Next Article 
Applied and Environmental Microbiology, February 2000, p. 455-466, Vol. 66, No. 2
0099-2240/00/$04.00+0
Copyright © 2000, American Society for Microbiology. All rights reserved.
Limitation of Bacterial Growth by Dissolved Organic
Matter and Iron in the Southern Ocean
Matthew J.
Church,1,*
David A.
Hutchins,2 and
Hugh W.
Ducklow1
School of Marine Science, The College of
William and Mary, Gloucester Point, Virginia
23062-1346,1 and College of Marine
Studies, University of Delaware, Lewes, Delaware
199582
Received 17 August 1999/Accepted 10 November 1999
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ABSTRACT |
The importance of resource limitation in controlling bacterial
growth in the high-nutrient, low-chlorophyll (HNLC) region of the
Southern Ocean was experimentally determined during February and March
1998. Organic- and inorganic-nutrient enrichment experiments were
performed between 42°S and 55°S along 141°E. Bacterial abundance, mean cell volume, and [3H]thymidine and
[3H]leucine incorporation were measured during 4- to
5-day incubations. Bacterial biomass, production, and rates of growth
all responded to organic enrichments in three of the four experiments.
These results indicate that bacterial growth was constrained primarily by the availability of dissolved organic matter. Bacterial growth in
the subtropical front, subantarctic zone, and subantarctic front
responded most favorably to additions of dissolved free amino acids or
glucose plus ammonium. Bacterial growth in these regions may be limited
by input of both organic matter and reduced nitrogen. Unlike similar
experimental results in other HNLC regions (subarctic and equatorial
Pacific), growth stimulation of bacteria in the Southern Ocean resulted
in significant biomass accumulation, apparently by stimulating
bacterial growth in excess of removal processes. Bacterial growth was
relatively unchanged by additions of iron alone; however, additions of
glucose plus iron resulted in substantial increases in rates of
bacterial growth and biomass accumulation. These results imply that
bacterial growth efficiency and nitrogen utilization may be partly
constrained by iron availability in the HNLC Southern Ocean.
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INTRODUCTION |
The factors that regulate the growth
of marine heterotrophic bacteria are ecologically and biogeochemically
important to the cycling of energy and materials in the ocean.
Bacterioplankton often dominate the biomass of planktonic food webs,
making their role in nutrient and energy fluxes crucial for the
organization of marine ecosystems (1, 12, 18, 21, 22, 23,
28). Constraints on the growth rates of bacterioplankton may not
be consistent in different marine environments. Bacterial
growth rates may be limited by dissolved organic matter (DOM)
quality (8, 11, 34), inorganic nutrients (51,
57), temperature (36, 45, 54, 61), viral infection
(48), or micronutrients such as iron (29, 30,
43). Bacterial stocks are the result of growth and removal
processes that include grazing (19, 63), viral infection
(48), and physical mixing (17). Each of these factors may limit bacterial growth over different temporal and spatial
scales (17).
The motivation behind this study was to identify the factors limiting
bacterial growth in the pelagic Southern Ocean. The Southern Ocean
(3, 42), the equatorial Pacific (7, 20, 33), and
the subarctic Pacific (34, 37) have all been characterized as high-nutrient, low-chlorophyll (HNLC) oceans. Many hypotheses have
been proposed to explain the existence of HNLC regions, but most
attention has focused on the importance of iron in limiting phytoplankton growth in HNLC systems (15, 38, 41, 42). HNLC
oceans typically have subnanomolar concentrations of dissolved iron
(41, 42, 52, 53). Such low concentrations of dissolved iron
are known to limit phytoplankton growth, but because bacteria are
widely regarded as more efficient competitors for limiting nutrients,
limitation of bacterial growth by dissolved iron has been relatively
neglected. Iron is an essential nutrient for bacteria because it is
part of cytochrome c, a component of the electron transport
chain in the respiratory system. Thus, iron deficiency might restrict
the growth efficiency of heterotrophic bacteria (58). The
few investigations into the dependence of bacteria on iron indicate
that iron may restrict bacterial conversion efficiency and growth rates
in oceanic and coastal regions of HNLC oceans (29, 43, 58).
Studies in the equatorial and subarctic Pacific indicated that the
microbial loop might serve an important role in regulating fluxes of
material in HNLC systems (7, 20, 33, 37). DOM constitutes a
potentially large, exportable pool of reduced carbon, and
quantification of DOM fluxes is essential for understanding the ocean
carbon cycle (7, 9, 62). The quality of available resources
may control bacterial growth rates and standing stocks in some systems.
The persistence of DOM in many marine systems suggests that DOM
production and subsequent bacterial consumption may be uncoupled in
either space or time. Such persistence indicates that some factor
limits bacterial utilization of the available DOM substrates (8,
9, 11, 57, 62, 64). Determination of whether bacterial growth
limitation results from poor-quality DOM requires experiments that
monitor the bacterial response to representative DOM substrates added
to seawater batch cultures (34, 36, 63).
