Environ. Sci. Technol. 2005, 39, 3289-3294
New Method for Assimilable
Organic Carbon Determination Using
Flow-Cytometric Enumeration and a
Natural Microbial Consortium as
Inoculum
FREDERIK A. HAMMES AND
THOMAS EGLI*
Department of Environmental Microbiology, Swiss Federal
Institute for Environmental Science and Technology (EAWAG),
Überlandstrasse 133, CH-8600 Dübendorf, Switzerland
The concentration of easily assimilable organic carbon
(AOC) largely determines the microbiological stability of
drinking water. However, AOC determination is often neglected
in practice due to the complex and tedious nature of the
conventional bioassay. The three major drawbacks of the
conventional method are (1) a long assay time of 9-12
days, (2) the use of a labor-intensive enumeration technique
(plating on growth media), and (3) limited information
supplied by the use of selected pure cultures (Pseudomonas
fluorescens P-17 and Spirillum NOX) for measuring a
complex pool of natural bioavailable carbon compounds.
A new method is proposed here, in which plating was replaced
with fluorescence staining of total nucleic acids combined
with flow cytometry as a rapid and straightforward
growth enumeration method. This approach also allowed
for the detection of inactive and/or unculturable microorganisms. Hence, the conventionally used pure cultures
were replaced in the new AOC assay with a natural microbial
consortium. It was shown that the flow-cytometric
enumeration method could be used to establish complete
growth curves for a natural microbial consortium growing
on AOC. Compared to the end-point measurements of the
conventional method, such kinetic data provide much
clearer insight into the actual growth potential of a water.
Introduction
Assimilable organic carbon (AOC) is a collective term
describing the fraction of labile dissolved organic carbon
that is readily assimilated by microorganisms, resulting in
growth. It consists of a broad range of low molecular weight
organic carbon molecules such as sugars, organic acids, and
amino acids. AOC is a critical parameter for drinking water
treatment and distribution processes. It represents only a
small fraction (0.1-9%) of the total organic carbon (TOC) in
water, but it is regarded as one of the main factors governing
heterotrophic growth, and thus biological water stability
(1, 2). Several studies have linked AOC directly to microbial
regrowth and biofilm formation in drinking water distribution
systems (2, 3). In addition, the present trend in drinking water
treatment is toward a combination of treatments which will
* Corresponding author phone: +41-44-823 51 58; fax:
+41-44-823 55 47; e-mail: [email protected].
10.1021/es048277c CCC: $30.25
Published on Web 03/31/2005
 2005 American Chemical Society
completely remove the nutrients that support microbial
growth, namely, carbon (C), nitrogen (N), and phosphorus
(P) (1). Based on the C:N:P molecular ratio of bacteria and
biomass (100:10:1), the growth determining factor in most
waters is usually carbon, i.e., AOC (1). These points emphasize
the need for a reliable, realistic, and easily applicable AOC
determination method.
Conventional AOC analysis is done with a bioassay that
was originally developed by Van der Kooij and co-workers
(4) and later adapted by others (1, 5, 6), of which a detailed
description is presented in Standard Methods (7). In short,
this bioassay quantifies, by plating, bacterial batch growth
as the number of colony forming units (cfu) in a water sample
from inoculation until stationary phase, usually performed
as an “end-point” measurement only. Pure cultures of either
Pseudomonas fluorescens (P. fluorescens) strain P-17 or
Spirillum strain NOX are most often used as test organisms,
due to their different nutritional capabilities. After inoculation
(prescribed at 500 cfu mL-1), the water sample is incubated
at 15 °C for 9 days, and microbial growth is quantified on
days 7, 8, and 9 with plating on nutrient agar. The result
(average net growth) is thereafter related to the growth of the
test organisms on pure solutions of acetate (P-17) or oxalate
(NOX) by means of prederived yield values, and the final
result is given as acetate-C equivalents (7). Typically, a water
sample is considered to be biologically stable if it has an
AOC concentration of less than 10 µg L-1 acetate-C equivalents, though this value depends on the presence of residual
chlorine in the system (3), and whether indeed the water is
limited in carbon and not phosphorus (8). Some improvements on the original method have been proposed by Kaplan
et al. (5) and LeChevallier et al. (6) and included the use of
a higher inoculum density ((2-4) × 104 cfu mL-1), a higher
incubation temperature (specifically 22 °C for P-17). LeChevallier et al. (6) also proposed adenosine triphosphate (ATP)
analysis instead for an estimation of bacterial growth, but
plating remains the most generally applied enumeration
method. Despite the relevance of AOC data to the industry,
AOC measurement is still not performed routinely in practice
and/or used often in the design and optimization of drinking
water treatment, storage, and distribution systems. We believe
that this is because the conventional method is somewhat
tedious, time-consuming, and labor-intensive. Moreover, we
question to which extent a pure culture(s), preconditioned
on a single pure AOC compound, is able to give a reliable
reflection of the AOC content of complex natural waters.
