Microb Ecol (1984) 10:335-344
MICROBIAL ECOLOGY 9 1984 Springer-Verlag
Benthic Bacterial Biomass Supported by Streamwater Dissolved Organic Matter Thomas L. Bott, t Louis A. Kaplan, t and Frank T. Kuserk 2 ~Stroud Water Research Center, Academy of Natural Sciences, R.D. #1, Box 512, Avondale, Pennsylvania 1931l, USA; and 2Department of Biology, Moravian College,Bethlehem,Pennsylvania 18018, USA
Abstract. Bacterial biomass in surface sediments of a headwater stream was measured as a function of dissolved organic carbon (DOC) flux and temperature. Bacterial biomass was estimated using epifluorescence microscopic counts (EMC) and A T P determinations during exposure to streamwater containing 1,788 t~g DOC/liter and after transfer to groundwater containing 693 #g DOC/liter. Numbers of bacteria and A T P concentrations averaged 1.36 x 109 cells and 1,064 ng per gram dry sediment, respectively, under initial DOC exposure. After transfer to low DOC water, biomass estimates dropped by 53 and 55% from E M C and ATP, respectively. The decline to a new steady state occurred within 4 days from A T P assays and within 11 days from EMC measures. A 4~ difference during these exposures had little effect on generation times. The experiment indicated that 27.59 mg/hour of natural DOC supported a steady state bacterial biomass of approximately 10 #g C/g dry weight o f sediment (from EMC determinations). Steady state bacterial biomass estimates on sediments that were previously muffled to remove organic matter were approximately 20-fold lower. The ratio of G T P : A T P indicated differences in physiological condition or c om m uni t y composition between natural and muffled sediments.
Introduction The sources o f nutrients supporting bacterial communities in surface sediments are not well described but m ay be o f diverse origin that includes deeper sediments, adjacent biota, and overlying water. Dissolved organic matter (DOM) has been shown to be a significant portion of the detrital carbon pool in lakes and rivers [33, 34], estuaries [35], and oceans [26]. Our studies are based on the recognized fact that bacteria are principal users of DOM. Many studies of the utilization o f individual compounds have been done for planktonic [3, 36, 37] and sediment [11, 22] bacteria. However, virtually nothing is known concerning the extent to which bacterial populations in surface sediments are supported by the heterogenous supply of D O M in overlying water. Here we have estimated this interaction in a small headwater stream. Our approach was to work with a bacterial community in steady state, that is, in
336
T . L . Bott et al.
equilibrium with DOM supply and grazing, and to examine the community response when transferred to a lower DOM supply. The approach was somew h a t a n a l o g o u s t o t h a t e m p l o y e d b y B r o c k a n d B r o c k [5] t o s t u d y t h e g r o w t h rate of thermophilic algae. In addition, we examined the response of some metabolic parameters to DOM manipulation.
Materials and M e t h o d s
Study Site Experiments were conducted in Saw Mill Spring, a woodland spring seep in the White Clay Creek drainage, Chester Co., PA. Concentrations of DOC increased downstream from the spring source. DOC concentrations at baseflow about 30 m downstream varied seasonally from 1.0-2.0 mg/liter, and were highest in summer and autumn. D O M sources within the seep were exudates from algae and higher plants, leachates of leaf litter and other particulate organic matter, and animal excretions primarily from invertebrates. Additional detail is found in Kaplan et al. [15].
Field Experiments Three plastic trays (38 cm long x 7.5 cm wide x 2.5 cm deep) were filled with sediment from the downstream site which had been sieved through a 4 m m screen to reduce its heterogeneity. The trays were placed individually in 75 cm long troughs of vinyl rain gutter fitted with end pieces. Spring water was gravity-fed to each trough through rubber tubing at a rate controlled by adjustable clamps. Inflowing water was filtered through glass wool placed in the upper l0 cm of the gutter in order to reduce the deposition of particles onto the sediments and the introduction of new organisms. The systems were covered with black plastic to eliminate algal growth and each day the trays were removed, the walls of the trough were cleaned, the glass wool changed, and the flow rates adjusted. After incubation for 19 days at the high DOC downstream site, the trays were transferred to another set of troughs at the low DOC spring source where they were incubated for another 36 days. A second experiment was performed using sediment from the downstream site that had been muffled at 550~ for 4 hours to eliminate organic matter. Trays of sediment were incubated at the downstream site for the entire 33 day experiment.
