Lehrstuhl für Mikrobiologie, Technische
Universität München, 85350 Freising,
Germany,1 and Department of
Microbial Ecology, Institute of Biological Sciences, University of
Aarhus, 8000 Aarhus, Denmark2
Fluorescence in situ hybridization (FISH) with rRNA-targeted
oligonucleotide probes has found widespread application for analyzing the composition of microbial communities in complex environmental samples. Although bacteria can quickly be detected by FISH, a reliable
method to determine absolute numbers of FISH-stained cells in
aggregates or biofilms has, to our knowledge, never been published. In
this study we developed a semiautomated protocol to measure the
concentration of bacteria (in cells per volume) in environmental
samples by a combination of FISH, confocal laser scanning microscopy,
and digital image analysis. The quantification is based on an internal
standard, which is introduced by spiking the samples with known amounts
of Escherichia coli cells. This method was initially
tested with artificial mixtures of bacterial cultures and subsequently
used to determine the concentration of ammonia-oxidizing bacteria in a
municipal nitrifying activated sludge. The total number of ammonia
oxidizers was found to be 9.8 × 107 ± 1.9 × 107 cells ml
1. Based on this value, the
average in situ activity was calculated to be 2.3 fmol of ammonia
converted to nitrite per ammonia oxidizer cell per h. This activity is
within the previously determined range of activities measured with
ammonia oxidizer pure cultures, demonstrating the utility of this
quantification method for enumerating bacteria in samples in which
cells are not homogeneously distributed.
 |
INTRODUCTION |
Fluorescence in situ hybridization
(FISH) using rRNA-targeted oligonucleotide probes is frequently applied
to quantify the composition of microbial communities in different
environments (1, 17, 21, 23, 33, 35). In such studies cell
numbers are generally obtained by manual counting in an epifluorescence microscope. Usually the relative abundance of a probe target population is determined by comparison of the obtained numbers (i) with counts of
all bacterial cells detectable by FISH via simultaneous hybridization with a bacterial probe (10, 11, 30, 34) or probe set
(9), or (ii) with counts of all organisms containing DNA
by simultaneous application of nucleic acid staining dyes (16,
29, 35, 38).
Although quantitative FISH has provided novel insights into the
structure and dynamics of microbial communities, it suffers from
tediousness and limited accuracy for samples containing densely aggregated cells like activated sludge flocs or biofilms. The latter
problem can in part be ameliorated by the use of confocal laser
scanning microscopy (CLSM) for the detection of probe-labeled cells
(36). However, even if optical CLSM sections are recorded, it is not feasible to manually count a sufficient number of cells in
each hybridization experiment in a reasonable time period to obtain
statistically reliable results. This limitation has two reasons. First,
manual counting itself is very time-consuming, and thus generally not
more than a few thousand cells per hybridization experiment were
counted in previous studies. Second, manual counting requires
high-magnification CLSM sections, which allow single-cell resolution
within clusters. However, such images contain relatively few cells, and
therefore many images need to be recorded, rendering the procedure even
more time-consuming. Therefore, more precise methods are required to
quantify the composition of the microflora in samples containing
clustered cells.
In principle, flow cytometry is a more efficient and accurate
alternative for quantification of fluorescently labeled bacterial cells
(39). However, for the analysis of microbial flocs and biofilms, flow cytometry is of limited use because it necessitates efficient dispersion of clustered bacteria prior to the measurement, a
requirement which frequently cannot be fulfilled (38, 39).
To overcome the limitations of manual cell-counting procedures,
semiautomated digital image analysis tools were recently developed which quantify fluorescently labeled bacteria in environmental samples
(6, 29). But such solutions are not able to efficiently count cells in dense clusters or biofilms because single-cell recognition within these structures cannot be automated. This problem
can be circumvented by measuring the areas of specifically stained
bacteria in randomly acquired optical CLSM sections. This approach only
requires the software to differentiate between labeled biomass
(including cell clusters) and unlabeled background but does not rely on
single-cell recognition within clusters. The abundance of a particular
population is then expressed as fraction of the area occupied by all
bacteria (8, 32).
For this purpose, an environmental sample is hybridized simultaneously
with different rRNA-targeted oligonucleotide probes: one specific probe
that targets the population which is to be quantified, and one
domain-specific probe set that detects most bacteria. The
population-specific and the domain-specific probes are labeled with
different fluorochromes. Following FISH, the fluorescence conferred by
the different probes is recorded in separate CLSM images. The areas of
the labeled cells in these images are measured by digital image
analysis. Since this approach analyses low-magnification images and can
be partly automated, it allows rapid quantification of large numbers of
bacteria, thereby significantly improving the statistical accuracy of
the measurement (8, 32).
