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Applied and Environmental Microbiology, February 2001, p. 760-768, Vol. 67, No. 2
0099-2240/01/$04.00+0 DOI: 10.1128/AEM.67.2.760-768.2001
Copyright © 2001, American Society for Microbiology. All rights reserved.
Pseudomonas stutzeri Nitrite Reductase
Gene Abundance in Environmental Samples Measured by Real-Time
PCR
Verónica
Grüntzig,1
Stephen C.
Nold,2,
Jizhong
Zhou,2,3 and
James
M.
Tiedje1,2,4,*
Department of
Microbiology,1 Center for Microbial
Ecology,2 and Department of Crop and
Soil Sciences,4 Michigan State University,
East Lansing, Michigan 48824, and Environmental Sciences
Division, Oak Ridge National Laboratory, Oak Ridge, Tennessee
378313
Received 17 August 2000/Accepted 4 December 2000
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ABSTRACT |
We used real-time PCR to quantify the denitrifying nitrite
reductase gene (nirS), a functional gene of biogeochemical
significance. The assay was tested in vitro and applied to
environmental samples. The primer-probe set selected was specific for
nirS sequences that corresponded approximately to the
Pseudomonas stutzeri species. The assay was linear from 1 to 106 gene copies (r2 = 0.999). Variability at low gene concentrations did not allow detection
of twofold differences in gene copy number at less than 100 copies. DNA
spiking and cell-addition experiments gave predicted results,
suggesting that this assay provides an accurate measure of P. stutzeri nirS abundance in environmental samples. Although P. stutzeri abundance was high in lake sediment and
groundwater samples, we detected low or no abundance of this species in
marine sediment samples from Puget Sound (Wash.) and from the
Washington ocean margin. These results suggest that P. stutzeri may not be a dominant marine denitrifier.
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INTRODUCTION |
Denitrification is one of the
important biogeochemical processes in that it is the main sink for
fixed nitrogen (28). In agriculture, it accounts for 20 to
30% of fertilizer losses (11), and in marine environments
it is thought to account for up to 80% of the loss of the nitrogen
load to coastal areas (38). Two of its products, NO and
N2O, are involved in global warming and the destruction of
the ozone layer. Denitrification is also used in waste treatment
facilities to remove excess combined nitrogen (41).
More accurate understanding and modeling of denitrification should be
possible if the catalyst can be quantified. We have evaluated the
application of real-time PCR to the quantification of the nitrite
reductase gene in Pseudomonas stutzeri. P. stutzeri is a
denitrifier commonly isolated from both soil (12) and
marine environments (44), and it may be of general
importance in global denitrification. Nitrite reductase catalyzes a key
step in the nitrogen cycle, in that the reduction of nitrite
(NO2
) to nitric oxide (NO) converts N to a
form no longer available to most of the biota. This enzyme is found as
two different variants. One contains copper and is encoded by
nirK, while the other contains the hemes c and
d1 and is encoded by nirS
(45). nirK is found in a wider range of
physiological groups, while nirS appears to be more abundant
in nature (8). P. stutzeri contains
nirS (29).
Several methods have been used to attempt to identify or quantify
denitrifiers, including the design of specific PCR primers (6,
17) or probes (39) for genes involved in
denitrification or ribosomal DNA genes (26) and
immunofluorescence assays using polyclonal antibodies against specific
denitrifying bacteria (44) or denitrification enzymes
(8). Recently, competitive PCR and most-probable-number
PCR were applied to the quantification of nirS-containing
denitrifying bacteria (34).
The real-time PCR technique is based on the use of the 5' nuclease
assay, first described by Holland et al. (20) and further improved by the use of fluorescent TaqMan methodology and the ABI Prism
7700 sequence detection system (PE Applied Biosystems, Foster City,
Calif.) (13, 19). The system requires the design of a
forward and a reverse primer, in addition to a probe that hybridizes
between them. The probe is fluorescently labeled at both ends
(31). The fluorescent dye at the 5' end serves as a
reporter, and its emission spectra are quenched by the dye at the 3'
end of the probe. During the elongation step of each PCR cycle, the DNA
polymerase cleaves the annealed probe with its 5' nuclease activity.