Physical and Chemical Measurements Water samples for DOC analyses were collected in triplicate 2 or 3 times weekly in precombusted (550"C for 5 hours) glass serum bottles 0 2 5 ml) and stored at 5~ Samples were then filtered through precombusted Gelman AE glass-fiber filters and the DOC concentration of the filtrate was determined using a Dohrmann DC 54 DOC analyzer. Water temperature was measured daily using a mercury thermometer and monitored continuously during a 5 day period using a Bristol thermograph. Discharge rates from each trough were measured using a graduated cylinder and stop watch.
Microbiological Analyses Surface sediments from each tray were sampled to a depth of 2-3 m m approximately 3 times weekly. Plastic rings (15 m m diameter) that were pressed into the sediment to delimit the sampling
Bacterial Biomass Supported by Streamwater DOM
337
area were left in place so that the area was not resampled. On each sampling date triplicate samples (approximately 0.5 g wet weight each) were taken for epifluorescence microscope counts (EMC) of bacteria and for ATP determinations. For EMC determinations, sediment was placed into a sterile test tube and sterile 0.2 M phosphate buffer (pH = 7.0) was added to a 5 ml volume. Samples were kept on ice, brought back to the laboratory, and then mixed vigorously and diluted serially either to 1:10 or l:100. Bacteria were stained for 10 min with 4,6-diamidino-2-phenylindole (DAPI, 0.5 #g/ml, Sigma Chemical Co., St. Louis, MO) and each sample was filtered through a 0.2 #m pore size Nucleopore filter stained with Irgalan black [25]. Filters were mounted in immersion oil and bacteria were counted using a Zeiss Universal Microscope equipped with epifluorescence illumination. Usually 20 fields were counted per filter for a total of 300-500 organisms. Cells on and offparticles were counted and the assumption was made that the number of cells on the underside of particles equaled that on the viewed side. Data were corrected for the small number o f cells associated with stains and Irgalan black treated filters. The sediments were dried at 600C and weighed. Samples for ATP analyses were placed into sterile test tubes containing 5.0 ml ice cold 3 N H3PO4 [18], vigorously shaken for 1 min, and frozen for further analysis. After thawing, I ml of 0.025 M Tris buffer (pH = 7.75) was added and the samples were filtered through preweighed Gelman AE glass-fiber filters into sterile shell vials. Tris-EDTA buffer (1 ml) was added to 4 ml of filtrate, and samples were neutralized to a pH o f 7.75 with NaOH. The sample volume was brought to 10 ml with 0.025 M Tris buffer. ATP was analyzed by the procedure of Stanley and Williams [28] using a Beckman liquid scintillation counter in noncoincidence mode. Aliquots (0.2 ml) of the neutralized extract were mixed with 0.1 ml of buffered MgC12 solution and 0.5 ml 0.025 M Tris buffer. To this, 0.5 ml of hydrated [19] firefly lantern extract (FLE 50, Sigma Chemical Co., St. Louis, MO) were added, and the solution was vortexed and bioluminescence was measured 15 sec after enzyme addition for 6 sec. Values were corrected for efficiency of the light reaction by t2C-ATP additions and ATP recovery from the sediments, determined by the recovery o f 14C-ATP initially added to the cold 3 N H3PO 4. Filters were dried for weight determinations. GTP (which may comain UTP and CTP as contaminants) was measured by conversion to ATP after ATP hydrolysis [16]. These analyses, corresponding ATP analyses and all ATP analyses on muffled sediments, were made using a Turner Model 20 Photometer. Light flux was integrated from 3.0-5.5 see.