Ecological studies of complex microbial communities may attempt to
determine not only relative abundances of probe-defined bacterial
populations, but also the respective cell concentrations in a sample.
This is particularly important if different samples which differ in
their prokaryotic biomass content are to be compared. Furthermore, cell
concentrations per volume or weight unit of an environmental sample are
needed to calculate key functional attributes of bacterial populations,
like in situ growth rates and in situ substrate turnover rates per
cell. Despite their importance, cell concentrations of FISH-stained
bacterial populations have rarely been determined for biofilms or
activated sludge flocs by manual counting because these measurements
required additional time-consuming and bias-introducing homogenization
and membrane filtration steps (17, 24, 26, 37). In
addition, it is impossible to directly apply the above-mentioned
area-based quantification methods (8, 32) to
semiautomatically determine absolute cell numbers in a sample after
membrane filtration because these methods cannot accurately measure the
entire biovolume of all cells of a probe-labeled population in the
filtered biomass on top of defined filter areas.
Recently, CLSM-based methods to semiautomatically measure the biovolume
of fluorescently labeled bacteria (15, 19) were published.
These methods could theoretically be applied to determine absolute cell
numbers of probe-defined bacterial populations on membrane filters.
However, biovolume-based quantification is only accurate if serial
optical sections are recorded using small vertical step intervals and
subsequently combined to image stacks. This procedure is extremely
time-consuming and leads to significant bleaching of FISH-labeled
bacterial cells.
In this study we thus developed a semiautomated procedure for
determining cell concentrations of bacterial populations in complex
samples by FISH and CLSM using the area-based quantification method
(8, 32). Spiking of the samples with known amounts of
Escherichia coli cells, which were used as internal
standards for the subsequent FISH analysis, allowed us to infer the
absolute cell numbers of probe-target bacteria from their measured
areas by digital analysis of CLSM images.
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MATERIALS AND METHODS |
Test strains, culture conditions, cell fixation, and activated
sludge sampling.
Type strains of Comamonas testosteroni
(DSM 1622) and Gluconobacter asaii (DSM 7148) were obtained
from the Deutsche Sammlung von Mikroorganismen und Zellkulturen GmbH
(DSMZ) (Braunschweig, Germany). Cells of C. testosteroni and
G. asaii were grown overnight under agitation at 30°C in
medium DSM M1 (0.5% [wt/vol] peptone, 0.3% [wt/vol] meat extract,
pH 7.0) and DSM M626 (5% [wt/vol] D-sorbitol,
1% [wt/vol] yeast extract, 1% [wt/vol] peptone, pH 6.0),
respectively. Pure cultures of Nitrosomonas europaea were maintained as described by Koops et al. (18). E. coli TOP10F' cells (Invitrogen, San Diego, Calif.) were grown
overnight in Luria-Bertani (LB) medium (31) at 37°C
under agitation. For fixation, cells of all species were washed in
phosphate-buffered saline (PBS) and then incubated for 3 h in 3%
paraformaldehyde (Sigma, Deisenhofen, Germany) as described by
Amann (3). Fixed cells were washed again in PBS and stored
at 4°C in PBS until they were used (normally within 3 days after
fixation). For long-term storage, the cells were resuspended in a 1:1
mixture of PBS and 96% (vol/vol) ethanol and kept at 
20°C.
Activated sludge was obtained from the secondary aerated nitrification
basin (27,144 m3) of the Munich II wastewater
treatment plant (one million population equivalents). Then 12.5 ml of
activated sludge was fixed immediately after sampling by adding 37.5 ml
of 4% paraformaldehyde. After 5 h of incubation at 4°C, the
activated sludge was centrifuged for 5 min at 4,550 × g, and the supernatant containing the fixative was
discarded. Subsequently, the sludge pellet was washed with PBS and
finally resuspended in 12.5 ml (the original sample volume) of a 1:1
mixture of PBS and 96% (vol/vol) ethanol. Samples were stored at
20°C.
Cell concentration of pure cultures.
The numbers of cells
per milliliter in the fixed pure cultures of C. testosteroni, G. asaii, and N. europaea were
determined with a Neubauer cell counting chamber (Paul Marienfeld GmbH,
Bad Mergentheim, Germany) following the instructions of the manufacturer.
The cell concentrations of E. coli cultures were inferred
from the optical density at 600 nm (OD600) using
a DU 650 spectrophotometer (Beckman, Fullerton, Calif.). For
calibration, an E. coli TOP10F' overnight culture was
diluted by factors of 105 to
107, the OD600 of these
dilutions was measured, and aliquots were streaked onto petri dishes
with solid LB medium. The numbers of CFU were correlated with the
OD600 for each dilution, and the resulting
conversion factor (5.5 × 108 CFU
ml1 per OD600 unit) was
used to calculate the concentration of E. coli cultures
based on their OD600 in all following
experiments. It is important to note that E. coli LB
overnight cultures contain insignificant numbers of nonviable cells
(40). In addition, microscopic observation of the E. coli culture used showed that the vast majority of cells occurred
as single cells.
