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Applied and Environmental Microbiology, September 1998, p. 3352-3358, Vol. 64, No. 9
0099-2240/98/$04.00+0
Copyright © 1998, American Society for Microbiology. All rights reserved.
Direct Determination of Carbon and Nitrogen
Contents of Natural Bacterial Assemblages in Marine
Environments
Rumi
Fukuda,*
Hiroshi
Ogawa,
Toshi
Nagata, and
Isao
Koike
Ocean Research Institute, University of
Tokyo, Nakano, Tokyo 164-8639, Japan
Received 30 January 1998/Accepted 17 June 1998
 |
ABSTRACT |
In order to better estimate bacterial biomass in marine
environments, we developed a novel technique for direct measurement of
carbon and nitrogen contents of natural bacterial assemblages. Bacterial cells were separated from phytoplankton and detritus with
glass fiber and membrane filters (pore size, 0.8 µm) and then
concentrated by tangential flow filtration. The concentrate was used
for the determination of amounts of organic carbon and nitrogen by a
high-temperature catalytic oxidation method, and after it was stained
with 4',6-diamidino-2-phenylindole, cell abundance was determined by
epifluorescence microscopy. We found that the average contents of
carbon and nitrogen for oceanic bacterial assemblages were 12.4 ± 6.3 and 2.1 ± 1.1 fg cell
1 (mean ± standard
deviation; n = 6), respectively. Corresponding values
for coastal bacterial assemblages were 30.2 ± 12.3 fg of C
cell1 and 5.8 ± 1.5 fg of N cell1
(n = 5), significantly higher than those for oceanic
bacteria (two-tailed Student's t test; P < 0.03). There was no significant difference (P > 0.2) in the bacterial C:N ratio (atom atom1) between
oceanic (6.8 ± 1.2) and coastal (5.9 ± 1.1) assemblages. Our estimates support the previous proposition that bacteria contribute substantially to total biomass in marine environments, but they also
suggest that the use of a single conversion factor for diverse marine
environments can lead to large errors in assessing the role of bacteria
in food webs and biogeochemical cycles. The use of a factor, 20 fg of C
cell1, which has been widely adopted in recent studies
may result in the overestimation (by as much as 330%) of bacterial
biomass in open oceans and in the underestimation (by as much as 40%)
of bacterial biomass in coastal environments.
| |
INTRODUCTION |
Because bacterioplankton play
important roles in the cycling of carbon and nitrogen within marine
environments (1, 14, 16), it is essential to assess
bacterial biomass accurately in order to better understand food webs
and biogeochemical fluxes. Previous studies have suggested that the
carbon biomass of bacteria generally exceeds that of phytoplankton in
oligotrophic oceans (11, 16, 20). These studies have
estimated bacterial biomass based on the assumption that one marine
bacterial cell contains 20 fg of carbon. This value was determined with
coastal bacterial assemblages grown in filtered seawater
(29) and has been commonly applied to oceanic bacterial
assemblages without much critical confirmation (11, 16, 25,
31). Recently, Christian and Karl (12) examined the
biomass distribution of microbial communities in the subtropical
Pacific Ocean by using biomass indicators of microorganisms and a
least-squares inverse method. They suggest that the average carbon
content of bacteria in the investigated area could be close to 10 fg
per cell, a value which is much lower than commonly used factors.
However, carbon contents of natural populations of oceanic
bacterioplankton have yet to be determined directly.
