Bacterial and phosphorus dynamics in profundal Lake Erken ...

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Hydrobiologia 364: 55–63, 1998.

H. L. Golterman (ed.), Sediment-Water Interaction 8. c 1998 Kluwer Academic Publishers. Printed in Belgium.

Bacterial and phosphorus dynamics in profundal Lake Erken sediments following the deposition of diatoms: a laboratory study Erik Törnblom & Emil Rydin Institute of Limnology, Uppsala University, Norbyvägen 20, 752 36 Uppsala, Sweden Received 5 November 1996; accepted in revised form 4 December 1997

Key words: bacterial activity, bacterial biomass, phosphorus fractionations

Abstract The benthic microbial response to the deposition of natural seston and the microbial impact on nutrient dynamics was studied in an experimental system using whole sediment cores equipped with flow-through systems for the overlying water. For 20 days, changes in sediment bacterial activity, total metabolic activity (heat production), bacterial biomass, phosphorus fractions and basic chemistry were followed, as well as the exchange of nutrients between sediment and water. Microbial activity and biomass increased immediately in response to the deposition of seston, peaked after seven days and then decreased linearly over the remaining time of the experiment. Cosettled bacteria were suggested to play an important role in the microbial response. Changes in bacterial biomass production, bacterial biomass and the NaOH-nrP extractable phosphorus fraction were concurrent in response to seston additions. The sediment acted as a trap for SRP from the overlying water when bacterial activity was high and as a source when the bacterial activity decreased. Altogether, the results suggest an important role of bacteria in the regeneration of seston P. Mineralization rates estimated from sediment heat production showed that ca. 11% of the added seston carbon was oxidized in the sediments during the experiment. Introduction Bacterial degradation and transformation of particulate and dissolved organic matter are key processes determining turnover rates of organic and inorganic substances in aquatic systems (Fenchel & Blackburn, 1979. In sediments from both marine and freshwater environments, bacterial productivity has been shown to be most closely linked to substrate supply and temperature (Cole et al., 1988; Sander & Kalff, 1993). In regions with a marked seasonality, pulses of organic matter input to the sediments occurring during spring and autumn as settling diatoms blooms, can therefore be expected to be of major importance for microbial sediment communities. Studies of pelagic-benthic coupling in marine environments have shown that benthic sediment communities quickly can respond to inputs of fresh organic matter in terms of: increasing microbial activity (Meyer-Reil, 1983; Graf et al., 1982), increasing O2 consumption causing a shift towards anaerobic conditions (Kelly & Nixon, 1984; Graf, 1987; van

Duyl et al., 1992) and a release of nutrients from the sediments to the overlying water (Garber, 1984; Kelly & Nixon, 1984; Enoksson, 1993). Few studies performed in freshwater have focused on the utilization of the organic matter by benthic organisms (Fitzgerald & Gardner, 1993; Goedkoop & Johnson, 1996) and little is known how the normally observed increase in bacterial activity and biomass in response to the deposition of organic matter, is related to fluxes of nutrients and especially the P-turnover. The mobilization/immobilization of phosphate in lake sediments has traditionally been considered to be governed mainly by redox-controlled interactions with Fe and Mn. The activity of microorganisms has, since a long time been known to indirectly affect P-cycling through the impact of mineralization processes on redox conditions (Einsele, 1936; Mortimer, 1941, 1942). More recent studies have suggested that uptake, storage and release of P by microorganisms in surface sediments can be of major importance in P-cycling in lakes (cf. Boström et al., 1988; Gächter et al., 1988; Gächter & Meyer, 1993). The amount

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56 of P in bacteria has been shown to be large enough to explain most of the P-release observed from some lake sediments (Boström, 1988). P may be bound or released by sediment bacteria through metabolic reactions, extracellular release and lysis of bacterial cells. Bacterial P-dynamics in sediments are dependent on factors like: type, amount and quality of organic matter, P-availability in the surrounding water, NO3 - concentration, P and redox (Boström et al., 1988). In oligotrophic and mesotrophic lakes, the P-content of settling seston alone is too low to fulfil the high P- requirements of the mineralizing bacteria (Gächter & Meyer, 1993). Hence, decomposing bacteria in these systems may use o-P from the surrounding water (Gächter & Mares, 1985). In oxidized surface sediments, possible P-sources for bacteria include o-P from the overlying water, P bound to sediment organic matter or inorganic compounds in the sediment (Boström, 1988; Istvánovics, 1993). This paper reports the results of a laboratory experiment using intact sediment cores collected in the profundal of Lake Erken. The study was performed in order to study the short-term effects of the deposition of natural seston on the microbial sediment community and the role of sediment bacteria in P-regeneration.