Several investigations have evaluated how the quality of organic
material affects bacterial growth in marine systems. Bacterial growth
efficiency and growth rates may depend on the stoichiometric C/N ratio
of the organic substrate rather than on the specific type of organic
substrates utilized for growth (24, 25). In the Sargasso
Sea, bacterial growth may be limited by input of high-quality (labile)
organic carbon (8). In contrast, Kirchman (34)
found that bacterial growth rates in the subarctic Pacific depended
more on specific organic substrates.
Bacterial growth can also depend on the supply and quality of inorganic
nutrients (51, 64). Bacteria have been shown to be effective
competitors with phytoplankton for ammonium and phosphate (14, 35,
56), and in some systems bacterial uptake of mineral nutrients
dominates nutrient fluxes (13, 60). The nitrogen and
phosphorus requirements of bacteria are large because of their high
cellular nucleic acid and protein contents, which require them to
sustain low intracellular C/N/P ratios (24, 25). Studies from both open ocean and coastal environments suggest that most marine
bacterial nitrogen requirements can be met by dissolved free amino
acids (DFAA) or ammonium (NH4+) (31, 32,
35). Specific carbon sources used to fulfill oceanic bacterial
demands are not as well defined, but monosaccharides like glucose
appear to support a large fraction of the bacterial carbon requirement
(49). The complexity of microbial communities requires
investigations that focus on the possibility that multiple factors may
interact to control bacterial growth. Resource limitation may construct
synergistic or antagonistic relationships within marine food webs,
potentially controlling the growth of marine bacteria. For example,
Hutchins et al. (29) hypothesized that iron-limited
phytoplankton growth could restrict carbon flow to heterotrophic
bacteria, resulting in an interaction between iron and carbon as
possible limiting resources.
The present study sought to determine whether resources limited
bacterial growth in the pelagic Southern Ocean. Specifically, we tested
whether additions of labile organic material (glucose and amino acids)
and/or inorganic nutrients (ammonium, phosphate, and dissolved iron)
stimulated rates of bacterial growth. By assessing how rates of
bacterial production change relative to changes in standing biomass, we
address the relative importance of DOM and iron in controlling rates of
bacterial growth in the Southern Ocean.
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MATERIALS AND METHODS |
Study site.
Sampling for these experiments took place aboard
the Australian vessel R/V Aurora Australis between 2 and 26 March 1998. The cruise followed a southerly transect along 141°E
between 42°S and 55°S (Fig. 1). The
cruise track intersected several frontal systems, including the
subtropical front (STF) (42°S), the subantarctic front (SAF)
(51°S), and the Antarctic polar front (APF) (54°S). Experiments
were performed in each of these fronts and at one location inside the
subantarctic zone (SAZ) at 47°S. Although they are variable in their
locations, each frontal system has a characteristic chemical and
physical hydrographic signature (5, 50). Generally,
concentrations of major nutrients (NO3
and
PO4+) increased along a southerly gradient,
while the trace nutrient iron was found in subnanomolar concentrations
throughout the entire study region (40, 52, 53;
P. W. Boyd, A. C. Crossley, G. R. DiTullio, F. B. Griffiths, D. A. Hutchins, B. Queguiner, P. N. Sedwick, and
T. W. Trull, submitted for publication).
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FIG. 1.
Study site and cruise track of R/V Aurora
Australis voyage 6, 28 February to 3 April 1998. Stars represent
stations where experiments were conducted (42°S, 47°S, 51°S, and
54°S at 141°E). Approximate locations of permanent frontal and
water-mass boundaries are indicated.
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Experimental design.
The overall design of these experiments
was to add amendments (organic and/or inorganic) to unmanipulated
(whole) seawater, incubate at in situ temperatures, and monitor changes
in bacterial growth, abundance, and biomass over a 4- to 5-day
incubation period. The decision to use unfiltered rather than
size-fractionated seawater (i.e., grazer-reduced treatments) was
made to minimize risks of potential contamination of samples with
metals or DOM as an artifact of filtration (8).
Water was collected, supplemented with amendments, and then placed in
darkened incubators for the duration of the experiment.
Incubation in
the dark obviated stimulation of phytoplankton growth
and subsequent
organic enrichment artifacts. Seawater was collected
from a depth of
approximately 15 m with an all-Teflon, trace-metal-clean
pump
system (
29). The seawater was pumped directly into a
trace-metal
clean incubation laboratory van, where water was dispensed
into
2-liter polycarbonate bottles. All bottles used in these
experiments
were soaked for 48 h in 10% HCl and then rinsed three
times with
sample water. The 2-liter polycarbonate bottles were filled,
capped,
and carried to a positive-pressure hood, where 175-ml
polyethylene
bottles were filled for each treatment. Duplicate
treatments were
prepared and sampled for all experiments. The sample
handling
and setup were designed to reduce potential metal
contamination,
although contamination was not directly measured.
Plastic gloves
were worn during all sample
transfers.
All substrates were prepared from commercially available reagents.