Spiking of pure culture mixtures and activated sludge with
E. coli cells.
Overnight cultures of C. testosteroni and G. asaii (100 ml) were harvested by
centrifugation for 10 min at 4,550 × g and fixed with
paraformaldehyde as described above. The cell densities in these
concentrated stock solutions were determined using the Neubauer chamber. Cell densities of E. coli overnight cultures were
determined photometrically, and E. coli cells were
concentrated and fixed with paraformaldehyde as described above.
Subsequently, different 1-ml cell mixtures containing 6.3 × 107 C. testosteroni and 3.7 × 107 G. asaii cells as well as
106, 107,
108, or 109 E. coli cells were prepared in 50% PBS-ethanol (EtOH) (vol/vol). In
addition, a 1-ml cell mixture containing 6.3 × 107 C. testosteroni and 3.7 × 107 G. asaii cells but without
E. coli cells was prepared in 50% PBS-EtOH (vol/vol).
Nitrifying activated sludge was spiked with different amounts of
E. coli using the following protocol. One milliliter of
paraformaldehyde-fixed activated sludge samples was centrifuged for 5 min at 10,000 × g. The supernatant was removed, and
the activated sludge was resuspended in 1 ml of PBS containing either
106, 107,
108, or 109
paraformaldehyde-fixed E. coli cells and mixed by vortexing
(10 s). In an additional experiment, 1.7 × 108 paraformaldehyde-fixed N. europaea
cells were added to the paraformaldehyde-fixed sludge prior to the
centrifugation step. Finally, all spiked activated sludge samples were
centrifuged for 5 min at 10,000 × g. The supernatant was carefully removed, and the sludge was resuspended in 1 ml of 50%
PBS-EtOH (vol/vol).
FISH.
The defined mixtures of pure cultures were spotted
onto microscope slides (Paul Marienfeld GmbH, Bad Mergentheim, Germany) and allowed to dry at 46°C. Afterwards, the slides were immersed for
2 to 3 s in molten 0.5% agarose (Gibco-BRL ultrapure agarose; Life Technologies, Paisley, Scotland) at 37°C. The slides were then
placed on ice until the agarose had solidified. Excess agarose on the
back side of the slides was removed, and the samples were dehydrated in
50, 80, and 96% (vol/vol) ethanol for 5 min each. The agarose coating
was applied to minimize cell loss during the following hybridization
and washing steps. After an additional drying step at room temperature,
whole-cell hybridization was performed as described by Manz et al.
(22).
A different protocol was developed for the treatment of activated
sludge samples prior to FISH. The sludge was prepared on the same type
of microscope slides, but the spotting and drying procedures were
repeated three times to obtain a thick layer of sludge flocs on the
slide surface. The average sample thickness was about 50 µm and thus
did not hamper laser penetration in the subsequent CLSM analyses.
Afterwards, the slides were immersed in agarose and hybridized
as described above. The thick layer of sludge flocs was required (i) to
record as many target cells as possible in order to improve the
accuracy of the measurements and (ii) to avoid bias in the
quantification of planktonic cells which otherwise would accumulate on
the slide surface.
The rRNA-directed oligonucleotide probes used for FISH were 5' labeled
with the dye Fluos
[5(6)-carboxyfluorescein-N-hydroxysuccinimide ester] or
with one of the sulfoindocyanine dyes indocarbocyanine (Cy3) and
indodicarbocyanine (Cy5). Labeled probes and unlabeled competitor
oligonucleotides were obtained from MWG (Ebersberg, Germany) or Thermo
Hybaid (Interactiva Division, Ulm, Germany). In all experiments,
group-specific probes labeled with Cy3 or Fluos were used together with
the Cy5-labeled EUB338 probe mix (consisting of probes EUB338,
EUB338-II, and EUB338-III) covering the domain Bacteria
(9). The probes used, their sequences, and their
specificities are listed in Table 1.
Microscopy and digital image analysis.
After in situ
hybridization, the microscope slides were embedded in Citifluor AF1
(Citifluor, Canterbury, United Kingdom). Pictures of fluorescent
cells were recorded using a CLSM (LSM 510, Zeiss, Oberkochen, Germany).