Materials and methods Sediment and seston collection, experimental setup During the autumn diatom bloom 1994, sediment cores were collected from a boat at 13 1 m depth in the profundal zone of the eastern basin Lake Erken, Sweden. Lake Erken, situated approximately 65 km north of Stockholm at 59˚ 250 N, 18˚ 150 E, has a surface area of 23.7 km2 , a mean depth of 9 m, a maximum depth of 21 m and can be characterized as mesoeutrophic (Blomqvist et al., 1994). 52 sediment cores (inner diameter 64 mm, length 500 mm) were collected using a plexiglass core sampler. The surface sediments (0–0.5 cm) were well oxidized upon collection with redox values well above + 300 mV. The water temperature at 13 m depth was 12.5 ˚C. The top rubber stoppers were removed and cores were placed in a dark water bath regulates at in situ lake temperature using lake water continuously pumped from 4 m depth. Magnetic stirrers (speed 50 rpm) were placed in the water 8 cm above the sediments. The cores were also equipped with a flow-through system operated by peristatic pumps connected to a 50 l container with

aerated lake water (daily collected at 4 m depth and filtered through phytoplankton nets (mesh size 20 m)). The renewal time of the water overlying the sediments (450–600 ml) was approximately 20 h. Cores were left to equilibrate to laboratory conditions for 48 h. Seston was collected at 4 m depth using photoplankton nets (mesh size 20 m), rinsed with lake water and again concentrated with phytoplankton nets. Subsamples were taken for the determination of bacterial abundance and production, C-, N-, P- and chlorophyll a concentrations. Live subsamples were also investigated microscopically in order to quantify the proportion of living diatoms. Before seston additions, peristaltic pumps and magnetic stirrers were stopped. 20 ml of the concentrated seston suspension was carefully pipetted into the water close to the sediment surface in 24 randomly chosen cores. The added seston immediately formed a flocculent green layer on the sediment surface. After 5 h this layer had settled into a thin 3–4 mm) and smooth-surfaced carpet. Water circulation was restarted and samples of in-and all outflowing water were collected for chemical analyses and to enable the determination of the exact flux in each core. The stirring was not restarted until day 1 in order to prevent resuspension of the added seston. When the stirring was restarted and during the experiment no visible resuspension of seston or sediment occurred. On day 0, four replicate cores (controls) and on days 1, 2, 4, 7, 12 and 20, four replicate seston and control cores were randomly chosen for analyses of microbial activity and biomass, sediment chemistry and Pfractionations (day 1 no P-fractionations). Before sectioning, the O2 -concentration in the overlying water and sediment redox potentials were determined in all cores. The overlying water in each core was then carefully removed using a syringe and the sediment was sectioned into 0–0.5, 0.5–2 and 2–4 cm layers. When sectioning cores with seston additions care was taken to include both the seston carpet and the underlying 0.5 cm of sediment. The experimental temperature followed changes in in situ lake temperature at 14 m depth but was on average 2 ˚C lower due to cooling in the ground-based part of the water pipe between the lake and the laboratory. The first 7 days, the experimental temperature varied around 9 ˚C ( 0.2 ˚C) and then decreased over time to 6.5 ˚C on day 20. Analyses Sediment redox potentials were measured using a platinum Eh-electrode (P 10401, Radiometer A/S) in con-