Previous studies have determined conversion factors by dividing
bacterial carbon by bacterial biovolume (3, 5, 18). Bacterial carbon is measured as particulate organic carbon retained on
a glass fiber filter, and biovolume is estimated by measuring bacterial
size by epifluorescence microscopy (7, 29, 33, 41), electron
microscopy (7, 30, 43), or particle counting (26). Bacterial assemblages have commonly been preincubated to minimize the effects of detritus and phytoplankton (34). The reported carbon-to-biovolume factors of marine bacteria vary widely, ranging from 0.18 to 1.61 pg of C µm3 (6,
26, 28, 29); some of these values are unrealistically high
(4). The variability may be related to errors in size measurement (28-30, 34), differences in the taxonomic
compositions and physiological states of bacterial assemblages
(28), or both. This large variability should introduce
significant errors into estimates of bacterial biomass calculated from
cell number, average cell size, and carbon-to-biovolume ratio. In
addition, preincubation before measurement could change the
compositions of bacterial populations (15, 22).
In this paper, we report our technique for estimating the carbon and
nitrogen contents of natural bacterial assemblages in oceanic and
coastal environments. Natural bacterial populations, separated from
phytoplankton and detritus, were concentrated by tangential flow
filtration. Carbon and nitrogen contents of bacterial cells were
determined by the high-temperature catalytic oxidation (HTCO) method.
Our novel approach made possible, for the first time, the direct
determination of the carbon and nitrogen contents of natural bacterial
assemblages in oligotrophic marine environments.
| |
MATERIALS AND METHODS |
Sampling.
Oceanic samples were collected in the Southern and
Pacific Oceans during a cruise (leg III to IV of KH-94-4, 10 January to 14 February 1995) of the R. V. Hakuho-maru (see Table
1). In the Southern Ocean (south), the water sample was collected at a
depth of 40 m with a Niskin sampler (volume, 12 liters). In the
other oceans a diaflame pump was used to collect water samples, while
the ship was steaming. The inlet of the pump was at about 4 m
below the surface. Surface coastal samples were collected from the
Tokyo Bay and the Otsuchi Bay with a Van Dorn water sampler (see Table
1). The vertical variability of bacterial carbon and nitrogen contents
was not examined in this study.
Prefiltration and concentration procedures.
We tested
several combinations of prefiltration filters for removal of
phytoplankton and other large particles from the sample waters. In the
prefiltration process, ca. 100 to 600 liters of samples were filtered
through 30-cm2 glass fiber filters (GA-200 for the Tokyo
Bay and GF-75 for all other stations; TOYO ROSHI Corp., Tokyo, Japan)
by gravity. Although the nominal pore sizes provided by the
manufacturer for the GA-200 and GF-75 filters are 0.8 and 0.3 µm,
respectively, a significant portion of coccoid cyanobacteria passed
through these glass fiber filters, except for the Otsuchi Bay samples.
Therefore, the oceanic samples and Tokyo Bay samples were further
filtered through a cartridge membrane filter (Micropore 8AU, type BS;
nominal pore size, 0.8 µm; size, 7 by 25 cm; Organo Corp., Tokyo,
Japan).
Particles in the prefiltered seawater were concentrated to
approximately 100 ml in volume with a Pellicon tangential flow filtration system (Millipore Corp., Bedford, Mass.) with 0.465 m2 of a 0.1-µm-pore-size Durapore membrane (hydrophilic
polyvinylidene difluoride; Millipore Corp.) attached (see Table 2). For
the Tokyo Bay sample, we used a hollow-fiber cartridge filtration system to which 0.03 m2 of a 0.1-µm-pore-size PM membrane
(polysulfone; Amicon, Inc., Beverly, Mass.) was attached. These systems
were run by a peristaltic pump with filter velocities of ca. 500 ml
min1 and a pressure of less than 1 kg cm2.