For detection of Cy3- and Cy5-labeled cells, two helium-neon lasers
(543 nm and 633 nm, respectively) and, for Fluos-labeled cells, an
argon laser (450 to 514 nm) was used. For each microscope field,
fluorescence conferred by the different probes was recorded in separate
images. For each hybridization experiment, 30 microscope fields at
random positions and in random focal planes were recorded using a Zeiss
Plan-Neofluar 40×/1.3 oil objective. This procedure (30 images at low
magnification) allows us to record a high number of probe-target cells
and thus to accurately determine the relative abundance of
heterogeneously distributed probe-target cells in activated sludge
samples (8). All pictures acquired corresponded to optical
sections of 1-µm thickness obtained by adjusting the pinhole diameter
of the CLSM accordingly. They were recorded as 8-bit images of 512 by
512 pixels with a resolution of 1.6 by 1.6 pixels per µm.
For each sample analyzed, detector gain, amplification offset, and
amplification gain settings were selected which allowed detection of
all probe-labeled cells with an intensity between 20 and 255. Special
attention was paid to optimize microscopic parameter settings so that
the images of those cells detected by the specific probes were
congruent with their counterparts in the picture with the EUB338 probe
mix-stained cells (Fig.
1B, C, and E). The cell
area quantification (see below) relies on this congruency, because it
is assumed that for each quantified cell the same area is measured with
the specific and with the universal probes.
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FIG. 1.
Principle of the cell quantification method developed
in this study. See text for an explanation of steps 1 to 7. (A)
Microscopic picture of activated sludge from the Munich II wastewater
treatment plant. (B and C) Aliquots of the same activated sludge after
spiking with 108 (B) and 109 (C) E.
coli cells per ml. FISH was performed with probe GAM42a (red)
and the EUB338 probe mix (green). The images containing the
fluorescence conferred by the probes were superimposed, and E.
coli cells appear yellow due to color blending. (D) Calibration
curve generated from corrected cell area fractions of E.
coli in spiked sludge aliquots. (E) The same microscopic field
as in A, showing simultaneous FISH using probes NEU and Nso1225 (red)
and the EUB338 probe mix (green). Cells of ammonia oxidizers appear
yellow due to color blending. (F) Graph showing the use of the
calibration curve (D) for converting the measured area fraction of an
autochthonous population to the equivalent E. coli
concentration. (G) Highly magnified optical section through a cell
aggregate of ammonia-oxidizing bacteria in activated sludge stained by
FISH using probes NEU and Nso1225 (red). (H) Highly magnified optical
section through E. coli cells from a pure culture
stained by FISH using probe GAM42a (red).
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It should be noted that cells of a population to be quantified will be
recorded as longitudinal sections as well as transverse sections (and
various intermediate forms). However, this fact has no significant
influence on the quantification accuracy, because both the cells
belonging to the indigenous bacterial population which is to be
quantified and the E. coli cells used as the internal standard are optically sectioned in random directions. Moreover, the
cell areas are determined by analyzing large numbers of probe-stained cells, a procedure significantly lowering the impact of the spatial orientation of individual cells.
The images were then exported as TIFF files by the image acquisition
software delivered with the microscope (Zeiss LSM 5, version 2.01).
These files were analyzed with the image-processing software (Zeiss
Kontron KS400, version 3.0) to measure the combined areas of stained
cells within each image as described by Schmid et al.
(32). The area fraction of specifically stained cells was
calculated as a percentage of the total area of bacteria stained by the
EUB338 probe mix in the same optical section.
Determination of cell density.
The cell concentrations of
probe-defined indigenous bacterial populations in a sample were
calculated using a seven-step procedure. First, aliquots of the sample
(e.g., activated sludge, Fig. 1A) were spiked with E. coli
(106 to 109 cells
ml1) as described above (Fig. 1, step 1), and
the spiked aliquots were stained by FISH using probe GAM42a and the
EUB338 probe mix (Fig. 1B and C). The area fraction of the E. coli cells in each spiked aliquot was determined by digital image
analysis (Fig. 1, step 2). For activated sludge samples, the area
fraction of the inherent 
-Proteobacteria was measured in
sludge aliquots without addition of E. coli cells. The area
fraction of these indigenous cells was subsequently subtracted from the
area fraction measured with probe GAM42a in the spiked aliquots to
obtain the area occupied by E. coli. Since spiking the
samples with E. coli increased not only the area fraction of
the cells stained by probe GAM42a but also the total area of all
bacteria, the measured E. coli cell area fraction must be
corrected to remain directly proportional to the number of added
E. coli cells. The corrected area fraction is calculated by
the formula
|
(1)
|
where Aec is the measured and
A
is the
corrected E. coli area fraction (in percent). The corrected area fractions were then plotted in a double-logarithmic graph against
the E. coli concentration, and a regression line was
calculated based on these data points (Fig. 1D). The double-logarithmic
transformation was necessary to meet a requirement for linear
regression, i.e., an equal variance for all measurements. The
regression line was used to calculate the "equivalent E. coli concentrations" from the area fractions of specifically
labeled bacterial populations (e.g., ammonia oxidizers stained within
activated sludge by FISH; Fig. 1, step 3, and Fig. 1E) in unspiked
aliquots of the samples by applying the following equation (Fig. 1,
step 4, and Fig. 1F):
|
(2)
|
where Ceq is the equivalent
E. coli concentration of the bacterial population (in cells
per milliliter), A is the measured area fraction of this
population, m is the slope, and b is the ordinate
intercept of the regression line.