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57 junction with a calomel reference electrode (K 401, Radiometer A/S). Measurements were performed at the sediment surface and at 0.5, 1, 1.5, 2, 3, 4 and 5 cm depth. Water samples were collected on days 0, 1, 2, 4, 7, 12, 15 and 20 from the experimental systems chosen for sediment analyses (2 replicates of inflowing and 4 for outflowing water). Heat production measurements were performed on subsamples from the 0–0.5 cm layer using a fourchannel heat conduction multichannel microcalorimeter (Thermal Activity Monitor 2277, Thermometric). Sample treatment and measurements were performed according to Boström and Törnblom (1990). All heat production measurements were performed at 12 ˚C. Heat production at in situ temperature was estimated assuming a Q10 -value of 3 according to Törnblom (1996). Heat production was converted to carbon units assuming RQ = 1 and using the aerobic catabolism of glucose as a model process assuming k H˚O2 = 469 kJ mol 1 O2 (Gnaiger & Kemp, 1990). Incubations of sediment samples with [3H]thymidine were performed at the ambient water temperature, in duplicates plus additional formaldehyde killed controls, according to Bell and Ahlgren (1987). 0.2 g of sediment and 0.2 ml seston suspension (only on day 0) was incubated in darkness was 20 Ci of [3 H]thymidine (45–55 Ci mmol 1 , Amersham) for 30 min. Additional [3 H]thymidine-incubations were performed at 9 ˚C on days 12 and 20 due to the temperature decrease during the last week of the experiment. The degree of participation of [3 H]thymidine was determined from isotope dilution experiments performed on day 4 according to Pollard and Moriarty 1984. An average degree of participation of 46 5% was found in both the seston treatment and the control. Bacterial production was calculated using a theoretical conversion factor of 0.44 1018 cells mol 1 assuming a DNA content of 2.5 fg per cell (Fuhrman and Azam, 1980) and corrected for the average degree of participation. The conversion of [3 H]thymidine incorporation rates into carbon units was made by applying the factor 2.2 10 13 g C m 3 (Bratbak & Dundas, 1984). Bacterial numbers and biomass were determined on sediment and seston samples preserved with filtersterilized (0.2 m) formaldehyde (final concentration 4%). Epifluorescense microscopy (Nikon filters Combination B-2A [Dichroic mirror 510, Excitation filter 450–490 and Barrier filter 520]) was used after sonication in an ice bath at 100 W for 1 min, dilution (final concentration of 0.005%), staining with acridine orange (Merck) and filtration onto 0.2 m prestained

(Sudan black (Sigma)) Nucleopore filters. 400 cells in at least 20 random fields were counted. Average bacterial volumes were calculated from size measurements by eyepiece graticule of a least 100 cells from each sample. Bacterial biomass was converted to carbon units by applying the factor 2.2 10 13 g m 3 of C (Bratbak & Dundas, 1984). Chlorophyll a in sediments and seston was analysed spectrophotometrically after freezing, freeze-drying and extraction with ethanol according to Hansson (1988). Sediment water content was determined after drying at 105 ˚C for 24 h. Sediment densities used in transformations into volume units were estimated using the equation: sediment density = 1.6174 0.0062 sediment water content (established for surface sediments at the sampling site). C- and Nconcentrations were determined on dry sediment using a CHN-Elemental Analyser (Carlo-Erba). Tot-P of sediment and seston was analysed as phosphate after acid hydrolysis at high temperature (340 ˚C) followed by Murphy and Riley procedure (1962). Sediment Pfractionation was performed using a modified Hieltjes and Lijklema (1980) scheme (Figure 1). In addition to the ordinary extraction scheme, digestion step was performed on the NaOH extractable P fraction in order to separate it into a reactive and a non-reactive form. Altogether, sediment P was separated into the following fractions (extractants and conditions are given within parenthesis): (a) loosely bound or labile P (1 M NH4 Cl at pH 7), (b) Fe- and Al-bound P (0.1 M NaOH) as reactive P, and non-reactive P after digestion of the extract (representing e.g. easily degradable organic P). By subtracting the concentration of NaOH-rP (reactive P) from NaOH-Tot P, NaOH-nrP (non reactive P) is determined (Furumai and Ohgaki, 1982), (c) Ca-bound P (0.5 M HCl) and (d) residual P (calculated by subtracting the sum of extracted P from the Tot-P, mainly consisting of refractory org-P but also including the inert P fraction). Extractions were made on the seston suspension on day 0 and on wet sediment immediately after sectioning of cores. Cpart and Npart , were captured on precombusted glass fibre filters (Whatman GF/C) and analysed using a CHN-Elemental Analyser (CarloErba). O-P was determined according to Murphy and Riley (1962). P in the overlying water is expressed as the net sediment retention (the difference between outand inflowing water).