Once separated from the quencher, the reporter fluorescence is
detected, resulting in an increase in fluorescence emission. The
fluorescence increases logarithmically as the PCR proceeds, until a
reagent becomes limiting. A threshold fluorescence intensity is defined
within the logarithmic phase. The higher the amount of initial template
DNA, the earlier the fluorescence will cross the defined threshold.
Copy number of the initial target DNA is thereby determined by
comparison to a standard curve.
The advantage of the real-time PCR method over other PCR-based
quantification methods is that it focuses on the logarithmic phase of
product accumulation rather than on the end product abundance. This
technique is therefore more accurate, since it is less affected by
amplification efficiency or depletion of a reagent. In addition, real-time PCR measures template abundance over a large dynamic range of
around 6 orders of magnitude (19). Finally, this method allows the simultaneous analysis of 96 samples in a short time and
reduces the risk of contamination, as no post-PCR manipulation is
required. The main disadvantage of real-time PCR is the need for a
special thermocycler and reagents that are expensive compared to the
equipment utilized by other PCR-based quantification methods.
Real-time PCR has been successfully applied in the medical field, for
example in the quantification of various DNA and RNA viruses in
patients (16, 23, 27, 33), in the detection of gene
amplification (3), mutations, or chromosomal
rearrangements (10, 32), and in the quantification of gene
expression (2, 43) or detection of various splice variants
(25). Recently, real-time PCR was applied to environmental
samples in studies that quantified conidia of a human pathogenic mold
in airborne samples (18) and for determining the abundance
of bacterioplankton in marine samples (40).
In this study, we tested real-time PCR for its use for quantifying an
important functional gene in environmental samples.
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MATERIALS AND METHODS |
Samples.
Soil samples were obtained from an agricultural
plot at the Kellogg Biological Station (KBS; Mich.) (1).
Groundwater samples were taken from the Schoolcraft (Mich.) and
Shiprock uranium mill tailings remedial action (N.Mex.) sites. The
uranium recovery process at the latter site used high concentrations of
ammonia, some of which entered the groundwater and led to the
accumulation of nitrate. The site is located on an elevated terrace,
along the south side of the San Juan River. Samples were taken on the terrace (samples 813 and 826), on the floodplain (samples 602, 603, and
619), and from a seep flowing from the base of the escarpment into the
floodplain (sample 425). The samples presented increasing concentrations of nitrate in the following order: 602, 826, 619, 425, 603, and 813 (22). P. stutzeri KC, a strain
that hydrolyzes carbon tetrachloride (9), was injected
into the Schoolcraft aquifer for a bioremediation field test 15 days
before the samples were taken from wells 2 m upstream and 1 and
2.5 m downstream from the injection site (M8, M11, and M19,
respectively) (21). The freshwater sediment sample was
collected from the surface 2 cm of sediment from Wintergreen Lake
(Mich.), a small hypereutrophic lake. The marine sediment samples were
obtained from the Washington margin of the Pacific Ocean and from Puget
Sound (Wash.). Sediment cores were sliced into sections, as described below.
Cultures.
Marine P. stutzeri isolates (strains
A3-5, D7-6, D9-1, E4-2, and F9-2) were obtained by L. Wu from the
Washington marine sediments (7). nirS clones
were obtained by G. Braker by amplification of DNA extracted from
Washington margin and Puget Sound sediments, using specific primers
(7). All culture collection strains and marine isolates
used in this study were grown in nutrient broth (Difco, Detroit,
Mich.). Escherichia coli transformants were grown in
Luria-Bertani broth (36) amended with kanamycin (50 µg/ml). Shewanella oneidensis MR-1, P. stutzeri
KC, and Pseudomonas aeruginosa were grown at 30°C, while
all other cultures were grown at 37°C.
DNA extraction and quantitation.
Genomic DNA was extracted
from late-exponential-phase cultures. Cells were harvested by
centrifugation and resuspended in lysis buffer (50 mM Tris [pH 8], 50 mM EDTA, 100 mM NaCl). The cells were incubated for 15 min at 37°C
with lysozyme (3.5 mg/ml), achromopeptidase (70 µg/ml), and RNase A
(30 µg/ml). After two freeze-thaw treatments, the cell suspension was
incubated for 5 min at 37°C with sodium dodecyl sulfate (1%),
followed by incubation with proteinase K (400 µg/ml) (1 h at 60°C).