Finally, Ceq was converted to the real
concentration of the bacterial population by taking into consideration
differences in size between E. coli and the probe-target
population. This conversion was accomplished by measuring the average
area of single E. coli cells and of single cells belonging
to the probe-target population of interest. For this purpose, images
that contained single cells of E. coli and of the
probe-target population were acquired at a high magnification (×5,000)
with a resolution of 55.6 by 55.6 pixels per µm (Fig. 1, step 5, and
Fig. 1G and H). Then a conversion factor was calculated as the ratio of
the average cell areas (Fig. 1, step 6):
|
(3)
|
where f is the conversion factor,
is the average single-cell area of the
population whose concentration was to be determined, and
ec is the average single-cell
area of E. coli. Eventually, Ceq was converted to the real
concentration by multiplication with the conversion factor (Fig. 1,
step 7):
|
(4)
|
where C is the concentration of the bacterial
population (in cells per milliliter).
Estimation of in situ substrate turnover rates.
Average
substrate turnover rates of ammonia-oxidizing bacteria were estimated
based on the measured cell concentrations. The total number of
ammonia-oxidizing cells in the aerated nitrifying basin of the
municipal Munich II wastewater treatment plant was calculated by
multiplying the number of ammonia-oxidizing cells per milliliter of
activated sludge by the reactor volume. The amount of ammonia-nitrogen
converted to nitrite (in milligrams per hour) was estimated according
to the formula
|
(5)
|
where
NH4+t
is the transformed ammonia-nitrogen (in milligrams per hour),
NH4i+ is
the ammonia-nitrogen concentration in the influent (in milligrams per
cubic meter),
NH4e+ is the
ammonia-nitrogen concentration in the effluent (in milligrams per cubic
meter), r is the reactor influent rate (7,858 m3 h1), and 0.9 is a
correction factor.
The estimated correction factor takes into account that ammonia is
removed from the sewage via autotrophic nitrification but also via
adsorption (27) and assimilation (activated sludge models
1 to 3 [12-14]). The estimated amount of ammonia
oxidized autotrophically was converted from milligrams per hour to
femtomoles per hour and was divided by the total number of
ammonia-oxidizer cells in the reactor to obtain the substrate turnover
rate in femtomoles of ammonia transformed to nitrite per hour and per cell.
| |
RESULTS |
Preparation of the samples and spiking with E.
coli
A relatively homogeneous distribution of the added
E. coli cells within a spiked sample is critical for
obtaining an accurate calibration curve. This is particularly important
for the measurements of sludge samples which were amended with
relatively small numbers of E. coli. Therefore, the area
fractions of E. coli cells added to activated sludge
were compared after vigorously vortexing the sludge for 1 min or after
homogenizing it with an Ultra-Turrax blender (IKA Labortechnik,
Staufen, Germany) treatment (1 min). The sludge was spiked with
E. coli either before or after these pretreatments.
The area fraction of E. coli was determined for each sample
by FISH with probe GAM42a and the EUB338 probe mix and digital image
analysis using the method previously published by Schmid et al.
(32). The area fractions were compared to the values measured with spiked sludge that had not been vortexed or homogenized. Neither the kind of pretreatment nor the order of sludge preparation and spiking affected the measured area fractions and their standard deviations (Table 2). Consequently,
E. coli cells were simply added to the fixed activated
sludge samples without additional pretreatments in all following
experiments.
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TABLE 2.
Effect of different activated sludge homogenization
procedures on area fraction measurements of E. coli
cellsa
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Evaluation of the quantification protocol using artificial mixtures
of pure cultures.
The developed quantification approach was first
tested with bacterial pure cultures. For this purpose, cultures of
G. asaii (
-subclass of Proteobacteria) and
C. testosteroni (
-subclass of Proteobacteria)
were mixed after their cell concentrations had been determined with a
Neubauer cell counting chamber. The final cell concentrations in
the mixture were 3.7 × 107 ± 0.5 × 107 cells ml1 for
G. asaii and 6.3 × 107 ± 0.2 × 107 cells ml1
for C. testosteroni (all the confidence limits indicate 95%
confidence intervals).