After a concentrate was recovered, a small volume of sodium chloride
solution (ca. 200 ml) was flushed through the system to wash out
bacteria adsorbed to the membrane. This solution was prepared with
precombusted sodium chloride and Milli-Q water, and the salinity was
adjusted to that of the samples. We repeated this washout procedure
four to five times with a total volume of approximately 1 liter and
combined all of the concentrated suspensions. Oceanic-sample
suspensions were further concentrated to less than 100 ml by using a
cell stir filtration system (model 202; Amicon, Inc.) with a
0.1-µm-pore-size Durapore membrane. The system was washed with
approximately 200 ml of sodium chloride solution to recover the
bacterial concentrate. The Otsuchi Bay samples obtained with a Pellicon
system were further concentrated by centrifugation (at 15,000 × g or 21,000 × g for 1 h) after samples
were weakly sonicated for 1 to 3 min in a sonication bath (IC 92;
Branson Ultrasonics Corp., Danbury, Conn.) to disperse bacterial
aggregates and detrital particles (44).
Enumeration of phytoplankton and bacterial abundance.
Chlorophyll a was extracted with
N,N-dimethylformamide after suspended materials
in the samples were collected on Whatman GF/F filters (25 mm) and
measured with a fluorometer (model 10R or 10-AU-005; Turner Designs,
Sunnyvale, Calif.) (40). Ten-milliliter volumes of bacterial
subsamples were preserved with buffered formalin (2%, vol/vol) and
stored at 4°C in the dark until the preparation of microscope slides
(within 1 week). Cells were stained with 4',6-diamidino-2-phenylindole,
filtered on Irgalan black-stained 0.2-µm-pore-size Nuclepore filters
(Costar, Cambridge, Mass.), and counted by epifluorescence microscopy
(Axioplan; Zeiss) (36). At least 240 cells were counted for
each filter. The precisions for estimates of bacterial abundance were
generally within 15% (coefficient of variation [CV]). Cyanobacteria
were counted by epifluorescence microscopy using blue excitation.
Counts were repeated 25 times for each filter, and total counts of 0 to
1,860 cells per filter were achieved.
Measurement of organic carbon and nitrogen.
Amounts of
organic carbon and nitrogen in the concentrated liquid samples were
determined by the HTCO method. This method has advantages over the more
conventional method of measuring bacterial carbon and nitrogen with an
elemental (CHN) analyzer after collecting particles on glass fiber
filters (29). First, we can avoid the loss of bacteria that
pass through glass fiber filters. Second, the HTCO method is more
sensitive and requires less sample by volume (100 µl for an analysis)
than CHN analysis (usually >10 ml). HTCO analysis for simultaneous
measurements of organic carbon and nitrogen was conducted by using a
modified total-organic-carbon analyzer (model TOC-5000; Shimadzu Corp., Kyoto, Japan) equipped with an aluminum oxide catalyst with 3% platinum, to which chromium oxide was added (35). Inorganic carbon was removed from the acidified samples by sonication in a
sonication bath (for 10 min; model 14; Branson Ultrasonics Corp.) under
a flow of nitrogen gas on the surfaces of the samples (400 ml
min1). This method, instead of more conventional bubbling
(42), was used to prevent the formation of organic flocs.
Nitrogen gas was prewashed with a barium hydroxide solution to
eliminate carbon dioxide contamination. Because a portion of inorganic
carbon sometimes remained in the samples after the above procedure
(usually less than 10% of the initial amount), the concentration of
organic carbon was obtained as the difference between the total- and
inorganic-carbon measurements. Inorganic carbon was detected on a
TOC-5000 analyzer without combustion, while total carbon was detected
after combustion. The procedural blank of organic carbon and total
nitrogen was obtained by using a sodium chloride solution which had
been used for suspension of bacterial concentrates, except for several
oceanic samples, for which Milli-Q water was substituted. The samples were kept frozen at 
20°C in sealed 10-ml glass ampoules until analysis. The precision (standard deviation [SD]) of total-carbon analysis was 0.4 to 8.6 µM (CV, 0.1 to 4.6%), and that of
total-nitrogen analysis was 0.2 to 1.9 µM (CV, 0.5 to 9.8%).
To examine oxidation efficiency for measuring carbon and nitrogen
contents of bacterial samples by our system, a comparison
was made
between the TOC-5000 analyzer and an elemental analyzer
(CHN analysis;
model NA 1500 NCS; Fisons Instruments, Milan, Italy).