The lysate was extracted twice with phenol-chloroform-isoamyl alcohol.
After isopropanol precipitation, the DNA was resuspended in Tris-EDTA
buffer (pH 8). Plasmid DNA was extracted from the E. coli
transformants with the Wizard Plus SV Miniprep DNA
purification system (Promega, Madison, Wis.), according to the
manufacturer's instructions. The genomic DNA extractions from the KBS
soil, the Shiprock aquifer, and the Wintergreen Lake sediment were
performed using the Ultra Clean Soil DNA kit (MO BIO, Solana Beach,
Calif.), following the manufacturer's instructions. Genomic DNA was
extracted from the Schoolcraft groundwater samples by the method of van
Elsas and Smalla (42). This method was also used to
extract genomic DNA from the Washington marine sediment samples, with
an additional proteinase K treatment (50 µl of a 20-mg/ml solution)
after the incubation with sodium dodecyl sulfate. The protocol of Gray
and Herwig (15) was used to extract genomic DNA from the
Puget Sound samples.
The quality of the extracted DNA was analyzed by electrophoresis on a
0.8% agarose gel. DNA concentrations were measured by absorbance at
260 nm. The P. stutzeri nirS gene copy number was estimated
based on the P. stutzeri Zobell genome size (4.29 Mbp) (14) and on the assumption that only 1 copy of
nirS is present per genome (24), i.e., 4.4 fg
of P. stutzeri DNA = 1 genome copy = 1 nirS copy.
Primers and probe.
Nine P. stutzeri nirS
sequences from isolates (7) and 52 non-P. stutzeri
nirS sequences from marine isolates, clones, and unrelated species
(7) were compared to select conserved regions within the
P. stutzeri nirS gene. The primers and probe were designed within conserved regions using the program PrimerExpress (PE Applied Biosystems) (Table 1). The probe was
dually labeled with the fluorescent dyes 6-carboxyfluorescein (FAM) and
6-carboxytetramethyl-rhodamine (TAMRA) at the 5' and 3' ends,
respectively, as recommended by the manufacturers. The primers and
probe were synthesized by Integrated DNA Technologies (Coralville,
Iowa).
Real-time PCR.
The increase in fluorescence emission, due to
the degradation of the probe by the DNA polymerase in each elongation
step, was monitored during PCR amplification using the 7700 Sequence Detector (PE Applied Biosystems). The fluorescence signal was normalized by dividing the emission of the reporter dye
(6-carboxyfluorescein) by the emission of the passive reference dye
6-carboxy-X-rhodamine. The parameter CT
(threshold cycle) is the fractional cycle number at which the
fluorescence emission crosses an arbitrarily defined threshold within
the logarithmic increase phase (0.1 in our reactions). The higher the
amount of initial template DNA, the earlier the fluorescence will cross
the threshold and the smaller will be the CT.
The CT values obtained for each sample were
compared with a standard curve to determine the initial copy number of
the target gene.
The reaction mixture for real-time PCR consisted of 1× TaqMan
Universal PCR Master Mix (containing AmpliTaq Gold DNA polymerase,
AmpErase uracil-
N-glycosylase, which degrades PCR carryover
products
from previous reactions, deoxynucleoside triphosphates with
dUTP,
a passive reference [6-carboxy-X-rhodamine], and optimized
buffer
components) (PE Applied Biosystems), 300 nM forward primer, 900
nM reverse primer, and 525 nM fluorogenic probe. MicroAmp optical
caps
and tubes were used (PE Applied Biosystems). A total volume
of 30 µl
was used for the optimization steps and 50 µl was used
for the final
reactions. PCR conditions were as follows: 2 min
at 50°C, 10 min at
95°C, then 40 cycles of 15 s at 95°C and 1
min at 60°C.
Negative controls with no template DNA or no probe
were run in each
reaction.
Specificity.
The DNA extracted from several strains, marine
isolates (7), and E. coli transformants
(7) was used as positive and negative controls to test the
specificity of the primer-probe set. Template DNA (18 ng) was added to
each reaction tube.