Flow cytometry coupled with fluorescent staining has
emerged as the leading tool for single-cell analysis in
microbiology. Multiple examples already exist of the application of flow cytometry for the enumeration of either
total bacteria or specific bacterial groups using either total
nucleic acid or selective RNA stains (9). Probably the biggest
advantage of flow cytometry is that it allows for rapid and
accurate enumeration of all cells, including those which are
unculturable or inactive. An obvious offspring of this is the
ability to quantify a natural microbial consortium consisting
largely of microbial cells that cannot be cultured on
conventional media. In this study we describe the use of a
recently patented method that entails fluorescent staining
and flow cytometry instead of conventional plating or ATP
analysis for quantification of microbial growth during the
AOC measurement (10). This method has the advantage that
it is fast, reliable, and reproducible. It is furthermore
demonstrated that when combined with a natural microbial
consortium and kinetic growth measurements, a more
realistic interpretation of AOC and of the microbial regrowth
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potential in natural water samples is obtained as compared
with only end-point data.
Experimental Section
Bacterial Strains and Natural Microbial Consortium. A pure
culture of P. fluorescens strain P-17 (ATCC 49642) (from here
on referred to as P-17) was obtained from D. Van der Kooij
(Kiwa, Nieuwegein, The Netherlands). The culture was stored
in 20% glycerol at -70 °C and cultivated on R2A agar (Oxoid)
(15 °C, 3-5 days) prior to use. AOC-free stock solutions of
the pure culture were prepared as described in Greenberg
et al. (7), and these solutions were stored for up to 3 months
at 5 °C. An AOC-free solution of a natural microbial
consortium was prepared as follow: 40 mL of water, sampled
directly after an activated carbon filter unit in a drinking
water treatment plant, was filtered (0.22 µm, Millex-GP,
Millipore) for removal of particulate organic carbon, inoculated with 100 µL of unfiltered water, and incubated in AOCfree vials without further amendments at 30 °C for 14 days.
The cells were subsequently harvested by centrifugation
(10 min, 3000g) and resuspended in HPLC-grade water (Fluka,
CH) amended with a mineral buffer as described by LeChevallier et al. (6). This solution was then incubated for a further
7 days to ensure that all residual organic carbon had been
degraded. Necessary controls were performed to validate
the AOC-free status of the inoculum solutions.
Preparation of AOC-Free Materials. Borosilicate glass
vials (40 mL) with screw caps containing TFE-lined silicone
septa were used for the assays. Carbon-free vials were
prepared as described in Greenberg et al. (7) as well as in
Charnock and Kjønnø (11). In short, the vials and screw caps
were first washed with common detergent and thereafter
rinsed thrice with deionized water. These were then submerged overnight in 0.2 N HCl and again rinsed thrice with
deionized water. The vials were subsequently heated in a
Muffel oven to 550 °C for at least 6 h (to remove all trace
organics). The screw caps were soaked in a 10% sodium
persulfate solution at 60 °C for at least 1 h, rinsed twice with
deionized water and once with HPLC-grade water and finally
air-dried.