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Figure 1. The modified Hieltjes & Lijklema (1980) scheme used for phosphorus fractionation, including a digestion step to distinguish reactive P (rP) and non-reactive P (nrP) in NaOH extracted P. Subtracting NaOH-rP from NaOH-Tot P gives NaOH-nrP. Table 1. Initial (day 0) values of chlorophyll a, bacterial biomass (B.B.), mean bacterial volume (MCV) and bacterial production (B.P.) in seston and in the 0–0.5 cm sediment layer (values are means 1 s.d. of 4 replicate samples and cores).

Chl. a (mg m Seston 146 Sediment 89.2

2)

B.B. (g m

2)

10.4 9.56 1.57 5.80 7.90 0.75

MCV (m3 ) 0.239 0.165

0.024 0.006

B.P. (mg C m 54.61 22.97

2

d

12.54 6.06

Characterization of seston and sediment The seston was dominated by large diatoms, a majority (> 80%) was alive and in good condition (intact cells with chloroplasts). Especially abundant were Aulacoseira islandica (Grun) O. Müller, Fragilaria crotonensis Kitt. and Asterionella formosa Hass. Seston additions corresponded to an input of 17.9 2.3 g C, 2.52 0.30 g N and 0.31 0.03 g P m 2 ( SD, n = 4). The chlorophyll a content of the added seston was 1.6 times higher than in the sediment (Table 1) or corresponding to the total sedimentation of seston in 13 m water depth with a mean chlorophyll a concentration of 11 g l 1 . Bacterial biomass was approximately equal in seston and surface sediments. The mean bacterial cell size, however, was significantly higher in the seston suspension (Mann-Whitney U-test p< 0.05) owing to the occurrence of large rod-shaped bacteria. Bacterial carbon in the seston suspension constituted

1)

Table 2. Temperature, redox potential (Eh) at 0.5 cm sediment depth and sediment water content of the 0–0.5 cm sediment layer in seston cores and controls (values are means 1 s.d. of 4 replicate cores).

Day Temp. Eh, seston Eh, control Water content Water content (˚C) cores (mV) cores (mV) seston cores control cores (%) (%) 0 1 2 4 7 12 20

9.1 9.0 9.0 9.0 9.0 7.2 6.5

324 260 190 324 298 272

22 33 47 15 21 36

321 294 228 315 320 305 307

36 15 102 43 12 28 25

94.70 94.21 95.52 95.79 96.04 95.64

0.84 0.62 0.58 0.86 0.19 0.55

93.82 93.59 93.86 94.61 94.96 94.26 94.88

0.40 0.56 0.63 0.50 1.37 0.44 0.29

ca. 10% of the total seston C. Bacterial production was approximately 2.4 times higher in the seston than in the original 0–0.5 cm sediment layer. Results Seston additions caused an immediate increase in bacterial production, cell-specific bacterial activity and total sediment metabolism (Figure 2). On day 1, bacterial production was 4.5 times higher in cores with seston addition than in control cores. Bacterial productivity at in situ temperature remained high the first week and then decreased linearly to approximately the same level as in the control cores on day 20. Cell-specific bacterial

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Figure 3. Temporal changes in (a) bacterial biomass (open boxes) and the NaOH-nrP phosphorus fraction (filled boxes) and (b) in the NH4 Cl phosphorus fraction (open dots) in the 0–0.5 cm sediment layers. Results are expressed as the differences between seston cores and controls (values are means 1 S.D. of 4 replicate cores). Results are expressed per g dry sediment. Also indicated is the C:P ratio (filled circles) calculated from bacterial biomass C and P in the NaOH-nrP fraction.

Figure 2. Temporal changes in (a) bacterial production, (b) bacterial specific growth rate and (c) heat production in the 0–0.5 cm sediment layers in seston cores (open symbols) and controls (filled symbols). Dots show bacterial production at 9 ˚C on days 12 and 20. All values are means of 4 replicate cores 1 S.D.

activity (specific growth rates) exhibited a slightly different pattern with maximum rates on day 12. Bacterial production at 9 ˚C (additional incubations performed due to decreasing experimental temperatures) on days 12 and 20 was low compared to values obtained at the same temperature during the first week of the experiment. Sediment heat production integrated over the experiment and assuming no diurnal variation could explain the oxidation of 3.20 and 1.33 g m 2 of C in the treatments with seston additions and in controls, respectively. Consequently, approximately 1.9 g m 2 of C or 11% of the Csest originally added was oxidized. The rapid increase in bacterial and overall sediment activity (sediment heat production) caused an increase in O2 consumption as shown by lower O2 concentrations in the overlying water on days 2, 4 and 7 (50–70% saturation in seston cores and 75–85% in the controls).