Sensitivity and detection limit.
Marine isolate E4-2 DNA was
chosen as a standard for measuring the sensitivity of the primer-probe
set and for generating standard curves in subsequent determinations.
This isolate was previously identified as P. stutzeri based
on 16S ribosomal DNA sequence identity and physiological
characteristics (7). Serial dilutions (10-fold) of
P. stutzeri E4-2 DNA were prepared in herring sperm DNA (1 µg · ml1 in water; Boehringer Mannheim,
Indianapolis, Ind.) as a carrier. All determinations were performed in
triplicate and 95% confidence intervals were determined (shown as
error bars).
Various calibration curves were constructed to determine the lower
detection limit of this assay and our ability to discriminate
twofold
differences in template concentration. A dilution series
of marine
isolate E4-2 DNA was prepared in a 1-µg · ml
1
solution of herring sperm DNA. Different volumes (2, 4, 10, and
20 µl) of template DNA from this dilution series were added to
individual reaction tubes, and the difference was made up with
water.
The maximum allowable error (MAE) in order to distinguish a twofold
difference in copy number was calculated as follows:
CT =
m · log(2), where
CT is the difference between
CT obtained
from samples with a twofold
difference in target copy number,
and
m is the slope of the
standard curve. The MAE is calculated
by this equation: MAE =
CT/2. The MAE was calculated for each
standard curve generated, and the mean MAE and the 95% confidence
interval were
determined.
Quantitation of nirS in environmental samples.
Reactions were performed using 100 ng of template DNA. The template
copy number was determined from CT values by
using a standard curve. Samples that exhibited
CT values equal to or higher than the negative
controls were considered as below the detection limit. All results were
normalized to the quantity of community DNA.
Accuracy.
In order to test for the presence of PCR
inhibitors, 106 strain E4-2 genome copies (4.4 ng of DNA)
were added to 100 ng of DNA extracted from one environmental sample
from each site. Samples with and without the addition of this positive
control DNA were compared, and the percent recovery of the added genome
copy number was calculated. P. stutzeri abundance was also
evaluated in microbial communities constructed from P. stutzeri KC, P. aeruginosa, and E. coli
JM109 in different proportions and added to 1 g of sterile quartz
sand (Sigma, St. Louis, Mo.). Direct cell counts were obtained before
mixing, using a Petroff-Hausser counting chamber. P. stutzeri KC cells were added to the mixtures in various ratios (1, 1/10, 1/102, 1/103, 1/104,
1/105, 1/107, 1/108, and 0). Equal
cell numbers of P. aeruginosa and E. coli JM109 were added to achieve 2 × 108 cells in each 300-µl
mixture. After cell addition, the mixture was vortexed for 3 s,
and DNA was extracted using the Ultra Clean Soil DNA kit (MO BIO),
following the manufacturer's instructions. Triplicate samples were
analyzed using 100 ng of template DNA. To avoid any problem from
different extraction efficiencies, the P. stutzeri KC
nirS gene copy number was expressed relative to total DNA
extracted. P. stutzeri, P. aeruginosa, and E. coli genome sizes were recently measured and reported as 4.29 Mbp
(14), 5.9 Mbp (35), and 4.6 Mbp
(4), respectively; their percent G+C content was
considered to be 63, 67, and 50%, respectively (30).
To measure the accuracy of real-time estimations of
P. stutzeri abundance in environmental samples, different numbers of
P. stutzeri cells were added to Wintergreen Lake sediment
and KBS
soil samples. The number of total cells in the soil or sediment
samples was determined by direct counts after staining with
5-(4,6-dichlorotriazine-2-yl)
aminofluorescein (
5).
Serially diluted cells (10
8, 10
7, and
10
6 cells) were added to 0.5 g of sediment or soil
samples. After
vortexing for 5 s, total DNA was extracted as
described above.
In order to normalize these values to the total
community DNA,
the average genome size of soil or sediment bacteria was
considered
equal to the
E. coli genome size, or 4.6 Mbp
(
4).
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RESULTS |
Specificity.