Growth of P-17 on Acetate-C. A pure acetate solution
containing 100 µg L-1 acetate-C (added as sodium acetate;
Fluka; technical grade) was prepared with bottled mineral
water. Bottled mineral water was specifically used because
it is well-buffered and contains all the required minerals for
microbial growth. Aliquots (40 mL) of this acetate solution
were then filtered with non-AOC-releasing filters (0.22 µm)
into AOC-free vials, which were capped tightly and then
pasteurized (70 °C, 30 min). Control samples were prepared
in exactly the same way except that no acetate was added.
For the non-AOC-releasing filters, we have used Millex-GP
(Millipore) syringe-mounted filters which were flushed with
ultrapure water before use. Triplicate vials were inoculated
with 100 µL of the AOC-free P-17 inoculum, giving a final
concentration of approximately 1 × 104 ((4 × 102) cells mL-1
(measured with flow cytometry) or 8 × 103 ((8 × 102) cfu
mL-1 (measured with plating), and incubated at 15 °C for 10
days. Before sampling, vials were shaken for 30 s and
thereafter 1 mL aliquots were removed with a sterile pipet
for both flow cytometry and plating. Samples were taken at
regular intervals and analyzed by means of plating and flow
cytometry. For plating, samples were taken on designated
sampling days, serially diluted in physiological salt solution
(8.5 g L-1 NaCl), and plated on R2A agar using the spreadplate method. The plates were incubated at 15 °C for 3-5
days before counting.
Growth of a Natural Microbial Consortium on AcetateC. An acetate solution containing 100 µg L-1 acetate-C and
an acetate-free control solution were prepared as described
above. Aliquots (40 mL) of these solutions were then filtered
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into AOC-free vials with a syringe-mounted filter (0.22 µm).
The vials were not pasteurized again, as this step was deemed
unnecessary when a natural microbial consortium is used as
inoculum. Triplicate vials were inoculated with 100 µL of the
AOC-free consortium, giving a final concentration of 5 × 103
((2 × 102) cells mL-1 (measured with flow cytometry) and
incubated at respectively 15 and 30 °C for 9 days. Samples
were taken at regular intervals and analyzed by means of
flow cytometry.
Kinetic Measurements with the Natural Consortium.
For a first kinetic experiment, an acetate-C calibration series
was prepared similar to the acetate solutions above, using
commercial mineral water as matrix and control, at the
following concentrations: 0 (control), 50, and 100 µg L-1
acetate-C. In a second experiment, natural river water
(Chriesbach river, Dübendorf, CH) was filtered (0.22 µm)
and diluted with commercial mineral water to final concentrations of 1, 5, 10, 50, and 100% (undiluted) river water.
Water samples from both experiments were filtered and
inoculated with the natural consortium as described above
and incubated at 30 °C. Samples were taken at t ) 0, 12, 14,
16, 18, 20, 22, 26, 48, and 52 h after inoculation for the acetate
experiment and at t ) 0, 8, 10, 12, 14, 16, 19, 21, and 23 h
for the river water experiment. Samples were stained and
analyzed with flow cytometry as described below. The specific
growth rates (µ) for the natural microbial consortium in each
sample were determined as follows:
µ ) (ln(x) - ln(x0))/∆t
(1)
where x, x0 are the microbial concentration measured at two
time points and ∆t is the expired time interval between these
points.