A small decrease in sediment redox potentials was also observed in both seston and control cores during the first days of the experiment (Table 2). During the last two weeks of the experiment, however, redox potentials at 0.5 cm depth varied around + 300 mV, indicating aerobic conditions in the top 5 mm of the sediment. Redox potentials decreased slowly with increasing sediment depth. The + 200 mV redox cline was found at approximately 1 cm sediment depth and the + 100 mV cline at 7 cm sediment depth in both seston and control cores. Bacterial biomass in the sediment approximately doubled after seston additions (Table 3), remained high until day 7 and then decreased by a factor 2 to day 20. The decrease in biomass between day 7 and 12 was caused by a decrease in bacterial abundance, as indicated by more or less constant mean bacterial volume over the first 12 days of the experiment (Table 3). Between day 12 and 20, however, the decrease in bacterial biomass can mainly be explained by a shift towards smaller cell sizes, shown as a decrease in volume from 0.24 to 0.17 m3 . A similar drop in bacterial

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60 Table 3. Changes in bacterial biomass and mean bacterial cell volume in the 0–0.5 cm sediment layer in seston cores and controls (values are means 1 s.d. of 4 replicate cores).

Day

Bacterial biomass (g m 2 ) Seston cores Control cores

0 1 2 4 7 12 20

17.5 21.4 17.3 21.2 22.3 14.7 11.1

1.74 3.99 5.60 5.56 3.48 3.55 1.02

Mean bacterial cell volume (m3 ) Seston cores Control cores

7.90 0.75 10.5 1.32 9.10 1.75 8.39 1.17 8.91 0.93 6.79 1.22 4.75 1.23

0.202 0.210 0.197 0.205 0.213 0.242 0.170

0.025 0.022 0.006 0.019 0.029 0.014 0.020

0.165 0.185 0.175 0.178 0.190 0.192 0.147

0.006 0.017 0.017 0.010 0.008 0.011 0.023

Table 4. The distribution of the different phosphorus pools in the added seston and in seston cores and contols (values are means 1 s.d. of 4 replicate cores, all values are expressed as g P/g D.W.) (Day 0 values in seston cores were caluclated as the weighted sum of sediment in control cores and the added seston).

Day

NH4 Cl-P (g P/g)

NaOH-rP (g P/g)

NaOH-nrP (g P/g)

HCl-P (g P/g)

Res-P (g P/g)

Tot-P (g P/g)

Seston

Seston cores 0 2 4 7 12 20 Control cores 0 2 4 7 12 20

260

1

260

2

1120

49

160

1

180

1

2000

178

172 97 219 150 115 154

9 33 42 25 16 55

497 497 625 540 587 629

21 15 86 60 52 235

863 641 774 800 795 806

23 15 76 92 96 111

297 350 395 380 412 368

31 16 44 24 15 29

312 526 379 545 416 661

63 274 130 121 209 132

2148 2110 2393 2415 2325 2618

78 225 146 94 214 322

117 85 108 90 103 93

15 25 30 19 13 6

645 612 575 562 750 574

34 123 127 51 113 2

702 541 553 522 627 650

23 90 134 34 59 69

382 361 358 402 405 381

51 27 99 11 36 15

394 746 546 556 452 761

103 153 197 137 94 271

2240 2345 2140 2133 2338 2460

28 67 237 172 99 306

biomass and volume was observed between day 12 and 20 in the control cores. The NaOH-nrP and NH4 Cl-P fractions showed a clear increase after seston additions (Table 4). The NaOH-nrP fraction in the added seston suspension, alone constituted more than 50% of the Psest . A significant relationship was found between mean values of the NaOH-nrP fraction and bacterial biomass expressed as the difference between seston supplied and control cores (linear regression, r2 = 0.858, p = 0.008). Expressed over time, both the NaOH-nrP fraction and bacterial biomass approximately doubled between day 0 and 7 and then decreased to the initial level by day 12 (Figure 3a). Assuming that 50% of the NaOH-nrP frac-