Glucose, ammonium, and phosphate additions were made from
dry stocks
dissolved in MilliQ-water. Stocks were sterilized through
0.2-µm-pore-size Acrodisc filters (HT Tuffryn membrane) which
had
been flushed several times with MilliQ-water prior to sample
filtration. No efforts were made to remove possible metal contaminates
in the glucose stocks; however, the effect of inadvertent contamination
may have been minimized, because differential responses to treatments
containing glucose plus Fe relative to treatments containing glucose
alone were observed (see Results). Initial iron stocks (17.9 µM
FeCl
2) were made in 0.01% N HCl. Controls consisted of
untreated,
whole seawater without amendments. Combined nutrient and
substrate
additions contained the same concentrations of glucose,
ammonium,
phosphate, and iron as for treatments where these amendments
were
added individually (Table
1). The
concentrations of glucose differed
in each experiment, while all other
treatment concentrations were
constant for all experiments. Amino acid
additions were from a
commercially available mixture of 20 amino acids
(Pierce Chemical),
from which metals had been removed using a Chelex
ion-exchange
resin column (
47).
All sampling was done in a positive-pressure, trace-metal-clean
incubation van. Daily samples of approximately 30 ml were
poured from
polyethylene incubation bottles into acid-cleaned,
MilliQ-water-rinsed
50-ml polycarbonate tubes. The tubes were
then transferred to a
radiation van for preparation of incubation
mixtures for
[
3H]thymidine (TdR) and [
3H]leucine (Leu)
incorporation assays. TdR incorporation may be
used as a proxy for cell
division and DNA synthesis, while Leu
incorporation estimated
prokaryotic protein production
rates.
TdR and Leu incorporation assays.
Incubations for
measurements of TdR and Leu incorporation were carried out in on-deck,
flowthrough incubators or in shipboard refrigerated incubators. On-deck
incubators were maintained at surface water temperatures with
circulating surface seawater. Incorporation of TdR and Leu was measured
by the microcentrifugation procedure described by Smith and Azam
(55). High-specific-activity TdR and Leu (TdR, 79 Ci
mmol1; Leu, 179 Ci mmol1) (New England
Nuclear) were added to 2-ml microcentrifuge tubes, followed by the
addition of 1.5 ml of sample water to start the incubations. The final
concentration of both TdR and Leu was 20 nM. Triplicate samples of both
TdR and Leu from each treatment bottle and each time point were
incubated. Time zero blanks for both TdR and Leu were killed with 5%
(final concentration) trichloroacetic acid. Microcentrifuge tubes for
TdR and Leu incorporation were placed in darkened floating racks in
shaded incubators and incubated for 4 to 16 h, depending on the
expected activity of the samples and the temperature of the water.
Incubations were terminated by the addition of 100 µl of 100%
trichloroacetic acid (final concentration, 5%). Samples were
immediately frozen for subsequent radioassay following the cruise.
Samples were processed as described by Smith and Azam (
55)
and
Carlson et al. (
10). Packard Ultima Gold scintillation
cocktail
was added to the pellet, and the radioactivity of each sample
was counted with a Wallac 1409 liquid scintillation counter. Rates
of
isotope incorporation for each sample were calculated as the
average
from three replicates minus the value of the blank. Disintegrations
per
minute were calculated based on counts per minute standardized
to
external quench parameters and the counting efficiency. Counting
errors
were less than 20%.
Cellular abundance and volumes.
Samples for determination of
bacterial abundance and cell volume were collected in 50-ml
polyethylene tubes and preserved in filtered glutaraldehyde
(0.2-µm-pore-size filter; 2% final concentration). Samples were
immediately filtered onto blackened 0.2-µm-pore-size polycarbonate
membrane filters (Poretics Corp.) and stained with a 0.05% solution of
acridine orange. The volume filtered varied depending on cell density,
with the objective of evenly distributing 100 to 300 cells per
microscope field for image analysis. Filters were affixed to
microscope slides with a small drop of Resolve immersion oil
and mounted with a cover slide (28). Filters were frozen
until processing.
Cell volumes were determined using a video image analysis system. Cells
were enumerated on a Zeiss Axiophot epifluorescence
microscope at a
magnification of ×1,000. Video images of the 24-
by 24-µm microscope
field were captured and stored using Zeiss
VIDAS VIDEOPLAN image
analysis software. Fluorescence was achieved
using blue excitation (450 to 490 nm) and a 520-nm emission filter
from a 200-W mercury lamp.
Sufficient video images were captured
from each filter to yield between
300 and 1,000 measurements of
individual cells. The length, width,
area, and perimeter of each
cell were measured, and cell volumes were
calculated using algorithms
calibrated with fluorescent beads
(
2). Algorithms included
edge detection to minimize halo
effects (
4). Cell abundance
was determined by visual cell
counts where at least 300 cells
per filter were
counted.
Conversion factors.