Fluorescent Staining and Flow Cytometry. All enumeration samples (1 mL) were amended with 1% lysis buffer
(10% Triton X-100, 5% Tween 20, 10 mM TrisHCl, and 1 mM
ethylenediaminetetraacetic acid (EDTA)) and directly stained
with 10 µg mL-1 SYBR Green stain (1:100 dilution in dimethyl
sulfoxide (DMSO); Molecular Probes), and incubated in the
dark for at least 20 min before measurement. Where
necessary, samples were diluted after staining in filtered
(0.22 µm) mineral water, so that the concentration measured
in the flow cytometer was always less than 3 × 105 counts
mL-1. Flow cytometry was performed using a PASIII flow
cytometer (Partec, Münster, Germany) equipped with a
25 mW argon ion laser, emitting at a fixed wavelength of
488 nm. Green fluorescence was collected in the FL1 channel
(530 ( 30 nm) and red fluorescence collected in the FL3
channel (>590 nm). All parameters were collected as
logarithmic signals. Data were analyzed using Flowmax
software (Partec). The specific instrumental gain settings were
as follow: FL1 ) 420, FL3 ) 700, speed ) 3 (implying a count
rate of less than 500 events s-1). All samples were triggered
on green fluorescence (FL1). Staining results of selected
samples were controlled with fluorescence microscopy, using
an inverted microscope (Olympus IX 51) equipped with the
appropriate filters.
Results
Flow-Cytometric Absolute Cell Counting. SYBR green stains
total nucleic acids and emits, upon excitation at 488 nm, a
bright fluorescent signal which was detected on the FL1
channel at 530 ( 30 nm. Due to the emission spectrum of
the stain (see www.probes.com for details), a corresponding
signal was also detected above 590 nm (FL3 channel). Parts
A-F of Figure 1 show examples of typical histogram and dot
plot data generated when a sample of P. fluorescens strain
P-17 (Figure 1A-C) and a natural microbial consortium
(Figure 1D-F) is analyzed with flow cytometry. Gates were
defined on the combined dot plot (Figure 1C,F), to discrimi-
FIGURE 1. Typical examples of fluorescence histogram data and enumeration gates for P. fluorescens strain P-17 in pure water (A-C)
and a mixed microbial consortium in river water (D-F), after staining with SYBR green and flow-cytometric analysis. The x-axis shows
green (FL1) or red (FL3) fluorescence intensity, and the y-axis shows the number of events recorded for a corresponding fluorescence
intensity. Gates R1 on the combined fluorescence dot blots (C and F, respectively) were used for enumeration purposes, allowing clear
distinction between the background and the stained sample in natural water.
nate between positive signals and background using both
green and red fluorescence intensity of the particles. Note
that the intensity of the background signal tended to vary
depending on the water used. While deionized laboratory
water resulted in practically no visible background (Figure
1C), some natural water samples displayed severe background, most probably as a result of mineral formation in
calcium-rich water (Figure 1F). Hence, for the sake of
standardization, the gating on the dot plot was used in all
cases. The enumeration method was validated with a stock
solution of P-17. This control revealed the standard error of
the enumeration method to be 4.8%, of which 1.4% could be
ascribed to machine error and the rest to error during
sampling, dilution, and staining. Moreover, the counts
obtained with flow cytometry were confirmed for P-17 with
conventional plate counts on R2A agar, and the plating
efficiency exceeded 90% (data not shown).
Flow-Cytometric Enumeration Compared to Conventional Plating. Flow cytometry coupled with fluorescent
staining is a simple, fast, and statistically reliable method for
bacterial enumeration. Figure 2 shows a comparison between
this method and conventional plate counting in an AOC assay
of 100 µg L-1 acetate-C, inoculated with P-17 and incubated
at 15 °C. It is significant to note that in the crucial stationary
phase period of the assay (ca. day 5 onward), the flow
cytometry data displayed much more stability than the plating
data. This could be ascribed to the plating method being
prone either to human error or to physiological changes to
the bacteria (e.g., loss of activity and/or culturability). In
terms of statistical accuracy, the plate count method can
reliably detect between 30 and 300 events/plate, which are
converted to the final result based on the appropriate dilution
factor. For example, a sample containing 3 × 105 cfu mL-1
would require at least two 10-fold dilutions. Flow cytometry
detects up to 6 × 104 events per analysis (the total volume
analyzed is 200 µL), which means that a sample containing
3 × 105 cells mL-1 can be analyzed without dilution. The
average net growth in the samples was determined as the
difference between the average concentration on days 9 and
10 and the concentration directly after inoculation (t ) 0).