Two bacterial
strains (
Marinomonas communis and
Listonella
[
Vibrio]
anguillarum) and the samples from
the Tokyo Bay and the Otsuchi
Bay were used. Bacterial strains were
cultured in half-strength
Zobell 2216E medium and harvested during
stationary phase. Cultured
bacteria do not represent the
characteristics of natural assemblages,
but they were used in order to
obtain a sufficient amount of carbon
and nitrogen for CHN analysis.
After the cells were washed and
suspended in sodium chloride solution,
subsamples were kept in
ampoules for HTCO analysis (HTCO total). Other
portions of samples
(50 ml) were filtered through precombusted glass
fiber filters
(Whatman GF/F) for CHN analysis. The GF/F filtrate was
kept in
ampoules and used for HTCO analysis (HTCO filt). Carbon and
nitrogen
values determined by the HTCO method (HTCO total minus HTCO
filt)
were compared with values obtained by CHN analysis.
| |
RESULTS |
Comparison between HTCO and CHN analyses.
We compared the
amounts of bacterial carbon and nitrogen measured by the HTCO method to
those measured by CHN analysis. The relationships between the values
obtained by those two methods were described as y = (0.92 ± 0.08) × x (33 ± 20) for carbon (r2 = 0.92) and y = (0.94 ± 0.04) × x (4.1 ± 1.8) for nitrogen (r2 = 0.98), where x and y
are the amounts of carbon or nitrogen (in micromolar concentrations)
measured by CHN and HTCO, respectively (± standard error;
n = 15). For both carbon and nitrogen, the slope was
not significantly different from 1 (P > 0.05 by the two-tailed Student t test). The difference between the
y intercept and zero was not significant (for carbon,
P > 0.1) or only marginally significant (for nitrogen,
P = 0.04). Therefore, we concluded that bacterial
carbon and nitrogen contents determined by the HTCO method did not
differ from those determined by CHN analysis.
Removal of phytoplankton by prefiltration processes.
The
abundances of bacteria and phytoplankton before and after prefiltration
are summarized in Table 1. The removal of
phytoplankton was assessed by determining the chlorophyll a
concentration and the abundance of cyanobacteria. The chlorophyll
a concentration before prefiltration was 0.078 to 0.49 µg
liter1 in oceanic samples and 3.6 to 11.8 µg
liter1 in coastal samples. The bacterial abundance of
coastal samples (2.0 × 106 to 6.1 × 106 cells ml1) was 1 order of magnitude
higher than that of oceanic samples (2.9 × 105 to
8.3 × 105 cells ml1). On the other
hand, the abundance of cyanobacteria (most of which were coccoid) in
oceanic waters was very low in the Southern Ocean (south) (<0.03 × 103 cells ml1) and relatively high in the
equatorial Pacific Ocean (3.5 × 104 cells
ml1), where it was almost as high as that in the Otsuchi
Bay in June 1995 (2.3 × 104 to 3.7 × 104 cells ml1) (Table 1).
After filtration with membrane filters (Micropore 8AU), chlorophyll
a concentrations of oceanic samples were close to the
detection limit (0.001 µg liter
1) and were less than
1% of concentrations before filtration, except
for two subtropical
samples from the Pacific Ocean (Table
1).
Oceanic bacteria mostly
passed through Micropore 8AU filters (87
to 96%), whereas most
cyanobacterial cells (>90%) were retained
on these filters (Table
1).
With coastal samples, chlorophyll
a concentrations after
prefiltration were less than 5% of concentrations
before
prefiltration. The abundance of recovered bacteria in filtrates
was 70 to 80% of that before filtration in the Tokyo Bay and 1995
Otsuchi Bay
samples, whereas the abundance was about 30% of that
before filtration
in the 1996 Otsuchi Bay samples.