DNA extracted from a range of denitrifying
isolates was used to test the specificity of the primer-probe
combination (Table 2). A logarithmic
increase in fluorescence intensity was readily detected by real-time
PCR in 17 of 21 P. stutzeri strains. However, we were not
able to detect by real-time PCR the P. stutzeri strains corresponding to genomovars 4, 5, and 7 and one strain in genomovar 1. Sequence analysis of nirS of the strains from genomovars 4, 5, and 7 indicated a higher similarity (80, 81, and 82%, respectively) with the nitrite reductase gene of P. aeruginosa than with
that of P. stutzeri. We identified several mismatches
between these strains and the primer-probe combination used in this
study (five to nine mismatches for the primers and four to five for the
probe). No non-P. stutzeri strain or clone gave a real-time
PCR signal.
Sensitivity and detection limit.
We tested the sensitivity of
the real-time detection system using a dilution series of P. stutzeri DNA (Fig. 1A). The
threshold value for this and all subsequent analyses was chosen to be
0.1. This value falls within the range of logarithmic fluorescence increase, yet avoids the signal from the no-template control. Nearly
all of our no-template controls showed some increase in fluorescence
intensity, similar to that shown in Fig. 1A. Since this increase was
not logarithmic and similar signals were detected in control reactions
without DNA polymerase (data not shown), this fluorescence increase is
likely due to probe degradation. The data obtained were used to draw a
standard curve relating CT values to the added
mass of P. stutzeri DNA and the number of gene copies (Fig.
1B). A linear response was observed over more than 6 orders of
magnitude, ranging from 14 to 4.05 × 106
nirS gene copies (r2 = 0.999;
Fig. 1B).
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FIG. 1.
Generation of standard curve. (A) Increase of
fluorescence intensity with cycle number for serially diluted P. stutzeri DNA. Symbols, from left to right:  , 17.8 ng;  , 5.9 ng;  , 0.59 ng;  , 59 pg;  , 5.9 pg; , 0.59 pg;
 , 59 fg;  ,
no-template control. CT, cycle at which the fluorescence
intensity crosses an arbitrary threshold value. (B) Standard curve.
Values represent means ± 95% confidence interval (n = 3).
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We constructed a series of calibration curves to study the detection
limit of the system and our ability to differentiate
similar
P. stutzeri DNA concentrations. Different volumes (ranging
from 2 to
20 µl) of a dilution series of template DNA (ranging
from 0.1 to
1,000
nirS gene copies · µl
1) were
added to the PCR. Given the 96-well capacity, the analyses
were done in
a low- and high-concentration set (Fig.
2A and
B,
respectively). Although the curves
remained linear down to 1
nirS copy (20 µl of 0.05
nirS gene copies · µl
1), the
variability associated with
CT values from low
copy numbers
precluded our ability to distinguish concentrations in
samples
with similar amounts. Only when the copy number was 100 or
greater
were we able to reliably differentiate a twofold difference in
P. stutzeri nirS concentration.
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FIG. 2.
Limit of detection of the P. stutzeri nirS
gene using real-time PCR. The denoted volumes of serially diluted
P. stutzeri DNA were added to different reaction mixtures.
(A) Low concentrations. (B) High concentrations. Values represent
means ± 95% confidence interval (n = 3).
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The increase in variability with cycle number is apparent when the
upper 95% confidence interval from all determinations is
plotted
against
CT (Fig.
3). Based on the slope of each individual
standard curve, we calculated the maximal error in the
CT value
that would still allow detection of a
twofold difference in gene
copy number (MAE). The average of the
different MAE values is
equal to 0.53 (Fig.
3). The corresponding 95%
confidence interval
is too small to be observed (Fig.
3).
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FIG. 3.
Relationship between CT and
error. The horizontal line represents the MAE to discriminate between
samples containing a twofold difference in P. stutzeri nirS
copy number. Values shown represent the means from 14 independent
standard curves. The 95% confidence interval is shown as a dashed
line.
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Accuracy.
P. stutzeri KC, E. coli JM109,
and P. aeruginosa cells were added to sterile quartz sand in
various known amounts. P. stutzeri KC contains the targeted
nirS gene, while P. aeruginosa has a nirS that is 67% similar in nucleotide sequence. The
correlation between the calculated and measured values was extremely
high (slope = 0.98, r2 = 0.992) (Fig.