Conversion factors from the literature
were employed to translate incorporation and biovolume (abundance × mean cell volume) into carbon-based estimates of production and
biomass. TdR incorporation rates were converted using 2 × 1018 cells mol of TdR1 and 120 µg of C
µm3 (18, 22, 39). Leu incorporation could
not be measured for the DFAA and DFAA plus Fe additions because the
amino acid mixture contained unlabeled leucine and extracellular
isotope dilution prevented signal detection. For this reason, estimates
of bacterial production (micrograms of C liter1
day1) were derived exclusively from TdR incorporation
rates. Leu incorporation was employed as an extra index of growth
limitation for all other treatments. Bacterial production estimates
were derived from TdR incorporation rates and estimates of mean cell
volumes. Biomass was determined as cell abundance × mean cell
volume × 120 fg of C µm3) (39).
Data analysis.
Duplicate incubation samples were analyzed
for each treatment. Data were analyzed statistically using multivariate
analysis of variance. Statistically significant results were analyzed
using Student-Newman-Kuels (SNK) multiple comparison tests, with
statistical significance determined at a P value of <0.05
(59). For these experiments, SNK tests were used to
distinguish differences between treatments and significant differences
over time for the variables cell abundance, cell volume, rate of
bacterial production, and specific growth rate (production rate divided
by standing biomass [P/B]).
The rates of change of various properties were estimated using
regression coefficients from model I least-squares fits on
the natural
logarithms of the individual data versus incubation
time. Growth rates
were derived by several methods. Calculation
of P/B for given time
points yielded estimates of the intrinsic
specific growth rates
(day
1), taking into account changes in both abundance and
biovolume.
Net accumulation rates were determined from the rate of
increase
of cell abundance over time, providing an indication of
population
growth based on cell division. Growth rates computed from
changes
in total biovolume used the rate of increase in the natural
logarithm
of (cell abundance × mean cell volume), which accounts
for increases
in cell size and cell division. Regressions were
performed over
appropriate intervals following inspection of the
experimental
time course plots. These net rates reflect the
intrinsic growth
rate as reduced by removal processes (e.g., grazing
and viral
lysis).
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RESULTS |
Bacterial production and distribution.
A strong zonal gradient
in surface water temperatures was observed along the cruise transect.
The most northern station had warm surface waters (14°C), while the
surface water temperature at the southern station fell to ~4°C.
Major nutrient concentrations increased from north to south. The
surface nitrate and phosphate concentrations in the subtropical zone
(STZ) were 7 and 0.6 µM, respectively, while concentrations in the
APF were 26 and 1.2 µM, respectively. Surface silicic acid
concentrations showed an opposite trend, ranging between 0.1 and 2.5 µM, with substantial depletion in the STZ and increasing in abundance
in the APF (53; Boyd et al., submitted). Dissolved
iron concentrations were subnanomolar (0.05 to 0.70 nM) within the
entire study region and generally decreased along a southerly gradient
(53; Boyd et al., submitted). The phytoplankton
community structure changed dramatically along the meridional transect;
waters in the STZ were dominated by cyanobacteria, with a shift to
eukaryotic dominance in the SAZ and APF (Boyd et al., submitted).
The rates of bacterial production in the upper mixed layer showed a
north-south gradient, declining nearly an order of magnitude
between
the STF and the APF (0.3 to 0.03 µg of C liter
1
day
1). The bacterial abundance in the upper mixed layer
displayed
a similar zonal gradient, with a greater cell abundance at
the
northern end of the transect and a decrease towards the
APF.
Responses to potential growth-limiting substances.
The major
result of the four addition experiments performed in this study was
that labile DOM stimulated rates of bacterial growth and enhanced
biomass production (Table 1). Bacteria responded favorably to
both glucose and amino acids, indicating that bacterial growth may be
constrained by both organic carbon and nitrogen. Additions of dissolved
iron or ammonium and phosphate alone had no significant impact on rates
of bacterial growth or biomass production in any experiment. The
two experiments in the SAZ and SAF indicated that bacterial growth
responded to combined additions of glucose plus Fe. In two of three
cases, bacterial growth showed dependence on the type of DOM substrate
(glucose versus DFAA) used in the growth media (see below). Results
were time dependent, with relative responses to each substrate
differing during some time courses.
(i) Experiment in the STF (42°S, 141°E).
The addition of
glucose and combinations of glucose with ammonium, phosphate, and/or
iron all significantly stimulated bacterial growth rates and the
production of bacterial biomass in seawater collected from the STF
(Fig. 2; Table
2). By day 4, all samples receiving
glucose treatments displayed elevated cell abundance relative to the
controls (Fig. 2A). All samples receiving glucose showed significant
increases in bacterial cell volume (micrometers3
cell1) (Fig. 2B) and rates of bacterial production (Table
2). Rates of Leu incorporation in the cells given glucose plus
NH4+ plus PO43
treatment were ~25 times greater than those in cells given control treatments (Fig. 2D). Additions of glucose plus
NH4+ plus PO43 and
glucose plus NH4+ plus
PO43 plus Fe dramatically increased rates of
TdR incorporation per cell after 2 days (Fig. 2E). Overall, additions
of NH4+ plus PO43
with glucose resulted in greater enhancement of cell growth rates than
did glucose treatment without NH4+ plus
PO43. Specific growth rates (P/B) for the
control, Fe, and NH4+ plus
PO43 treatments were statistically
indistinguishable throughout the experiment.