tion represented bacterial poly-P (Hupfer et al., 1995), a bacterial C:P ratio (bacterial biomass in carbon unit:P in the NaOH-nrP fraction) was calculated on a weight basis. This C:P ratio peaked on day 2 (36:1) and then decreased almost linearly to approximately 20:1 on day 20 (Figure 3a). The NH4 Cl-P fraction (representing labile P in the sediment) also exhibited significant changes during the experiment (Figure 3b). Between day 0 and 4 this P fraction approximately doubled and then quickly decreased from 110 g g 1 on day 4 to approximately 10 g g 1 on day 12. All phosphate in the overlying water of both seston supplied and control cores was trapped in the sediments during the first week of the experiment and released

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Figure 4. Mean net sediment retention (mg m 2 d 1 ) of P, (mean and 1 S.D. of 4 replicate cores) in seston cores (open bars) and controls (filled bars) and the amounts of P (open circles) supplied through the inflowing water during the different sampling intervals.

from day 12 on (Figure 4). Integrated over the whole experiment, the net P-release from the sediments corresponded to a release of 14 and 9 mg m 2 for the treatments with and without seston additions, respectively. Compared to the P-supply of 21 mg m 2 with the inflowing water, however, a P-fixation occurred in both treatments. Discussion Sediment bacteria have been suggested to play an important role in the uptake, storage and release of P by sediments through the incorporation into and release from intracellular poly-P granules (Gächter et al., 1988; Uhlmann & Bauer, 1988). This suggestion is supported by the recent discovery of bacterial polyP in lake sediments by Hupfer et al. (1995) and that bacterial poly-P was found to constitute between 31 and 50% of the NaOH-nrP sediment P fraction. The NaOH extraction step used in our and the above mentioned studies, will also when present, extract phytate (de Groot & Golterman, 1993; Golterman et al., 1998), a stable cyclic P-compound probably associated with bacterial metabolism. The NaOH-nrP fraction may therefore contain an unknown portion of phytate-P. However, P bound in sediment bacteria may be released through metabolic reactions, extracellular reactions and lysis of bacterial cells in response to changing environmental conditions, for example shift

towards anoxia (Boström et al., 1988) and should therefore constitute an important and potentially mobile P pool in sediments with a large NaOH-nrP fraction like the ones investigated in this study. The most noticeable result in this study is the close coupling and the concomitant changes in bacterial activity, bacterial biomass, the NaOH-nrP and labile P pools and the sediment uptake and subsequent release of phosphate as a result of the deposition of seston on the sediment surface. Altogether, these results illustrate an important role of bacteria in the P-regeneration in the oxic surface layers of the sediment and in the P-exchange between sediment and water. P-fractionations, normally performed to characterize sediments regarding the composition of the different P-pools were used to investigate possible shortterm changes within these pools. The results show that significant changes in some sediment P-pools can be detected within a time scale as short as one or a few days. C:P ratios of sediment bacteria, calculated from bacterial biomass carbon and the NaOH-nrP fraction assuming a 50% bacterial P content, decreased from 36 on day 2 to approximately 20 by the end of the experiment, indicating an active bacterial uptake and storage of P. In spite of the rough estimate of bacterial P, the ratios are in agreement with literature values of 18.4 (Fenchel & Blackburn, 1979), 14.9 (Gächter et al., 1988) and the 3 to 29 with a median value of 11 found by Vadstein et al. (1988). Even though the estimated bacterial C:P ratios were within the range described in literature they may be understimates, since algal polyP and other organic P-forms in the seston suspension may have been included in the NaOH-nrP fraction. The large size of this fraction (approximately 50% of the Tot-P) in the seston may indicate that non-bacterial P forms were included, but could also be explained by low C:P ratios of the seston-associated bacteria. Moreover, the P fractionation techniques using NaOH (0.1 M) early in the sequential extraction procedure, in this study only preceded by NH4 Cl, have disadvantages. Org-P may to some extent be hydrolyzed by NaOH (Golterman, 1988), resulting in an overestimation of NaOH-rP on behalf of Org-P (NaOH-nrP). Bacterial mortality was high in both seston supplied and control cores. This was shown as a decrease in bacterial biomass by a factor two between day 7 and 20. This increased bacterial mortality may directly and indirectly have been responsible for the observed Prelease from the sediments. Firstly, a direct P-release can be expected at bacterial lysis (Boström et al., 1988).