Aliquots of this mixture were supplemented with increasing
concentrations of E. coli cells, and a regression line was
generated from a graph depicting the relative cell areas of E. coli in the spiked aliquots versus the amount of E. coli cells added (Fig. 2a). This
regression line should have a slope of approximately 1, as the
corrected area fraction of E. coli cells is expected to be
directly proportional to the amount of E. coli cells added. The slope in this and all other determined calibration curves (see also
below) was slightly higher than 1 (e.g., 1.25 in Fig. 2a). This implies
that small cell additions had a large effect on the area fraction,
whereas large additions only had a more moderate effect. Subsequently,
the unspiked mixture was hybridized with the EUB338 probe mix and with
probe ALF1b or BET42a. It should be noted that the intensities of the
fluorescent signals varied considerably among the individual cells of
either species, G. asaii and C. testosteroni.
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FIG. 2.
Calibration curves used to convert cell area fractions
to equivalent E. coli concentrations for the
determination of cell concentrations in pure culture mixtures of
C. testosteroni and G. asaii (A), in
activated sludge from the Munich II wastewater treatment plant (B), and
in the same activated sludge after addition of 1.7 × 108 N. europaea cells per ml (C). The
general linear equation of these curves is log
Ceq = m × log A + b, where
Ceq is the equivalent E. coli concentration, A is the cell area fraction,
m is the slope, and b is the ordinate intercept
of the line. The values of m are 1.253317 (A),
1.105818 (B), and 1.004006 (C). The values of b are 5.27404 (A), 6.429176 (B), and 6.83411 (C). Error bars which are smaller
than the marker symbols are not shown.
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The cell area fractions of G. asaii and C. testosteroni were measured, and the calibration curve and area
correction factor were used to convert the areas to cell concentrations
as described above. Table 3 shows the
results of this experiment. The concentration determined for G. asaii deviates by 0.4 × 107 cells
ml1 (or 10.8%) from the Neubauer cell chamber
count, while the difference between the two quantification methods
amounts to 0.8 × 107 cells
ml1 (or 12.7%) for C. testosteroni.
Quantification of ammonia-oxidizing bacteria in activated
sludge.
The number of autochthonous ammonia-oxidizing bacteria was
determined in a nitrifying activated sludge from the Munich II wastewater treatment plant. An equimolar mixture of probes Nso1225 and
NEU was used for the in situ detection of ammonia oxidizers of the
-subclass of Proteobacteria. Probe Nso1225 targets all recognized ammonia oxidizers of the -subclass with the exception of
Nitrosococcus mobilis (28). The single central
mismatch of N. mobilis is discriminative under stringent
conditions, so probe NEU was used in addition. This probe targets
N. mobilis and other not yet described ammonia oxidizers
from activated sludge which are also not detected by probe Nso1225
(unpublished results).
A first inspection of the analyzed nitrifying activated sludge by FISH
with probes NEU and Nso1225 confirmed the occurrence of large amounts
of ammonia-oxidizing bacteria of the -subclass of
Proteobacteria. Most ammonia oxidizers were relatively small and formed spherical, tightly packed cell clusters, but others were
slightly larger and formed looser aggregates with narrow intercellular cavities.
The cell concentration of the ammonia oxidizers was determined and
confirmed in two experiments. First, their area fraction was measured
after FISH with both probes NEU and Nso1225 and the EUB338 probe mix.
The cell concentration of the autochthonous ammonia oxidizers was then
calculated based on a calibration curve that had been generated by
spiking of the sludge with E. coli (Fig. 2b). In the second
experiment, 1.7 × 108 ± 0.3 × 108 N. europaea cells per ml were
added to the activated sludge. The concentration of the ammonia
oxidizers (consisting of the autochthonous and the added ammonia
oxidizers) in the modified sample was measured by the same procedure as
in the original sludge, but a new calibration curve was generated after
aliquots of the modified sludge had been spiked with E. coli
(Fig. 2c). Finally, the concentration of the autochthonous ammonia
oxidizers in the original sludge was subtracted from the concentration
of the ammonia oxidizers in the sludge supplemented with the additional
N. europaea cells.
For the autochthonous ammonia oxidizers, a cell area fraction of 8.4% ± 1.4% was measured, which corresponds to a concentration of 9.8 × 107 ± 1.9 × 107
cells per ml of activated sludge. Following the addition of 1.7 × 108 ± 0.3 × 108
N. europaea cells per ml, the area fraction of the
ammonia oxidizers increased slightly to 9.4% ± 1.4%. It should be
noted that different image acquisition parameters were used in the two
experiments, making it impossible to compare the area fractions
directly. This was necessary because the added ammonia oxidizers showed
a weaker fluorescence after FISH than the autochthonous ammonia
oxidizers. Furthermore, the added N. europaea cells
affect the calibration curve so that the small change in area fraction
corresponds to a large difference in cell numbers when using the
appropriate new calibration curve. The resulting absolute cell
concentrations, however, are comparable.