Concentrating bacteria by tangential flow filtration.
By using
approximately 100 to 600 liters of sample, tangential flow filtration
achieved a 7- to 200-fold concentration of natural bacterial
assemblages (Table 2). The total number
of bacteria recovered in the concentrate (retentate) was 4.5 to 84% of
the initial number. The numbers of bacteria recovered in the filtrate
(permeate) were 7.3 to 68% and 0.02 to 31% of the initial numbers for
oceanic and coastal samples, respectively. The total recovery
(retentate plus permeate) of bacteria after tangential flow filtration
was 13 to 96% (average, 60%) for oceanic samples and 9.3 to 83%
(average, 41%) for coastal samples. The concentrated samples, with the
exception of that from the Tokyo Bay, were further concentrated by cell
stir filtration (for the oceanic samples) or centrifugation (for the
Otsuchi Bay samples) before measurement of organic carbon and nitrogen.
The total number of bacteria finally recovered was 1 to 26% of the
total number of bacteria in the original seawater (Table 2).
In order to examine the possible contribution of phytoplankton to total
organic carbon in the concentrated samples, the amount
of phytoplankton
carbon was estimated from the chlorophyll
a concentration
and the abundance of cyanobacteria (Table
1). Here we assumed
that the
carbon-to-chlorophyll
a weight ratio is 50 and the carbon
content of cyanobacteria is 200 fg cell
1 (
9),
although the carbon content of phytoplankton may be variable
depending
on its physiological state. Since the chlorophyll
a concentrations of most oceanic samples were close to the detection
limit after prefiltration (Table
1), we did not determine chlorophyll
a concentrations in the concentrated samples. By assuming
the
same recovery level for bacteria and phytoplankton during the
concentration procedure, the contribution of phytoplankton carbon
to
total carbon in the concentrate was estimated to be minor (<11%)
for
both oceanic and coastal samples, except for the 1996 Otsuchi
Bay
samples (Table
1).
We examined if bacterium-sized detritus (submicron particles; size
range, 0.4 to 1 µm) (
27) contributed to total organic
carbon and nitrogen in the concentrates. Because previous studies
have
shown that submicron particles are less dense than bacteria
and that
these particles can be separated by centrifugation (
27),
we
compared bacterial carbon and nitrogen contents in the precipitates
with those in the samples before centrifugation, using the Otsuchi
Bay
samples. We found that 63 to 80% of bacteria in the samples
concentrated by tangential flow filtration were precipitated by
centrifugation (
n = 3). The carbon and nitrogen
contents of the
samples concentrated by tangential flow filtration were
96 to
109% of those of the centrifuged samples (
n = 3), indicating that
contamination with nonliving organic particles of
low density
was minor in the Otsuchi Bay samples.
Carbon and nitrogen contents of marine bacterial assemblages.
Values for bacterial carbon and nitrogen contents are summarized in
Table 3. These quotients were calculated
by using the concentrations of organic carbon or nitrogen as determined
by the HTCO method and the numbers of bacterial cells in the
concentrated samples. The bacterial carbon contents of oceanic samples
varied between 5.9 and 23.5 fg cell1 (average ± SD,
12.4 ± 6.3 fg of C cell1; n = 6).
The corresponding values in coastal samples ranged from 15.7 to 47.9 fg
cell1 (average ± SD, 30.2 ± 12.3 fg of C
cell1; n = 5), significantly higher than
those in oceanic samples (P < 0.03 by the two-tailed
Student t test). Estimates of nitrogen content for oceanic
samples were 1.2 to 3.9 fg cell1 (average ± SD,
2.1 ± 1.1 fg of N cell1), significantly
(P < 0.002) lower than those of coastal samples (range, 3.7 to 7.3 fg of N cell1; average ± SD,
5.8 ± 1.5 fg of N cell1). The atomic ratios of
carbon to nitrogen content (C:N ratios) in oceanic samples varied
between 5.4 and 8.3, with an average value of 6.8 (SD = 1.2;
n = 6) (Table 3). The C:N ratios for coastal bacterial
assemblages ranged from 5.0 to 7.7 (average ± SD, 5.9 ± 1.1; n = 5), and there was no significant
(P > 0.2) difference between oceanic and coastal
samples. The values for carbon and nitrogen contents and C:N ratios
obtained at short intervals in the Otsuchi Bay were quite similar to
each other (Table 3).