The control values were then subtracted from the acetatecontaining samples. On the basis of the empirical yield values
of 4.1 × 106 cfu (µg of acetate-C)-1 (7), the net growth recorded
in Figure 2 yields AOC values of respectively 137 ( 26 µg L-1
acetate-C for the plating method and 124 ( 9 µg L-1 acetate-C
for the flow-cytometry method. Evidently, the expected AOC
concentration was slightly overestimated in both instances.
It may well be that the pasteurization step used in these
analyses resulted in some minor carbon contamination of
the vials. The specific growth rate (µ) with acetate was
approximately 0.07 h-1, and in the control it was approximately 0.01 h-1.
Natural Microbial Consortium and Temperature. A
combination of fluorescent staining and flow cytometry
enabled the enumeration of a natural microbial consortium,
with flow cytometry data similar to those presented in Figure
1D-F. This method was used to assess the effect of different
incubation temperatures on the AOC assay with pure acetate
solutions. Figure 3 shows the consortium proliferating much
more rapidly with acetate at 30 °C than at 15 °C. Stationary
phase was reached at 30 °C between 30 and 40 h after
inoculation in both the control and the acetate-containing
sample. In the samples inoculated at 15 °C, stationary phase
was only reached after 70-130 h of incubation. The maximum
specific growth rates at 30 °C were approximately 0.18 and
0.14 h-1 for the acetate-containing and control samples,
respectively, and at 15 °C it was 0.09 and 0.07 h-1, respectively.
Moreover, the empirical yield values of 6.08 × 106 cells
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FIGURE 4. Batch growth curves for a natural microbial consortium
growing with acetate during 52 h of incubation at 30 °C (where
9e+5 equals, for example, 9 × 105). The maximum specific growth
rates in these samples were respectively 0.199 h-1 (r2 ) 0.992; b),
0.210 h-1 (r2 ) 0.997; O), and 0.202 h-1 (r2 ) 0.999; 1). Error bars
indicate standard deviation on triplicate samples.
FIGURE 2. Batch growth curves for P. fluorescens P-17 growing
with and without acetate as determined with plating (A) and flow
cytometry (B), respectively (where 9e+5 equals, for example, 9 ×
105). Error bars indicate standard deviation on triplicate samples.
FIGURE 3. Batch growth curves for a natural microbial consortium
growing with and without acetate at 15 and 30 °C, respectively
(where 1e+6 equals, for example, 1 × 106). Error bars indicate
standard deviation on triplicate samples.
(µg of acetate-C)-1 at 30 °C and 4.96 × 106 cells (µg of acetateC)-1 at 15 °C were higher than the values given in Standard
Methods (4.1 × 106 cfu (µg of acetate-C)-1) (7) but in a similar
range to those recorded for P-17 growth on acetate in this
work (5.1 × 106 cells (µg of acetate-C)-1) (Figure 2). Temperature also affected the yield measured as cell numbers.
This was evidenced by the yield in cell numbers at 30 °C,
which was notably higher than that recorded at 15 °C. The
significantly higher specific growth rate at 30 °C, which
resulted in stationary phase being reached more rapidly, was
the reason this incubation temperature was favored in further
experiments.
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Kinetic Analysis of Growth on AOC. Batch growth curves
of the natural microbial consortium on a single AOC
compound (acetate) at two concentrations were considerably
different from batch growth curves on a natural AOC mixture
(river water). Exponential growth was detected between 12
and 26 h after inoculation in the acetate-containing solutions
(Figure 4), with no significant differences in the specific
growth rates recorded for the various concentrations
(0.199-0.202 h-1; Figure 4). Note that the end-point cell
concentrations (t ) 48 and 52 h) correspond perfectly to the
added acetate concentrations. When the average cell concentration of the control was subtracted from the average
values for the acetate-containing samples, the resulting cell
concentration in the 50 µg L-1 acetate-C sample was 49.8%
of the concentration in the 100 µg L-1 acetate-C sample.