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FIG. 2.
Responses of various bacterial properties to enrichment
treatments in the STF (42°S, 141°E). (A) Cell abundance; (B) mean
cell volume; (C) TdR incorporation; (D) Leu incorporation; (E) TdR
incorporation per cell; (F) Leu incorporation per cell. Open symbols,
organic amendments; closed symbols, inorganic amendments and unamended
controls. Error bars are standard errors for duplicate samples. Letters
represent results of SNK multiple-comparison tests at the final time
point. Results for treatments with the same letter are statistically
indistinguishable (P > 0.05).
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(ii) Experiment in the SAZ (47°S, 141°E).
DFAA additions
significantly stimulated rates of bacterial growth and enhanced
bacterial biomass in amendment experiments performed in the SAZ (Fig.
3). Additions of DFAA and DFAA plus Fe
produced roughly ten times more biomass than all other treatments, including glucose (Table 3). The addition
of DFAA and DFAA plus Fe resulted in cell volumes
(micrometers3 cell1) approximately eight
times greater than those with all other treatments by day 3 (Fig. 3B).
DFAA and DFAA plus Fe additions resulted in rates of TdR incorporation
~45 times higher than in situ rates measured at day 0. Rates of
isotope incorporation (picomoles liter1
day1) with the glucose and glucose plus Fe additions
increased slightly by days 4 and 5 but never attained the levels
observed with the DFAA additions. The addition of glucose alone did not
affect cell growth rates throughout the experiment, but the addition of
glucose plus Fe increased growth rates more than five times above the control value by day 5. Samples receiving glucose plus Fe also showed
significantly greater rates of TdR and Leu incorporation than those
receiving glucose alone (Fig. 3C and D). Combined additions of Fe and
DFAA had no measurable effect on growth rates relative to additions of
DFAA alone.
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FIG. 3.
Responses of various bacterial properties to enrichment
treatments in the SAZ (47°S, 141°E). (A) Cell abundance; (B) mean
cell volume; (C) TdR incorporation; (D) Leu incorporation; (E) TdR
incorporation per cell; (F) Leu incorporation per cell. Open symbols,
organic amendments; closed symbols, iron-only amendments or unamended
controls. Error bars are standard errors for duplicate samples. Letter
designations are the same as for Fig. 2.
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(iii) Experiment in the SAF (51°S, 142°E).
Bacterial
growth in the SAF increased in response to treatments with both amino
acids and glucose (Fig. 4; Table
4). Bacterial cell abundance and volume
increased with both glucose treatments and amino acid treatments;
however, the amino acid treatments produced larger cells than the
glucose treatments (Fig. 4A and B). There were no significant
differences in abundance, volume, biomass, production, or growth rates
between Fe and control treatments (Table 4). DFAA and DFAA plus Fe
treatments produced about three times more biomass than the glucose
treatment and about twice the biomass of the glucose plus Fe treatment
(Table 4). The glucose plus Fe addition resulted in nearly twice as
much as biomass as the glucose-alone addition (Table 4). Additions of
both glucose and amino acids stimulated rates of bacterial production
in the SAF (Table 4). The glucose plus Fe treatment increased rates of
TdR and Leu incorporation significantly above those with the glucose
treatment (Fig. 4C and D). DFAA and glucose treatments resulted in more
than a 100-fold increase in TdR uptake per cell relative to control and
Fe treatments (Fig. 4E). DFAA additions provoked large increases in
rates of TdR incorporation per cell through the first 3 days of the
incubation (Fig. 4E), while glucose treatments significantly increased
thymidine incorporation through all 4 days of the incubation. Bacterial
growth was stimulated most dramatically by additions of amino acids and
by the treatments containing glucose plus Fe (Fig. 4; Table 4).
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FIG. 4.
Responses of various bacterial properties to enrichment
treatments in the SAF (51°S, 142°E). (A) Cell abundance; (B) mean
cell volume; (C) TdR incorporation; (D) Leu incorporation; (E) TdR
incorporation per cell; (F) Leu incorporation per cell. Symbols are
same as in Fig. 3. Error bars are standard errors for duplicate
samples. Letter designations are the same as for Fig. 2.
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(iv) Experiment in the APF (54°S, 141°E).
The most
striking feature of the experiment conducted in the APF (54°S) was
the lack of bacterial response to any of the treatments (Fig.
5). Bacterial abundance, cell volume, and
rates of isotope incorporation all increased slightly with every
treatment over time, but none of the measured cell properties changed
significantly relative to one another. The initial biomass at day 0 was
lower than at any other station by a factor of three, while in situ rates of TdR and Leu incorporation were nearly the same as those observed at the SAF (Fig. 4C and D and 5C and D). TdR incorporation ranged between 0.01 × 106 and 0.03 × 106 pmol cell1 day1 for all
treatments over all time points (Fig. 5E).
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FIG. 5.