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62 Secondly, the diminishing bacterial community should have a decreasing ability of to take up and store and P liberated from the diatoms and the underlying sediment. A redox-controlled P-release from the sediment is less probable since oxic conditions were prevailing during the last two weeks of the experiment. A diffusion of P from deeper sediment layer is probable since the + 200 mV redox cline was at approximately 1 cm depth. A diffusion of P from deeper sediment layer may be an explanation to the tendency of increasing Tot-P observed over the experiment. This should, however, not affect our assumptions of an active role of bacteria in the regeneration of P. The uptake and release of P in control cores was similar to that in seston supplied cores but less pronounced. Also bacterial biomass and production in the control exhibited very similar but less pronounced patterns as in the seston treatment. These results can most probably be explained by the fact that the sediment cores were collected at the end of the autumn diatom bloom (the seasonal maximum in chlorophyll a in profundal sediments of the lake was recorded only a few days later) and it has to be assumed that an unknown amount of fresh seston was already deposited when sediment cores were collected. High numbers of large rod-shaped bacteria with a high cell-specific activity in the seston suggest an important role of these bacteria in the observed increase in bacterial activity after seston addition. A co-sedimentation of actively growing pelagic bacteria with settling diatoms has been observed in Lake Erken both during spring and autumn and have been suggested to play an important role in the high bacterial activity normally observed in the sediments in response to sedimentation events (Goedkoop & Törnblom, 1996). Some indirect evidence for the significance of cosettled bacteria in our study is given by the observations that mean bacterial cell volumes were higher in the seston than in the sediments and that mean bacterial cell volumes in the treatment with seston addition remained high or even increased until day 12. An active role of bacteria originating from the sediments before seston additions cannot be ruled out, however, since the high and increasing mean bacterial cell volumes observed also could be explained by an increased growth of these bacteria. Bacterial production was affected by the decrease in temperature as shown by bacterial production measurements at a higher temperature (9 ˚C) on days 12 and 20. The difference in production between 9 ˚C and the lower experimental temperature, however, was too small to explain the more than threefold

decline in bacterial production between days 7 and 20. Since the cell-specific bacterial activity remained high at least until day 12, the strong decrease in bacterial biomass can most probably be explained by grazing by meio- and microfauna and/or viral infection. A similar rapid decline in bacterial abundance and bacterial activity in Lake Erken sediments has been observed within days of the deposition of a spring diatom bloom (Goedkoop & Törnblom, 1996). Sediment heat production could explain the oxidation of ca. 11% of the Csest originally added and a comparison between sediment heat productions and bacterial production expressed in carbon units, suggests that the total activity was largely bacterial. For comparison, Goedkoop & Johnson (1996) showed that between 1.9 and 12.4% of the seston C deposited during spring was mineralized by bacteria during the same period. Further comparisons with natural conditions are difficult to make since the amount of seston added was too large to mimic a natural diatom sedimentation event. To summarize, the microbial sediment community responded quickly to the deposition of natural seston in terms of increased activity and biomass. Changes in bacterial activity and biomass were directly related to changes in the NaOH-nrP and labile P-pools in the sediment and to the uptake and subsequent release of phosphate by the sediments. Decreasing bacterial C:P ratios over time indicated an active uptake and storage of P by the bacterial community. Althogether, our results imply that uptake and storage of P by sediment bacteria and/or bacteria introduced with seston assemblages can be of importance in the P-turnover following the sedimentation of natural seston. Acknowledgements We would like to acknowledge Anu Toom for assistance in P-fractionations, Stefan Djurström for help in the construction of the experimental setup, Jan Johansson for laboratory assistance, Kjell Hellström for counting and measuring bacteria and Kurt Petterson for providing space at the Erken Laboratory. This work was supported by grant nr 13416 from the Swedish Environmental Protection Agency. References Bell, R. T. & I. Ahlgren, 1987. Thymidine incorporation and microbial respiration in the surface sediment of a hypereutrophic lake. Limnol. Oceanogr. 32: 476–482.

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