4B). The lowest proportions of P. stutzeri (1/107 and 1/108) overlapped with
the background in the real-time PCR measurements.
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FIG. 4.
Quantification of nirS DNA in
artificial mixtures of P. stutzeri KC, P. aeruginosa, and E. coli cells. (A) Comparison between
P. stutzeri nirS gene copies per µg of DNA measured values
by real-time PCR (gray bars) and calculated values based on cell counts
(black bars). (B) Correlation between calculated and measured values
(slope = 0.98, r2 = 0.992). Error bars
represent 95% confidence intervals (n = 3) for both
panels.
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To further test the accuracy of the system, a similar experiment was
performed adding known amounts of
P. stutzeri cells to
different soil and sediment samples. Measured values of
nirS
correlated
well (slope = 0.97,
r2 = 0.971) with the values calculated from direct cell counts (Fig.
5).
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FIG. 5.
Quantification of nirS in KBS soil () and
Wintergreen Lake sediment () samples spiked with P. stutzeri cells. The line shows the correlation between
P. stutzeri nirS gene copies per microgram of DNA measured
by real-time PCR and calculated values. Error bars represent 95%
confidence intervals (n = 3).
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Analyses of environmental samples.
DNA extracted from
environmental samples exhibited a wide range of P. stutzeri
nirS gene abundance, as measured by real-time PCR (Table
3). The groundwater and freshwater
sediment samples had the highest P. stutzeri nirS copy
numbers. Marine sediments consistently displayed low P. stutzeri
nirS abundance.
In order to test for the presence of any PCR inhibitors in the
environmental samples, 10
6 genome copies of strain E4-2
were added to each sample. Five
of the six samples yielded the expected
amount of
nirS (Fig.
6).
The
KBS soil sample showed slightly fewer
nirS copies than
expected.
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FIG. 6.
P. stutzeri nirS copy number measured by
real-time PCR without (black bars) and with (gray bars) the addition of
106 P. stutzeri nirS copy numbers to DNA
extracted from the following environmental samples: KBS, soil from KBS
(Mich.); SCH, groundwater from Schoolcraft bioremediation site (Mich.);
SHI, groundwater from Shiprock (N.Mex.); WIN, freshwater sediment from
Wintergreen Lake (Mich.); WAS, marine sediment from Washington margin
(Wash.); PUG, marine sediment from Puget Sound (Wash.).
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DISCUSSION |
Good detection methods share four features: specificity,
sensitivity, precision, and accuracy. Real-time PCR appears to satisfy these requirements. The primer-probe set we designed for the P. stutzeri nirS gene amplified a group of strains that generally corresponded to the P. stutzeri species (Table 2). Except
for one strain, all the representatives of the two more commonly
isolated genomovars, 1 and 2, were readily amplified. The sequence
similarity of the nirS gene of the P. stutzeri
strains in genomovars 4, 5, and 7 to the P. aeruginosa
nitrite reductase gene explains the inability to detect these strains
by real-time PCR. In addition to cultured strains, we selected a range
of isolates and cloned nirS sequences from the Pacific
Northwest marine environment for the primer-probe design. This makes
our system especially appropriate for application in this environment.
Although there may be cross-reactivity with related DNA in the natural
microbial communities, none of our habitat-specific negative controls
gave a positive reaction. Furthermore, the phylogenetically closest
relative, Pseudomonas balearica, was not detected. The probe
needed in real-time PCR requires the identification of three specific
regions in the DNA sequence, rather than two, which provides for the
high specificity. This, however, can make the design of an appropriate
primer-probe set more difficult. The 16S rRNA gene would be an
alternative target which is more conserved but that would sample a
larger organismal group, which may not correlate with function.
The method was linear over more than 6 orders of magnitude and
sensitive down to 1 gene copy, similar to the results obtained in other
studies (18, 27). However, the high variability associated with low target copies limits precision near the detection limit (Fig.
2). We are able to detect the presence of only 1 copy, but not
precisely, e.g., we cannot discriminate a twofold difference at 100 copies or less. The theoretical upper 95% confidence interval to allow
the discrimination of a twofold difference in copy number is 0.53 (Fig.