Responses of various bacterial properties to enrichment
treatments in the APF (54°S, 141°E). (A) Cell abundance; (B) mean
cell volume; (C) TdR incorporation; (D) Leu incorporation; (E) TdR
incorporation per cell; (F) Leu incorporation per cell. Symbols are the
same as in Fig. 3. Error bars are standard errors for duplicate
samples. Letter designations are the same as for Fig. 2.
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DISCUSSION |
Predominance of DOM stimulation of bacterial growth.
Based on
these experiments, rates of bacterial growth in the Southern Ocean
appeared to be controlled primarily by glucose or DFAA availability.
Dissolved iron may have been important in regulating the complete
response to labile DOM, but the primary limitation to bacterial growth
was availability of labile organic substrates. These experiments also
indicate that bacterial growth rates were most severely constrained by
supplies of substrates that contain reduced nitrogen.
The most pronounced trend in three of the four studies was that the
addition of DOM, as either glucose or amino acids, enhanced
rates of
bacterial growth and resulted in significant increases
in bacterial
abundance, production, and biomass. Additions of
either glucose or
amino acids increased production rates and resulted
in the accumulation
of larger bacterial cells than control treatments.
Nine of the 16 treatments with DOM additions showed significant
increases in bacterial
abundance and biomass yields (Table
1).
Fourteen out of the 16 treatments with either glucose or DFAA
showed significantly increased
net accumulation rates (abundance)
relative to the control and Fe
treatments (Table
1). Mean rates
of TdR incorporation per cell for the
glucose and DFAA treatments
were ~300% higher than those
for control treatments in three of
the four experiments. Finally, large
increases in specific growth
rates (P/B) were seen with nearly all of
the DOM additions (Fig.
6).
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FIG. 6.
Specific growth rates (P/B) at day 4 for experimental
treatments. In situ rates are estimated at day 0 from control
treatments. Error bars represent standard errors for duplicate
treatments. Results with similar bar fill patterns are statistically
indistinguishable (P > 0.05) for each experiment.
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Patterns of response to DOM.
Patterns of bacterial growth
changed in response to the types of organic substrates provided in
these experimental treatments. Treatments with dissolved organic
nitrogen (DFAA) or glucose plus NH4+ plus
PO43 stimulated rates of bacterial growth and
production of biomass to a greater extent than additions of glucose
alone. Six of the eight treatments with either DFAA or glucose plus
NH4+ plus PO43
resulted in larger maximal specific growth rates (P/B) and produced between 2 and 25 times more biomass than glucose and glucose plus Fe
treatments. Mean bacterial biomass, production, and growth rates were
greater for treatments with DFAA or glucose plus
NH4+ plus PO43 than
for treatments with glucose alone (Tables 2 to 4). In two of the four
experiments, maximal rates of TdR incorporation occurred with either
DFAA or glucose plus NH4+ plus
PO43 (Fig. 2C and 3C).
One explanation for the observed bacterial growth response may be that
bacterial growth in the Southern Ocean is limited by
the availability
of reduced nitrogen. Increased bacterial growth
efficiency (BGE) on
DFAA or glucose plus NH
4+ plus
PO
43 substrates may have spurred the
increases in growth rates with
these treatments relative to treatments
with organic carbon alone.
Amino acids can be assimilated into cellular
constituents (proteins
and nucleic acids), providing cellular energy
conservation by
circumventing the cell's need to expend energy in
construction
of amino acids from simple carbon, nitrogen, and
phosphorus substrates
(
11,
34,
44).
Reduced nitrogen substrates may have triggered substantial increases in
BGE, resulting in large increases in growth rates.
Recent efforts to
measure BGE with bacteria growing on naturally
occurring substrates
indicate a range of 4 to 40% (
6,
16).
BGE is dependent in
part on the quality of the substrate used
to support bacterial growth.
Goldman et al. (
25) and Goldman
and Dennett (
24)
observed no differences in BGE in laboratory
enrichment cultures grown
on several substrates, including DFAA
or glucose plus
NH
4+. They found that the BGE ranged between
~40 and 95%, with lower
conversion efficiency on substrates with
higher C/N ratios. Carlson
and Ducklow (
8) estimated BGE in
amendment experiments in the
Sargasso Sea and found a range of 4 to
30% for DFAA and glucose
treatments. Kirchman (
34)
estimated a BGE of 34% for bacteria
grown on DFAA in the subarctic
Pacific.
In the present study, four of the eight treatments enriched in reduced
nitrogen resulted in higher rates of bacterial production
than glucose
additions. Mean biomass yields and growth rates with
the DFAA and
glucose + NH
4+ additions were consistently
larger than those with glucose additions
alone (Table
1). These results
hint that bacterial growth on
glucose alone may have been less
efficient than growth on organic
amendments containing reduced
nitrogen. Carbon-rich substrates
such as glucose provide energy for
cellular maintenance but may
not provide essential nutrients needed to
facilitate growth. Additions
of reduced nitrogen (as either DFAA or
NH
4+) may have alleviated possible nitrogen
limitation and increased
BGE.