Growth Rate Experiment Growth rates were determined from increases in cell number o f bacterial microcolonies [4]. Microscope slides were etched with a diamond marking pencil to provide identifiable microscopic fields. The slides were placed at the downstream field site at 15~ for 16 hours for attachment of the flora after which they were brought to the laboratory. Slides were covered with site water and phase contrast photomicrographs were taken using a water immersion lens, Zeiss Universal microscope and camera. To determine the effect of temperature on growth rates, slides were incubated either at 10 or 15~ in sterile flasks containing water from the spring source. Slides were rephotographed at intervals of 5-8 hours and incubated in fresh water for the intervening periods. Water used to cover the slides and for incubation was membrane-filter sterilized (0.45 ttm pore size).
Results
Environmental Variables Exposure conditions for sediment communities during each phase of the exp e r i m e n t a r e p r e s e n t e d i n T a b l e 1. D O C c o n c e n t r a t i o n , w a t e r t e m p e r a t u r e , a n d
338
T . L . Bolt et al.
Table 1.
E n v i r o n m e n t a l p a r a m e t e r s at the study locations 5: + SD (n) Parameter
Downstream
DOC concentration (#g/liter)" T e m p e r a t u r e (~ ~ Discharge (liter/hour) h
Sourcc
1,788 + 175 (7) 16.4 _+ 2.1 (18) 37.18 _+ 5.70 (57)
693 + 204 (14) 't 12.5 _+ 0.6 (32) d 34.60 + 6.01 (93)'
For D O C c o n c e n t r a t i o n and t e m p e r a t u r e , n is the n u m b e r o f d a i l y m e a n s a ve ra ge d dur ing each phase of the e x p e r i m e n t : for discharge, n is the n u m b e r of i n d i v i d u a l m e a s u r e s m a d e on i n d i v i d u a l troughs. Tested for significance using t test b Tested for significance using 2 level nested A N O V A , c~ = 0.05 ~lc~ = 0.001
1 X 1010
~_ 1
Ful
•
10 4
A r
gl
0~ ev-
"o e
1 X 10 9 \
~
MICROBIAL ECOLOGY 9 1984 Springer-Verlag
Benthic Bacterial Biomass Supported by Streamwater Dissolved Organic Matter Thomas L. Bott, t Louis A. Kaplan, t and Frank T. Kuserk 2 ~Stroud Water Research Center, Academy of Natural Sciences, R.D. #1, Box 512, Avondale, Pennsylvania 1931l, USA; and 2Department of Biology, Moravian College,Bethlehem,Pennsylvania 18018, USA
Abstract. Bacterial biomass in surface sediments of a headwater stream was measured as a function of dissolved organic carbon (DOC) flux and temperature. Bacterial biomass was estimated using epifluorescence microscopic counts (EMC) and A T P determinations during exposure to streamwater containing 1,788 t~g DOC/liter and after transfer to groundwater containing 693 #g DOC/liter. Numbers of bacteria and A T P concentrations averaged 1.36 x 109 cells and 1,064 ng per gram dry sediment, respectively, under initial DOC exposure. After transfer to low DOC water, biomass estimates dropped by 53 and 55% from E M C and ATP, respectively. The decline to a new steady state occurred within 4 days from A T P assays and within 11 days from EMC measures. A 4~ difference during these exposures had little effect on generation times. The experiment indicated that 27.59 mg/hour of natural DOC supported a steady state bacterial biomass of approximately 10 #g C/g dry weight o f sediment (from EMC determinations). Steady state bacterial biomass estimates on sediments that were previously muffled to remove organic matter were approximately 20-fold lower. The ratio of G T P : A T P indicated differences in physiological condition or c om m uni t y composition between natural and muffled sediments.
