4o ' ~ 20

o

,,0

r~

400

350

300

250

200

150

100

50

0

torn Oxic decay

cyano 5 ~- --_~ ---~---_ o

0 20 40 60

o/~ Anoxic decay

----0-..%

0 20 40 60 80 100 120 140 160 180 200

Time (Days) Fig. 4. The concentration of total bacterial fatty acids over the course of oxic and anoxic incubations of either diatom (e) or cyanobacterial (O). In all incubations fatty acids attributed to bacteria peaked early during incubations. Note longer incubation times for anoxic exper-

iments.

biomass in the late stages of oxic diatom decay approaches that of incoming waters (Fig. 1), and the collection of particles by filtration removed suspended bacteria as well as those associated with the small number of detrital particles present. If bacterial biomass present in incoming waters

H. R. Harvey and S. A. Macko

( l . 6 x 106ml - l ) is subtracted from the total bacterial carbon based upon the volume filtered, the contribution of bacterial carbon to POC is reduced from 25.1 to 9.2% of the total. In other incubations, bacterial carbon present at the conclusion of incubations as observed cells accounted for less than 7% of total organic carbon and would be further reduced if bacteria suspended in incoming waters were excluded.

Although several fatty acids have been assigned to specific bacterial groups or even individual bac- 80 teria (e.g. Ratledge and Wilkinson, 1989; Kaneda, 1991), previous attempts to link specific fatty acids with bacteria in natural communities have often been equivocal. Taylor and Parkes (1985) used additions of either acetate, propionate, lactate or hydrogen to anoxic marine sediment slurries together with fatty acid profiling to examine changes in the bacterial community. They followed the changes in fatty acid distribution in slurries over time compared to fatty acids of pure cultures of sulfate reducing bacteria. Additions of acetate, considered a primary substrate for sulfate reduction, to slurries resulted in no significant changes in fatty acids concentrations attributed to sulfate reducing bacteria. Propionate and L-lactate additions, however, led to elevated concentrations of the 17:1 (co6) fatty acid considered diagnostic of the presence of Desulfobulbus sp. in the slurries, yet total biomass showed no significant increase. We also observed the (~)6 17:1 acid in trace amounts during the degradative process in anoxic waters, but this acid was also seen in oxic incubations where sulfate reduction did not occur. While a viable population of sulfate reducers seems a reasonable source for this acid in anoxic environments, its presence in oxic environments

-26

~ -24

-22 .o

• ~- -20

~ -18

-16

Oxic diatom decay

0 10 20 30 40

- -o - - 14:0 - . © . 15:0

16:1 ---~.. 16:0

18:1

T ime (Days)

Fig. 5. The 3~3C isotopic signatures of important fatty acids present during the active stage of oxic diatom decay. All fatty acids showed significant shifts during the incubation period with an average 1.6%

enrichment at day 42.


suggests that it may not be restricted to this group of microorganisms alone. Instead, aerobic or facultative bacteria growing under oxic conditions must also contain this acid in significant quantities, albeit in much lower amounts than observed for Desulfobulbus.

One difficulty in documenting sources for individual bacterial fatty acids is a likely consequence of the small sub-set of bacteria whose lipid distribution has been established. It is generally believed that at most about 1% of the viable bacterial community in environmental samples can be successfully cultured, with even lower success (0.001-0.1%) for bacteria in seawater cultures (see Amann et al., 1995 for review). As a result, our ability to consider fatty acids as specific markers is limited by a pau- city of data on the distribution of fatty acids among the true bacterial consortia and reliance on those that can be successfully cultured. Although several techniques have been developed for match- ing fatty acid distributions with likely bacterial populations in soils and sediments (e.g. Findlay and Dobbs, t993) or phospholipids as surrogate measures of microbial biomass (Findlay et al., 1989) they are ultimately limited by the ability to culture the great diversity of bacteria which inhabit environmental samples.

To evaluate the relationship of bacterial fatty acids and bacterial carbon without assignment to specific groups, the concentration of bacterial acids at any time point during oxic and anoxic degradation experiments can be plotted vs bacterial carbon derived from direct counts of intact bacterial cells. Because bacterial fatty acids represent conservative membrane components (largely as phospholipids) and not storage products, a simple relationship with bacterial biomass should be seen. For oxic incubations during diatom or cyanobacteria decay, no evidence for a direct relationship is seen (Fig. 6, left panels) even though higher amounts of bacterial carbon (Fig. 1) often occur at times of elevated bacterial fatty acids (e.g. day 15 and 20 in Fig. 4). Although at first examination bacterial carbon and identified bacterial fatty acids under anoxic conditions appear coincident over time, again a relationship based upon concentration is not seen (Fig. 6, right panels). Anoxic decomposition of diatoms showed a significant relationship for a simple regression relationship only if the time point of greatest bacterial fatty acid (day 8 -368 #g l - l) is excluded. The fact that the point in time of greatest bacterial fatty acids does not co- incide with the time of greatest bacterial carbon is troublesome, and brings into question that such a

.-t g

CO

1500

1200

9O0

6OO

300

0

1200

1000

800

600

40O

200

0

~ c ~ y Anoxic decay

l I 0 30 60 90 120 150 180 200 300 400

S

• •

I'5 30

3500 . diatom 3000.

2500 .

2000

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1000 • Q

5 0 0

o ••% I 0 100

5OOO

I cyanobacteria

1000

cyauobacteria

0~ o 45 ~ 7'5 o zoo 2oo 3oo ~o

Summed "baet~al" fatty acids (~tg)

Fig. 6. The concentration of total bacterial carbon (/~g) vs the summed concentration of bacterial fatty acids at all time points during oxic and anoxic decay of diatom (o) or cyanobacterial (0) cells. Note

different scales for bacterial fatty acids.


simple relationship based on these parameters exist. A treatment using bacterial abundance alone saw similar trends (data not shown).