Similar to Cherrier et al. (
11) in the eastern north Pacific
and Kirchman (
34) in the subarctic Pacific, we observed that
combined additions of organic carbon with reduced nitrogen stimulated
bacterial growth rates. However, we also observed that additions
of
glucose and glucose plus Fe frequently resulted in large increases
in
biomass. Clearly, bacterial growth in the Southern Ocean is
not limited
simply by organic carbon or by nitrogen. Labile organic
matter appeared
to be the primary control over bacterial growth
rates in the Southern
Ocean, with three of the four experiments
showing specific limitation
by reduced nitrogen containing
DOM.
The major difference between this study in the HNLC region of the
Southern Ocean and similar studies in the HNLC subarctic
Pacific
(
34,
37) and equatorial Pacific (
33,
36) was that
we observed large increases in bacterial biomass in response to
treatments with DFAA and glucose plus NH
4+. In
the north Pacific, where water temperatures are comparable
to
temperatures in this study, additions of DFAA and glucose plus
NH
4+ resulted in large increases in bacterial
production but only
small increases in bacterial abundance
(
34). Similarly, amendment
experiments in the equatorial
Pacific indicated that DFAA and
glucose plus
NH
4+ treatments stimulated rates of production
but had no effect on
bacterial abundance (
36). Removal
processes (predation and viral
lysis) apparently had sufficient
capability to respond to increases
in bacterial production in these two
systems, whereas in the Southern
Ocean, removal processes appear to be
temporally uncoupled from
bacterial
growth.
The most dramatic increases in bacterial biomass observed in this
experiment were in response to organic additions containing
reduced
nitrogen. In two of the four experiments, glucose and
glucose plus Fe
treatments also significantly increased cell abundance
over that with
the control treatment, but the response was lower
than that with
treatments containing reduced nitrogen. Furthermore,
DFAA and glucose
plus NH
4+ plus PO
43
treatments frequently led to production of larger cells. Studies
in
other HNLC regions found that bacterial growth rates and rates
of
production are a function of DOM, while biomass is tightly
constrained
by removal processes (
11,
34,
36). However,
in the Southern
Ocean, both biomass production and growth rates
appear to be functions
of the specific types of organic substrates
that support bacterial
growth.
Differences between the apparent coupling between DOM and bacterial
biomass in the Southern Ocean and other HNLC oceans may
be a reflection
of fundamental differences in the structures of
the microbial food webs
among the different HNLC systems. Kirchman
(
34) and Kirchman
and Rich (
36) observed a close coupling
between bacterial
growth and removal in the subarctic and equatorial
Pacific despite
additions of labile DOM. Although our experiments
were manipulative and
may not accurately model in situ processes,
they provide evidence that
increased fluxes of labile DOM in the
Southern Ocean might result in a
temporal uncoupling between bacterial
growth and removal
processes.
Colimitations: DOM and iron.
Another important difference
between this study and other studies in the subarctic and equatorial
Pacific is that the studies conducted in the subarctic and equatorial
Pacific did not investigate the potential role of dissolved iron in
regulating bacterial growth, and the resulting experimental treatments
should be considered iron replete. The work described in this study was
done under trace-metal-clean conditions, providing insight into
possible limitation by both iron and organic material.
Additions of iron alone did not significantly increase bacterial growth
rates above those with the control treatments in any
of these
experiments, unlike the results of Pakulski et al. (
43)
from
another part of the Southern Ocean. However, a combined addition
of
glucose and iron frequently resulted in higher growth rates,
rates of
production, and accumulation of biomass than control
treatments. In
three of the four experiments, glucose alleviated
growth limitation,
while in two of the four experiments, the combined
additions of
dissolved iron and glucose resulted in larger increases
in bacterial
biomass than additions of glucose alone. Iron appeared
to have a less
important effect on bacterial growth with DFAA
treatments. The
bacterial growth responses to DFAA plus Fe and
glucose plus Fe
treatments suggest that BGEs in the DFAA treatments
were already near
maximal levels, and the addition of iron had
no measurable impact on
bacterial growth. In laboratory cultures,
Tortell et al.
(
58) showed that BGE could be limited by the
availability of
iron. In iron-limited systems such as the HNLC
Southern Ocean,
bacterial cells may lack iron necessary for optimal
functioning of the
electron transport chain, reducing the total
energy yield
produced by organic matter oxidation. Additions of
iron to glucose
treatments may have increased the conversion efficiency
of glucose,
resulting in increased biomass production
rates.
Our experiments indicate that bacterial growth in the iron-limited HNLC
Southern Ocean is partly constrained by the availability
of reduced
nitrogen. One possible pathway through which iron and
reduced nitrogen
interact is assimilatory nitrate reduction. The
cytoplasmic enzymes
catalyzing the reduction of nitrate to ammonium
require iron. Bacterial
utilization of nitrate is generally low
due to the high energetic costs
associated with the reduction
of nitrate to ammonium (
36).
We did not observe any response
to experimental treatments amended with
only iron, indicating
that direct iron inhibition of bacterially
mediated nitrate reduction
was not the primary factor controlling rates
of bacterial growth.