Accounting for error propagations, estimates bear analytical errors
(SDs) in the range of 0.4 to 8.4 fg cell
1 (CV, 7 to
38%), 0.1 to 1.9 fg cell
1 (CV, 7 to 38%), and 0.1 to
1.0 (CV, 1 to 10%) for carbon content,
nitrogen content, and C:N
ratio, respectively. There was no systematic
difference
(
P > 0.05) in the precision of these estimates between
coastal and oceanic samples.
| |
DISCUSSION |
The method described in this paper has advantages over previous
methods of estimating bacterial biomass in seawater. First, our
estimation of bacterial biomass does not rely on biovolume and the
carbon-to-biovolume ratio of bacterial cells. The validity of the
carbon-to-biovolume ratio of natural bacterial populations has not been
fully established due to the difficulty of measuring bacterial cell
size accurately (28, 34). Second, our method does not
require preincubation of seawater samples, which may result in changes
in bacterial physiology and the taxonomic compositions of the samples
during incubation (15, 22, 34). The use of the large-scale
tangential flow filtration technique (2), combined with
sensitive detection of carbon and nitrogen by HTCO analysis (35), made it possible to determine directly, for the first time, the elemental compositions of natural populations of bacteria in
oligotrophic oceans. In the following discussion, we first examine
potential problems of our method and then discuss the implications of
our results for the assessment of microbial biomass in diverse marine
environments.
Methodological problems.
Possible errors involved in our
estimation of bacterial biomass include inclusion of nonbacterial
organic matter such as phytoplankton and detritus in the concentrate.
If such organic matter were included in the concentrate, our estimates
of bacterial carbon and nitrogen content would be too high. However, in
all the samples except for those taken from the Otsuchi Bay on 17 May
1996, the contribution of phytoplankton carbon (estimated from the
chlorophyll a concentration and cyanobacterial abundance)
was minor (<15% of total organic carbon in the concentrate [Table
1]). Although we did not count the abundance of prochlorophytes, which
are dominant in the oligotrophic oceans (10), our estimation
of phytoplankton carbon from chlorophyll a probably accounts
for the contribution by prochlorophytes; prochlorophytes are generally
retained by GF/F filters (23, 37). We also examined if
bacterium-sized detritus (submicron particles) (27)
contributed to our measurement of bacterial carbon. Results from the
centrifugation experiment showed that bacteria and organic carbon were
precipitated at the same ratio (the difference was within 5%),
indicating that the contribution of submicron particles to the
measurement of total organic carbon in the concentrates was minor.
After concentration by tangential flow filtration, recovered bacteria
were 5 to 84% (average, 27%) of total bacteria in prefiltered
seawater (Table
2). This result is consistent with previous
observations
that the recovery of bacteria and picoplankton from a
large volume
of seawater (50 to 8,000 liters) by tangential flow
filtration
is typically 37 to 80% (
2,
13,
19,
21). Low
levels of
recovery would be a consequence of (i) passage of a portion
of
bacterial populations through the filters or (ii) incomplete removal
of bacterial populations which firmly attached to filters, or
both. The
first factor (passage of bacteria through filters) was
significant only
for some oceanic samples from the equatorial
and subtropical Pacific;
51 to 68% of bacteria were found in the
permeate. We hypothesize that
relatively small bacteria were abundant
in these oligotrophic waters
(
32) and that these small bacteria
passed through the
filters. If this is correct, the carbon and
nitrogen contents of
bacteria that we determined for these samples
would be too large.