The concentration of ammonia oxidizers after amendment was 2.3 × 108 ± 0.3 × 108
cells ml1. The difference between this value
and the concentration of the autochthonous ammonia oxidizers amounts to
1.3 × 108 cells ml1
which should be equal to the 1.7 × 108
added N. europaea cells per ml. The deviation between the
two values, 4 × 107 cells
ml1, is 22.4% of the cell addition and thus
about twice as high as the differences between the cell concentrations
measured by the newly developed quantification method and obtained by
using the Neubauer chamber for the pure culture mixtures (see above).
Estimated activity of ammonia-oxidizing bacteria in activated
sludge.
The average rate of ammonia oxidation per autochthonous
ammonia-oxidizer cell within the activated sludge was calculated based on the determined concentration of autochthonous ammonia oxidizers. As
described above, 9.8 × 107 ± 1.9 × 107 ammonia-oxidizer cells
ml1 were found in the activated sludge. Thus,
with a total reactor volume of 27,144 m3 the
total amount of ammonia oxidizers in the reactor was 2.7 × 1018 ± 0.5 × 1018 cells.
During the last 2 weeks before sampling, the amount of
NH4+-N was in the range of 10 to
16 mg liter1 in the influent and in the range
of 0.05 to 0.3 mg liter1 in the effluent of the
plant, respectively (Fig. 3). The average concentrations of NH4+-N in the
influent (12.1 mg liter1) and the effluent
(0.08 mg liter1) of the plant during the last 6 days before sampling were used for the activity estimation. Thus, with
a flow rate of 7,858 m3 h1, 9.5 × 107 mg of
NH4+-N
h1 was transformed in the basin. Assuming that
10% of the ammonia was not removed by autotrophic oxidation, the
ammonia oxidizers oxidized 8.5 × 107 mg of
NH4+-N
h1 which equals 6.1 × 1018 fmol of
NH4+ h1.
Consequently, each ammonia-oxidizer cell converted 2.3 ± 0.4 fmol
of NH4+ to
NO2 per hour if equal activity
of all ammonia-oxidizer cells is assumed.
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|
FIG. 3.
Concentrations of ammonia nitrogen in the influent ( )
and the effluent ( ) of the nitrification basin of the Munich II
wastewater treatment plant measured during the last 2 weeks before
sampling.
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DISCUSSION |
Cell quantification methods suitable for microbial ecology should
provide precise results for environmental samples containing bacteria
which are not homogeneously distributed. Furthermore, they should be
independent of cultivation, as most bacteria in natural or engineered
systems have hitherto not been isolated (5, 34). A
generally applicable method should allow specific enumeration of both
single species and higher phylogenetic taxa (e.g., genera or
subclasses). New protocols must be tested with regard to these
requirements by use of suitable model systems.
In this study, mixtures of pure cultures and an activated sludge sample
were used, because these model systems have different advantages when
evaluating a new quantification method. The selected pure cultures
consisted of uniformly shaped cells, which can easily be counted in a
counting chamber to verify the results obtained with the new
quantification protocol. On the contrary, activated sludge contains
numerous different cultivated and uncultivated prokaryotic species
(2, 7, 33) with different morphologies and abundance and
thus represents a challenge for any quantification method.
Applicability and accuracy of the newly developed quantification
method.
The new quantification method was successfully applied to
measure cell concentrations of specific populations in bacterial pure
culture mixtures as well as in activated sludge. The results obtained
were not expressed as relative abundance based on a reference value
such as total cell counts, but as absolute cell numbers per volumetric
unit. Since the developed quantification procedure is based on FISH
with rRNA-directed oligonucleotide probes, it can be used in any
environment that is amenable to FISH analysis. It makes no difference
whether the quantified organisms grow as single cells or in dense
aggregates as long as individual cells can be resolved microscopically.
However, the accuracy of the results obtained with this method depends
on a uniform cell size of the target population (see below). Size
variations within a target population can be caused by polymorphism of
a single species or if probes with a broader specificity were applied
for detection of morphologically different bacterial taxa. It should
also be noted that the accuracy of this method depends on a relatively homogeneous distribution of reference cells (used for spiking) within
the environmental sample. For the analyzed activated sludge sample,
this was easily achieved by adding the E. coli cells to the
sample followed by a short mixing step. Since composition and density
of aggregates or flocs might vary between different environments,
special pretreatment (e.g., homogenization) of other samples may be
necessary to ensure optimal dispersal of the cells used for spiking.