However, our results indicate that bacterial
growth is at least
partly constrained by the availability of labile
organic matter.
This suggests that if nitrate reductase is inhibited by
low in
situ iron concentrations, its expression may be linked to the
availability of an energy source such as labile
DOM.
The possible inhibition of nitrate reductase by low iron availability
could also affect the phytoplankton community and thus
indirectly
control bacterial growth. Phytoplankton growth on nitrate
as a nitrogen
source requires nitrate reductase. One possible
implication of
iron-inhibited nitrate reductase activity might
be a decrease in inputs
of reduced-nitrogen-containing DOM from
phytoplankton, thus indirectly
limiting bacterial growth. Various
investigations into the factors
limiting bacterial growth in other
HNLC oceans support our observations
that bacterial growth may
be largely constrained by
reduced-nitrogen-containing DOM (
34,
35,
37).
Bacterial growth in the APF showed no stimulation by organic
enrichments or dissolved iron, indicating that some factor other
than
DOM exerted specific control over rates of bacterial biomass
production. It is possible that treatments in this experiment
were
contaminated with dissolved iron. Boyd et al. (submitted)
noted
significant iron contamination of water obtained at the
same time as
water for our experiment in the APF. As a result
of this potential
contamination, no firm conclusions about the
role of Fe alone as a
limiting factor may be drawn from this experiment;
however, potential
iron contamination does not detract from the
striking lack of bacterial
response to organic amendments observed
in this experiment. In situ
cell abundance and incorporation rates
were lower at the APF than at
any other station (Fig.
5). Furthermore,
temperatures in the APF were
~4°C cooler than those in the SAF.
Kirchman and Rich
(
36) noted that the time scale of the bacterial
response to
increased DOM concentrations was longer at lower temperatures
in the
equatorial Pacific during normal conditions relative to
El Niño
conditions, when the temperature was nearly 5°C higher.
It is
possible that the 5-day incubation period used in this study
was
insufficient to observe a significant bacterial response to
the
amendments.
Pomeroy and Deibel (
45) observed significant suppression of
microbial metabolism at cold temperatures (
1.8 to +1°C). Pomeroy
et
al. (
46) and Wiebe et al. (
61) concluded that an
interaction
between temperature and organic substrate concentrations
might
partly constrain bacterial growth rates. Both investigations
found
that bacterial growth rates were sensitive to manipulations of
both temperature and substrate, providing evidence that when bacteria
are grown at the low end of their temperature range, they may
require
higher substrate concentrations to maintain cellular activity
(
45,
61). The APF marks the confluence of warm subantarctic
water and
cold polar water. Bacterioplankton sampled in the APF
may be adapted to
the higher water temperatures characteristic
of the subantarctic waters
(~8°C). Those organisms adapted to
warmer water would show optimal
rates of growth at temperatures
greater than in situ temperatures found
in the APF (~4°C). Bacterial
growth in the APF may have been partly
constrained by the low
water temperatures observed in the APF. Past
investigations into
bacterial dynamics in the pelagic Southern Ocean
indicated that
metabolic activity and biomass tended to decrease south
of the
STF, increasing again south of the APF (
26,
27).
We hypothesize that no single factor limited rates of bacterial growth
in the Southern Ocean. Rather, a combination of factors,
including
carbon, nitrogen, temperature, and iron, combined to
restrict the
growth of heterotrophic bacteria. Growth-limiting
factors may stem from
an array of resource limitations across
several trophic levels whose
combined effect is to reduce in situ
bacterial growth rates. Overall,
bacterial growth in the HNLC
Southern Ocean appeared to be limited by
inputs of dissolved organic
nitrogen and carbon. However, bacterial
growth in the APF may
be more tightly constrained by low temperatures
than by resources.
In the SAF and SAZ, additions of carbon, nitrogen,
and iron may
all interact to control bacterial stocks and rates of
bacterial
growth.
| |
ACKNOWLEDGMENTS |
We gratefully acknowledge Peter Sedwick and Tom Trull (University
of Tasmania) for providing the opportunity to conduct this work. We are
grateful to the officers and crew of the R.V. Aurora Australis and the Australian Antarctic Division of CSIRO. Philip Boyd (University of Otago) provided equipment and provocative insights
into the ecology of HNLC oceans. Flynn Cunningham provided technical
assistance in the image analysis part of this work. Scott Polk provided
the study site map, and Jessica Morgan aided with statistical analyses.
This work was supported by NSF grants OPP 95-30734 to H.W.D. and INT
9802132 and OCE 9730334 to D.A.H.
| |
FOOTNOTES |
*
Corresponding author. Mailing address: The College of
William and Mary, School of Marine Science, P.O. Box 1346, Gloucester Point, VA 23062-1346. Phone: (804) 684-7401. Fax: (804) 684-7399. E-mail: mattc{at}vims.edu.
U.S. J.G.O.F.S. contribution number 545. V.I.M.S. contribution number 2277.
| |
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Applied and Environmental Microbiology, February 2000, p. 455-466, Vol. 66, No. 2
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Abstract
Introduction