Concerning the second factor (incomplete removal),
we know little about
the mechanism of recovery of bacterial populations
from tangential flow
filtration systems. In our experience, bacteria
recovered in the
retentate tend to be in the form of large aggregates,
which are
probably formed during vigorous flushing (see Materials
and Methods).
We assume that selective accumulation of specific
bacterial populations
in these large aggregates did not occur,
i.e., that bacterial recovery
was a nonselective process. Consistent
with this notion, Giovannoni et
al. (
21) observed that the taxonomic
composition of
picoplankton in the concentrate made by tangential
flow filtration did
not differ from that in original seawater.
Direct determination of carbon and nitrogen contents of natural
bacterial assemblages in marine environments.
Our estimates of
bacterial carbon content in samples collected from distant and diverse
marine ecosystems (Table 1) varied widely (5.9 to 47.9 fg of C
cell1 [Table 3]). This result suggests that the
geographical and seasonal variability of bacterial carbon content is
quite large. Notably, we found that the carbon content of oceanic
samples (average ± SD, 12.4 ± 6.3 fg cell1)
is much lower than that of coastal samples (30.2 ± 12.3 fg
cell1), even though the carbon content of oceanic
bacteria could be overestimated, as mentioned above. This pattern
probably can be explained by differences in the physiological state of
bacterial populations between oligotrophic and productive waters. In
support of this hypothesis, previous work has shown that bacteria
growing faster under rich nutritional conditions are generally larger than those under conditions of starvation (34).
Table
4 summarizes the carbon contents of
marine bacteria reported in the literature. Our estimates in coastal
environments
(15.7 to 47.9 fg of C cell
1) are within the
range previously reported at a Long Island beach
(20 fg of C
cell
1) (
29) and the Otsuchi Bay (17.3 to 53.3 fg of C cell
1) (
26). Lower values (7 to 19 fg
of C cell
1) have been obtained by X-ray microanalysis for
samples collected
in Raunefjorden and Knebel Vig (
17).
Christian and Karl (
12)
estimated indirectly, by a
least-squares inverse method, that
bacterial carbon content was close
to 10 fg of C cell
1 in the subtropical Pacific Ocean. Our
estimate of 13 fg of C
cell
1 in the subtropical Pacific
Ocean is consistent with their estimates.
Caron et al. (
9)
suggested a bacterial carbon content of 15
fg of C cell
1
in the Sargasso Sea, and our average estimate of 12.4 fg of C
cell
1 for oceanic environments is also consistent with
their value.
We found that C:N ratios of bacteria varied in the range of 5.0 to 8.3 (average ± SD, 6.4 ± 1.2; CV, 19%;
n = 11)
(Table
3).
This result is consistent with those of previous studies,
which
reported natural bacterial C:N ratios in aquatic environments
that averaged around 5 to 7 (Table
4). Although the carbon and
nitrogen
contents of bacteria in coastal samples were significantly
different
from those in oceanic samples, no difference in C:N
ratio was detected.
Some researchers concluded that the bacterial
C:N ratio was unaffected
(CV, 15 to 36%) by the substrate C:N
ratio (
7,
24,
34). Our
data support the hypothesis that
the bacterial C:N ratio is relatively
invariant in marine environments.
Ratio of carbon in bacteria to carbon in phytoplankton in marine
environments.
We estimated bacterial carbon biomass in the surface
waters of several marine environments by using the obtained carbon
content and abundance and then compared it with phytoplankton biomass (Fig. 1). Phytoplankton carbon was
estimated from the chlorophyll a concentration by assuming
carbon-to-chlorophyll a weight ratios of 20 to 100. Estimates derived by using a fixed carbon-to-chlorophyll a
weight ratio of 50 will be used in the following discussion to
facilitate comparison with other studies (11, 16, 25). Note
that, for estimating bacterial biomass, we use the carbon content (5.9 to 47.9 fg of C cell1) which was directly determined for
natural populations in each region, whereas previous studies set the
value at 20 fg of C cell1. Our estimates of bacterial
biomass were 3.9 to 8.5 µg of C liter1 in oceanic
waters and 33 to 290 µg of C liter1 in coastal waters.