3). Confidence intervals below this value are achieved more frequently
at lower CT values, equivalent to higher numbers of target copies. The error values associated with the measurements of
P. stutzeri nirS abundance in various environmental samples were consistent with this characterization of precision, i.e., high
concentrations have an error value at least 1 order of magnitude smaller than the measured value, while the error was of the same order
of magnitude when nirS copy number was small (Table 3).
The method proved to be extremely accurate. When P. stutzeri
was mixed in various proportions with E. coli and P. aeruginosa cells, it could be detected when present at 100% to
0.001% in the mixture, presenting a high correlation between expected
and measured results over this whole range. Furthermore, spiked samples gave the expected results.
Environmental samples analyzed with real-time PCR displayed a wide
range of P. stutzeri nirS abundances. P. stutzeri
was highly abundant in freshwater sediment from Wintergreen Lake
(Mich.), which agrees with a study by Gamble et al. (12),
who identified P. stutzeri as the dominant denitrifier in
these sediments. P. stutzeri was either absent or present at
extremely low population densities in marine sediments from the
Washington margin and Puget Sound (Wash.). Ward and Cockcroft
(44) measured the abundance of P. stutzeri in
the water column of Monterey Bay, Calif., and found that it represented
only 0.02 to 0.08% of the total bacterial community. Our findings
indicate that P. stutzeri is also not abundant in marine
sediments. This is in agreement with the studies by Braker et al.
(7), who observed nirS heterogeneity and the lack of a dominant group in nirS clones isolated from these
sites. Furthermore, these results are also consistent with studies done on other denitrification genes. Scala and Kerkhof (37)
observed a high diversity among nitrous oxide reductase
(nosZ) genes in marine sediments, with no overlap between
environmental nosZ sequences and cultured denitrifiers.
Groundwater samples from the Schoolcraft bioremediation site (Mich.)
followed expected trends, corresponding with the injection of
substrates and P. stutzeri strain KC into a contamination
plume. We detected no P. stutzeri upgradient from the
inoculation site and found the highest P. stutzeri abundance just downgradient from the inoculation site. P. stutzeri was
also prevalent at Shiprock, perhaps resulting from the high nitrate contamination at that site.
Real-time PCR is a specific, sensitive, precise, and accurate method
for quantifying a gene or organism group in a broad range of
environmental sample types. It remains to be seen whether functional gene sequences are conserved enough in a habitat so that a reasonable number of probe-primer sets can provide useful quantitative information for components of a group or process. This same method and primer-probe regions should be effective in quantifying mRNA and hence in
determining which group of organisms or gene families are active in denitrification.
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ACKNOWLEDGMENTS |
This work was supported by Department of Energy grants
DE-FG02-98ER62535 and DE-FG02-97ER62469. Oak Ridge National Laboratory is managed by the University of Tennessee-Battelle LLC for the Department of Energy under contract DE-AC05-00OR22725.
We kindly thank Jorge Lalucat for providing the P. stutzeri
strains with genomovar classifications, Allan Devol for collecting the
Puget Sound and Pacific Ocean sediment samples, Phillip Long and the
UMTRA personnel for the Shiprock groundwater samples, Liyou Wu for the
marine isolates, Gesche Braker for the nirS clones and for
providing us DNA extracted from the Pacific Ocean sediments, and
Katrina Linning for DNA extracted from the Schoolcraft samples.
| |
FOOTNOTES |
*
Corresponding author. Mailing address: Center for
Microbial Ecology, 540 Plant and Soil Science Building, Michigan State
University, East Lansing, MI 48824-1325. Phone: (517) 353-9021. Fax:
(517) 353-2917. E-mail: tiedjej{at}msu.edu.
Present address: Department of Biology, University of
Wisconsin-Stout, Menomonie, WI 54751-0790.
| |
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Applied and Environmental Microbiology, February 2001, p. 760-768, Vol. 67, No. 2
0099-2240/01/$04.00+0 DOI: 10.1128/AEM.67.2.760-768.2001
Copyright © 2001, American Society for Microbiology. All rights reserved.
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Abstract
Introduction