Independent triplicate vials that were only sampled after
72 h confirmed the end-point values, which also indicates
that no significant carbon contamination occurred as a result
of continuous sampling during the experiment. Figure 5A
shows batch growth curves for the natural consortium on
five dilutions of river water. Throughout the measuring
period, the different concentrations could be distinguished
from one another, and exponential growth was recorded in
all samples from as early as 8 h after inoculation (Figure 5B).
In fact, contrary to the acetate result above, it was apparent
that different natural AOC concentrations resulted in different
specific growth rates (Figure 5B). The lowest detected specific
growth rate was 0.164 h-1 for the 1% dilution, and the highest
detected specific growth rate was 0.351 h-1 for the 100%
river water sample. End-point measurements predicted less
accurately the AOC concentration than when acetate was
used as carbon source, especially at higher dilutions of the
river water. This can be ascribed to the influence of the
dilution water, e.g., a different pH, additional carbon or
additional micronutrients. Alternatively, it is conceivable that
different growth conditions might have affected the final cell
size. Thus, growth measured only as cell numbers might err
slightly in the estimation of total biomass increase.
Discussion
The conventional AOC bioassay is not used as commonly in
practice as one would expect from its importance for drinking
water production. This apparent neglect is in our opinion
due to the tedious nature of the conventional method and
also because of inevitable questions arising from the use of
a pure culture(s) to accurately assess a wide range of natural
AOC components. The aim of this work was to examine
fluorescence staining of total microbial nucleic acids coupled
FIGURE 5. Batch growth curves for a natural microbial consortium
growing on five dilutions of river water (A) and calculation of the
maximum specific growth rates in these samples (B), which were
respectively 0.16 h-1 (r2 ) 0.994; b), 0.24 h-1 (r2 ) 0.995; O),
0.25 h-1 (r2 ) 0.991; 1), 0.3 h-1 (r2 ) 0.995; 3), and 0.35 h-1
(r2 ) 0.998; 9) (where 6e+5 equals, for example, 6 × 105). Error bars
indicate standard deviation on triplicate samples.
with flow-cytometric quantification and a natural microbial
consortium as a means of improving the AOC bioassay.
In this study (e.g., Figure 2) it was shown that flow
cytometry can be used in combination with the conventional
bioassay (P-17, 15 °C) instead of other enumeration methods.
While plating is relatively easy and does not require specific
equipment, flow cytometry is more reproducible and less
susceptible to variations in culturability of the cells, and, as
an additional benefit, the cell count is obtained within
15 min (10 min staining, 5 min analysis) instead of after 1-3
days, as are required for the incubation of the plates. Total
nucleic acid fluorescence staining and flow cytometry also
allows the detection and enumeration of all bacteria in a
sample, including those that are inactive or unculturable.
This means that growth is correctly assessed as all organisms
that proliferated in the sample during the assay and not only
those that are both viable and culturable at the sampling
end points. For example, large fluctuations in plate count
results were reported by LeChevallier et al. (6) when P-17
was grown on acetate at 30 °C. In fact, using acridine orange
direct microscopic counts, Kaplan et al. (5) demonstrated
that only between 60 and 80% of the cells in pure-culture
AOC assays are culturable and that this might affect the
determined AOC value when predetermined yield values are
used. Compared to ATP analysis, flow cytometry is more
sensitive and less prone to background interference, and it
analyses specifically the numerical growth of a culture, which
is the fundamental principle of the AOC concept, rather than
a cell-associated parameter (ATP). This is of particular
significance since different bacteria can have widely different
intracellular ATP concentrations during the various stages
of growth. For example, LeChevallier et al. (6) reported nearly
10-fold differences in mean ATP values of P-17 (1.85 fg cell-1)
and NOX (0.213 fg cell-1), respectively. Moreover, the use of
ATP analysis requires two separate conversions to achieve
AOC data: a first conversion from total ATP to total cell
numbers and a second conversion from total cell numbers
to AOC.