Introduction The sources o f nutrients supporting bacterial communities in surface sediments are not well described but m ay be o f diverse origin that includes deeper sediments, adjacent biota, and overlying water. Dissolved organic matter (DOM) has been shown to be a significant portion of the detrital carbon pool in lakes and rivers [33, 34], estuaries [35], and oceans [26]. Our studies are based on the recognized fact that bacteria are principal users of DOM. Many studies of the utilization o f individual compounds have been done for planktonic [3, 36, 37] and sediment [11, 22] bacteria. However, virtually nothing is known concerning the extent to which bacterial populations in surface sediments are supported by the heterogenous supply of D O M in overlying water. Here we have estimated this interaction in a small headwater stream. Our approach was to work with a bacterial community in steady state, that is, in
336
T . L . Bott et al.
equilibrium with DOM supply and grazing, and to examine the community response when transferred to a lower DOM supply. The approach was somew h a t a n a l o g o u s t o t h a t e m p l o y e d b y B r o c k a n d B r o c k [5] t o s t u d y t h e g r o w t h rate of thermophilic algae. In addition, we examined the response of some metabolic parameters to DOM manipulation.
Materials and M e t h o d s
Study Site Experiments were conducted in Saw Mill Spring, a woodland spring seep in the White Clay Creek drainage, Chester Co., PA. Concentrations of DOC increased downstream from the spring source. DOC concentrations at baseflow about 30 m downstream varied seasonally from 1.0-2.0 mg/liter, and were highest in summer and autumn. D O M sources within the seep were exudates from algae and higher plants, leachates of leaf litter and other particulate organic matter, and animal excretions primarily from invertebrates. Additional detail is found in Kaplan et al. [15].
Field Experiments Three plastic trays (38 cm long x 7.5 cm wide x 2.5 cm deep) were filled with sediment from the downstream site which had been sieved through a 4 m m screen to reduce its heterogeneity. The trays were placed individually in 75 cm long troughs of vinyl rain gutter fitted with end pieces. Spring water was gravity-fed to each trough through rubber tubing at a rate controlled by adjustable clamps. Inflowing water was filtered through glass wool placed in the upper l0 cm of the gutter in order to reduce the deposition of particles onto the sediments and the introduction of new organisms. The systems were covered with black plastic to eliminate algal growth and each day the trays were removed, the walls of the trough were cleaned, the glass wool changed, and the flow rates adjusted. After incubation for 19 days at the high DOC downstream site, the trays were transferred to another set of troughs at the low DOC spring source where they were incubated for another 36 days. A second experiment was performed using sediment from the downstream site that had been muffled at 550~ for 4 hours to eliminate organic matter. Trays of sediment were incubated at the downstream site for the entire 33 day experiment.
Physical and Chemical Measurements Water samples for DOC analyses were collected in triplicate 2 or 3 times weekly in precombusted (550"C for 5 hours) glass serum bottles 0 2 5 ml) and stored at 5~ Samples were then filtered through precombusted Gelman AE glass-fiber filters and the DOC concentration of the filtrate was determined using a Dohrmann DC 54 DOC analyzer. Water temperature was measured daily using a mercury thermometer and monitored continuously during a 5 day period using a Bristol thermograph. Discharge rates from each trough were measured using a graduated cylinder and stop watch.