The cell concentrations measured with the newly developed method
deviated in the mixed pure culture experiments by approximately ±10 to
13% and in the experiments with activated sludge by approximately ±22% from the Neubauer chamber counts. In addition to the measurement error of the Neubauer chamber, several difficulties with the area measurement of FISH-stained cells could have caused these
discrepancies. First of all, the intensity of the FISH signal is a
function of the ribosome content of the target cells. Although most
cells in actively growing pure cultures contain high ribosome numbers, we observed that a fraction of the FISH-stained C. testosteroni and G. asaii cells emitted less
fluorescence than the majority of the labeled cells. Such differences
in the fluorescent signal intensity were even more pronounced between
different bacterial populations in the activated sludge sample. The
presence of very bright and relatively dark cells in the same sample
makes it difficult to find appropriate microscope parameter settings
and intensity thresholds during image analysis to differentiate between
cells and background. Under such conditions, either the areas of the bright cells are overestimated when the threshold is too low, or the
darker cells are not included in the analysis when the threshold is too
high. Such errors may still be higher in samples from
oligotrophic environments, where the growth rates and ribosome content
of indigenous bacterial populations may differ more
pronouncedly than in activated sludge.
Problems with fluorescence intensities are also responsible for the
upper limit of cell concentrations that can be quantified by the newly
developed method. Very high concentrations of probe-target cells (e.g.,
108 cells per ml) require that the sample be
spiked with E. coli concentrations of between
107 and 109 cells per ml to
obtain a calibration curve that spans at least one order of magnitude
above and below the cell concentration to be measured. After addition
of 109 E. coli cells per ml, however,
the E. coli cells formed thick layers of stacked cells on
the microscope slides. Following FISH with the EUB338 probe mix, the
local fluorescence intensity within these layers of E. coli
cells was far higher than the fluorescence intensities observed for
most aggregates of autochthonous bacteria. As a consequence, the CLSM
detector collected too much light at the locations of the E. coli layers, and the E. coli cells appeared too large
in the images with the EUB338 probe mix-stained cells. The sensitivity
of the detector could not be reduced to overcome this problem, because
then the darker autochthonous bacteria would not have been detected
anymore. This problem hampered the precise determination of area
fractions for high E. coli cell densities and in consequence
affected the precision of the calibration curves. Thick E. coli cell layers were not observed after spiking with smaller
amounts of E. coli than 109 cells per
ml. The quantification accuracy can thus be expected to be higher for
lower concentrations of probe-target cells that do not require spiking
of the sample with 109 E. coli cells
per ml to obtain a suitable calibration curve.
Furthermore, the conversion of the equivalent E. coli
concentration to the real concentration of the quantified population is
a possible source of measurement error. The ratio of the average cell
areas is used as the conversion factor, but even in pure cultures cell
size, and therefore the cell area, may vary considerably. This problem
could have contributed to the observed differences between Neubauer
chamber counts and the counts inferred from the novel quantification
method. In complex systems like activated sludge, cell size variation
of probe-target bacteria is frequently observed. Therefore, application
of the developed quantification method is recommended for
quantification of probe-defined groups of microorganisms that do not
show pronounced differences in size, like the two populations of
ammonia-oxidizing bacteria detected in this study.
Despite these possible sources of error, the novel FISH-based
quantification method constitutes a straightforward and precise method
to determine the absolute numbers of microorganisms in different
environments and is especially useful for samples containing biofilms
or aggregates. The accuracy of the method is demonstrated by the highly
similar cell concentrations obtained using the well-established Neubauer chamber counts and the novel FISH-based quantification method
for pure culture mixtures as well as for an activated sludge which was
amended with a defined number of N. europaea cells.
The utility of the developed quantification method to enumerate
bacteria in samples where cells are not homogeneously distributed was
illustrated by quantification of autochthonous ammonia-oxidizing bacteria in a nitrifying activated sludge. Based on the absolute numbers of ammonia-oxidizing bacteria obtained, their average activity
in the municipal activated sludge sample was estimated to be 2.3 ± 0.4 fmol of NH4+
cell1 h1, a value which
is within the range of per cell activities measured with pure cultures
of N. europaea (1.24 to 23 fmol of
NH4+
cell1 h1)
(20). Compared to manual counting of probe-labeled cells
by microscopy (21, 24, 34, 35), the new semiautomatic
method is less tedious and not negatively affected by aggregates or
cell clusters, and its measured standard error is lower.
This study was supported by Sonderforschungsbereich 411 from the
Deutsche Forschungsgemeinschaft (Research Center of Fundamental Studies
of Aerobic Biological Wastewater Treatment). The International Workshop
on New Techniques in Microbial Ecology (INTIME), where the basic
concept of this study was outlined, is acknowledged as a forum
encouraging the realization of joint projects between the University of
Aarhus, the University of Aalborg, and the Technische Universität
München.
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Abstract
Introduction