At least two sources for error can explain the lack of agreement between observed fatty acid concentrations and estimates of bacterial carbon and these are not mutually exclusive. The first is that the bacterial consortia contains fatty acids in addition to those considered strictly bacterial, i.e. those in common with both terrestrial and auto- chthonous organic sources which are not unique. Both algal (e.g. Viso and Marty, 1993; Dunstan et al. , 1994) and terrigenous (Cranwell, 1991; Simoneit et al. , 1991) sources of organic material commonly contain fatty acids which overlap with those known to occur in bacteria (Goossens et al. , 1989), especially among the C16 and C~8 fatty acids. Bacteria isolated from natural sedimentary environments commonly contain a significant, and often major, portion of their fatty acids as the saturated 16:0 and 18:0 acids (e.g. Perry et al. , 1979; Parkes and Taylor, 1983) even though "bacterial" fatty acids occur. This leads to an under-estimate of the contribution of bacterial fatty acids compared to the observed biomass, since a fraction of bacterial fatty acids are excluded during summation.

To illustrate the difficulty in assigning specific fatty acids to source material, fatty acids considered unique to diatoms can be considered in an analo- gous way as those derived from bacteria. Using the concentration of polyunsaturated acids present in particles during diatom incubations, an estimate of the minimum contribution of algal carbon to the total POC pool can be obtained. The abundance of these acids decreases dramatically during the degradative process under both oxic and anoxic conditions (Fig. 3), with calculated first-order rate constants of 34.3 y-i and 13.6 y-l, respectively. These rates are much higher than values for the POC pool as a whole (12.8 y-1 and 2.9 y-i for oxic and anoxic respectively--Harvey et al. , 1995), but are useful to demonstrate the potential flaws in assigning any specific compound to source material in degraded material. During oxic diatom decomposition, the concentration of polyunsaturated fatty acids which persist would allow only 3.8% of the POC pool to be considered algal in origin. During anoxic diatom decay, degradative rates of POC and polyunsaturated acids differ even more, and as a result < 1% of remaining particulate carbon would be assigned as algal. In contrast, if fatty acids common in both algae and bacteria were included in the estimate of either algal or bacterial contributions to POC, the proportions would be substan- tially greater since these acids account for the largest fraction of total fatty acids in both groups (Tables 1-4).

A second consideration is the presence of bacterial fatty acids which do not represent bacteria

themselves, but secondary consumers. The demon- stration of the ability of marine ciliates to incorpor- ate fatty acids from the bacteria they consume (Ederington et al., 1995) allow "bacterial" fatty acids to occur in the absence of bacteria. Although the activities of microzooplankton were not quantified in the present study, tetrahymanol, a triterpenoid alcohol which appears ubiquitous in bacterivorus ciliates (Harvey and McManus, 1991), was observed during oxic incubations of both phytoplankton, suggesting that bacterial predation by ciliates did occur. Bacterial consumption would tend to over-estimate the presence of viable bacterial populations as diagnostic fatty acids of bacteria remain as membrane components of microzooplankton long after digestion of the bacterial cell.

In either of these scenarios, bacterial fatty acids and measures of bacterial biomass are uncoupled, and could account for the absence of obvious agreement seen in Fig. 6. Several other possibilities also exist, and have been discussed in previous reports including changes in microbial community structure during manipulation (e.g. Parkes and Taylor, 1983), and variability in bacterial fatty acid content or retention of bacteria detritus (Henrichs and Doyle, 1986; Peulve et al., 1996). The examination of isotopic signatures from major fatty acids during oxic diatom decay suggests that both bacterial and algal fatty acids undergo isotopic shifts during the degradative process (Fig. 5). Although the 15:0 acid began and concluded as the least depleted acid during the incubation, it showed an isotopic shift comparable to other fatty acids during the incubations. If the 15:0 fatty acid is solely bacterial, its varying signal indicates that the isotope changes in the more common fatty acids are a combination of new bacterial biomass in addition to remaining algal fatty acids. A similar shift to more enriched values is observed for the total lipid pool (data not shown), suggesting that fatty acids are selectively lost during the decomposition process and at least a fraction resynthesized into bacterial biomass. Other possibilities for enrichment could involve intramole- cular fractionation and loss of particular molecular groups such as carboxyls (Abelson and Hoering, 1961).

As these incubations demonstrate, bacteria are important catalytic agents during early diagenesis, but show large variations as contributors to the pool of organic carbon as intact cells. At times bacterial biomass can be a substantial fraction of the total particulate carbon pool, particularly during periods where microbial activity is greatest (Figs 1 and 2). As degradation progresses and labile substrates are depleted, bacterial biomass as evident from intact cells declines, with the overall contribution of bacteria dependent upon the pool of rem- nant organic matter which is retained. Fatty acids


which can be directly a t t r ibuted to bacter ia appear as a significant fract ion of fatty acids dur ing diagenesis, but show little re lat ionship with more direct measures o f bacterial biomass. As a result, the use of organic markers such as fatty acids to trace bacterial con t r ibu t ions remain a quali tat ive tool for the geochemical communi ty until fat ty acids present f rom diverse sources of organic mat te r can be assigned to their respective sources with some certainty.

Acknowledgements--We thank B. Millsaps for culturing the phytoplankton, T. Bell for technical assistance and J. Tuttle for stimulating discussions. Helpful comments by John Parkes and one anonymous reviewer significantly improved the final manuscript. This work was supported by the NSF Marine Chemistry Program (OCE 9203241) and by the Donors to the Petroleum Research Fund of the American Chemical Society. We also thank Brenda Yates for manuscript preparation. Contribution No. 2806, Center for Environmental Science of the University of Maryland System.

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