Microbiological Analyses Surface sediments from each tray were sampled to a depth of 2-3 m m approximately 3 times weekly. Plastic rings (15 m m diameter) that were pressed into the sediment to delimit the sampling
Bacterial Biomass Supported by Streamwater DOM
337
area were left in place so that the area was not resampled. On each sampling date triplicate samples (approximately 0.5 g wet weight each) were taken for epifluorescence microscope counts (EMC) of bacteria and for ATP determinations. For EMC determinations, sediment was placed into a sterile test tube and sterile 0.2 M phosphate buffer (pH = 7.0) was added to a 5 ml volume. Samples were kept on ice, brought back to the laboratory, and then mixed vigorously and diluted serially either to 1:10 or l:100. Bacteria were stained for 10 min with 4,6-diamidino-2-phenylindole (DAPI, 0.5 #g/ml, Sigma Chemical Co., St. Louis, MO) and each sample was filtered through a 0.2 #m pore size Nucleopore filter stained with Irgalan black [25]. Filters were mounted in immersion oil and bacteria were counted using a Zeiss Universal Microscope equipped with epifluorescence illumination. Usually 20 fields were counted per filter for a total of 300-500 organisms. Cells on and offparticles were counted and the assumption was made that the number of cells on the underside of particles equaled that on the viewed side. Data were corrected for the small number o f cells associated with stains and Irgalan black treated filters. The sediments were dried at 600C and weighed. Samples for ATP analyses were placed into sterile test tubes containing 5.0 ml ice cold 3 N H3PO4 [18], vigorously shaken for 1 min, and frozen for further analysis. After thawing, I ml of 0.025 M Tris buffer (pH = 7.75) was added and the samples were filtered through preweighed Gelman AE glass-fiber filters into sterile shell vials. Tris-EDTA buffer (1 ml) was added to 4 ml of filtrate, and samples were neutralized to a pH o f 7.75 with NaOH. The sample volume was brought to 10 ml with 0.025 M Tris buffer. ATP was analyzed by the procedure of Stanley and Williams [28] using a Beckman liquid scintillation counter in noncoincidence mode. Aliquots (0.2 ml) of the neutralized extract were mixed with 0.1 ml of buffered MgC12 solution and 0.5 ml 0.025 M Tris buffer. To this, 0.5 ml of hydrated [19] firefly lantern extract (FLE 50, Sigma Chemical Co., St. Louis, MO) were added, and the solution was vortexed and bioluminescence was measured 15 sec after enzyme addition for 6 sec. Values were corrected for efficiency of the light reaction by t2C-ATP additions and ATP recovery from the sediments, determined by the recovery o f 14C-ATP initially added to the cold 3 N H3PO 4. Filters were dried for weight determinations. GTP (which may comain UTP and CTP as contaminants) was measured by conversion to ATP after ATP hydrolysis [16]. These analyses, corresponding ATP analyses and all ATP analyses on muffled sediments, were made using a Turner Model 20 Photometer. Light flux was integrated from 3.0-5.5 see.
Growth Rate Experiment Growth rates were determined from increases in cell number o f bacterial microcolonies [4]. Microscope slides were etched with a diamond marking pencil to provide identifiable microscopic fields. The slides were placed at the downstream field site at 15~ for 16 hours for attachment of the flora after which they were brought to the laboratory. Slides were covered with site water and phase contrast photomicrographs were taken using a water immersion lens, Zeiss Universal microscope and camera. To determine the effect of temperature on growth rates, slides were incubated either at 10 or 15~ in sterile flasks containing water from the spring source. Slides were rephotographed at intervals of 5-8 hours and incubated in fresh water for the intervening periods. Water used to cover the slides and for incubation was membrane-filter sterilized (0.45 ttm pore size).
Results
Environmental Variables Exposure conditions for sediment communities during each phase of the exp e r i m e n t a r e p r e s e n t e d i n T a b l e 1. D O C c o n c e n t r a t i o n , w a t e r t e m p e r a t u r e , a n d
338
T . L . Bolt et al.
Table 1.
E n v i r o n m e n t a l p a r a m e t e r s at the study locations 5: + SD (n) Parameter
Downstream
DOC concentration (#g/liter)" T e m p e r a t u r e (~ ~ Discharge (liter/hour) h
Sourcc
1,788 + 175 (7) 16.4 _+ 2.1 (18) 37.18 _+ 5.70 (57)
693 + 204 (14) 't 12.5 _+ 0.6 (32) d 34.60 + 6.01 (93)'
For D O C c o n c e n t r a t i o n and t e m p e r a t u r e , n is the n u m b e r o f d a i l y m e a n s a ve ra ge d dur ing each phase of the e x p e r i m e n t : for discharge, n is the n u m b e r of i n d i v i d u a l m e a s u r e s m a d e on i n d i v i d u a l troughs. Tested for significance using t test b Tested for significance using 2 level nested A N O V A , c~ = 0.05 ~lc~ = 0.001
1 X 1010
~_ 1
Ful
•
10 4
A r
gl
0~ ev-
"o e
1 X 10 9 \
~