The ratios of bacterial carbon biomass to phytoplankton biomass were
1.7 and 1.9 for two samples in the subtropical Pacific Ocean, the only
region where bacterial carbon exceeded phytoplankton carbon. In other
regions of the oceans, the carbon biomass ratios range from 0.2 to 0.6, which is similar to those for coastal regions (0.1 to 0.5). Cho and
Azam (11) noted that in the North Pacific Gyre, the ratio of
bacterial carbon biomass to phytoplankton biomass increased with a
decrease in chlorophyll a concentration when the chlorophyll
a concentration was <0.2 µg liter1, and the
carbon biomass ratio was as high as 8. (See also Simon and Azam
[38].) Ducklow and Carlson (16) also
reported that bacterial carbon exceeded phytoplankton carbon when the
chlorophyll a concentration was below ca. 0.1 to 1 µg
liter1. Although our estimates support the previous
propositions, i.e., that bacterial carbon biomass contributes
significantly to total biomass in the oceans (8, 11, 16, 20,
39), the degree of the predominance is not as large as previously
thought. Buck et al. (8) suggested that the dominance of
bacterial biomass should not be recognized as a general rule in oceanic
waters, and our results are consistent with their opinion. On the other hand, our results suggest that the use of a fixed factor (20 fg of C
cell1) could result in great underestimation of bacterial
biomass in productive coastal environments. Previous works (16,
39) showed a negative correlation between the logarithm of the
chlorophyll a concentration and the logarithm of the ratio
of bacterial carbon biomass to phytoplankton biomass. Our data also
yield a strong negative correlation between these variables if we
assume that a fixed factor of bacterial carbon content is applicable to
all the investigated regions (r = 0.90;
P < 0.0002; n = 11). However, if we
use our measured values for bacterial carbon content for each region,
the correlation is not significant (P > 0.05).
Therefore, the relationships between the ratios of bacterial carbon
biomass to phytoplankton biomass and productivity in marine systems
should be reevaluated by taking into account the regional variability of bacterial carbon content.
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FIG. 1.
Ratio of bacterial carbon biomass to phytoplankton
biomass in the study area. Bacterial carbon was estimated by using the
data in Table 3. Phytoplankton carbon was estimated by assuming that
the carbon-to-chlorophyll a ratio is 50. Error bars indicate
the ranges of estimates with carbon-to-chlorophyll a ratios
ranging from 20 to 100. Regions for which results are shown are
indicated either by latitudinal positions (for oceanic samples) or by
geographic names (see Table 1). Note that there are four results from
the Otsuchi Bay.
|
|
| |
ACKNOWLEDGMENTS |
We thank the captain and crew of the R.V. Hakuho-maru
and the scientists on board for help with CTD sampling. We also thank E. Tanoue for lending us the Pellicon system and Organo Corp. for
providing Micropore 8AU. We are also grateful to H. Otobe and K. Morita
for help in collecting samples in the Otsuchi Bay and to K. Ishikawa
and S. Sukisaki for help with sampling in the Tokyo Bay.
This work was supported by a grant from the Ministry of Education,
Science, Sports, and Culture of Japan (07404044).
| |
FOOTNOTES |
*
Corresponding author. Mailing address: Ocean Research
Institute, University of Tokyo, 1-15-1 Minami-dai, Nakano, Tokyo
164-8639, Japan. Phone: 81-3-5351-6457. Fax: 81-3-5351-6461. E-mail:
rfukuda{at}ori.u-tokyo.ac.jp.
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Abstract
Introduction