The use of a pure culture for detection of natural AOC is
contentious as such. Indeed, although P-17 was initially
recognized and used as a typical heterotrophic strain
representing drinking water bacteria, Spirillum strain NOX
was subsequently added to the AOC determination method
as it was discovered that P-17 does not adequately detect
ozonation products (1). Flavobacterium sp. strain 12 has also
been suggested for specific waters (targeting sugars in
particular), while several other natural isolates have also been
tested (1, 11). However, because AOC covers a wide range
of assimilable substrates including for example organic acids,
sugars, alcohols, amino acids, and oligopeptides in various
molecular sizes, a pure culture will probably never suffice to
detect all of these (12). Using combinations of these pure
cultures as a mixed inoculum has been examined but presents
experimental problems in terms of both the enumeration
and interpretation of the results (1). As a result, researchers
often prefer to use only one of these strains (5). Even though
it was not specifically tested in this work, it is possible to
measure with flow cytometry analysis any other pure culture
or combinations of two or more pure cultures in the AOC
bioassay. In the latter case it should be noted that the total
nucleic acid staining method described in this study would
suffice only for the detection of the total number of grown
cells. However, application of fluorescent in situ hybridization
(FISH) probes and fluorescent antibodies together with flow
cytometry would enable the separation and enumeration of
the individual subpopulations of a natural consortium (9).
Though the latter option may not yet be lucrative for industrial
applications, it does offer immense research opportunities
in the field of microbial ecology, e.g., following microbial
population dynamics in the presence of specific AOC
compounds.
An alternative approach to pure cultures is to use a natural
microbial consortium. Such a consortium possesses a much
broader and diverse substrate range than a single pure culture
and should thereby inherently be able to offer a more realistic
interpretation of the actual AOC content when natural
substrates are assayed. This approach adds the possibility to
use an in situ natural microbial consortium in the AOC
analysis, in other words, a natural microbial consortium
autochthonous to the water being analyzed and, therefore,
adapted to the types of AOC in such a water sample. Flow
cytometry is a straightforward method whereby a natural
microbial consortium can be enumerated accurately, obviating the problem of nonculturability. This work demonstrated that when a natural microbial consortium, which
was adapted to 30 °C, was used in the assay, stationary phase
end-point data could be achieved within 30-40 h following
inoculation. As such, this is significantly faster than any other
previously reported method (6). It remains to be tested
extensively the degree to which different natural microbial
consortia would produce similar AOC results on the same
water. Preliminary experiments in this regard with four
inoculums produced from different sources showed no
statistical difference in the predicted AOC value (data not
shown).
Using a natural microbial consortium, however, poses a
question as to how microbial growth should best be related
to carbon equivalents, because standardizing the yield of a
natural consortium on natural AOC entails a rather complicated argument. Empirical yield values for pure culture
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growth on acetate are used in the conventional AOC method,
but these are subject to several factors. First, these yield factors
are dependent on temperature (15 °C in the conventional
method) and require recalculation if a different incubation
temperature is used (6, 7). Second, the yield values differ
significantly for different microbial strains when used under
otherwise similar conditions. The best examples of this are
P-17 and NOX, which have yield values of 4.1 × 106 and
1.2 × 107 cfu (µg of acetate-C)-1 respectively. Third, the yield
values will also differ significantly when different pure carbon
sources are used. For example, NOX yields 1.2 × 107 cfu
(µg of C)-1 on acetate, while only yielding 2.9 × 106 cfu
(µg of C)-1 on oxalate (1). Kaplan et al. (5) illustrated with
DOC mass balances during AOC measurements that P-17
yield on acetate is about 1.25 times lower than its yield during
growth on complex, natural substrates. In fact, Van der Kooij
(1) reported various yield values for five different pure cultures
growing on various carbon sources (acetate, oxalate, glucose,
starch, lactate) ranging between 2.9 × 106 and 1.2 × 107 cfu
(µg of C)-1. This latter point is even further complicated by
the use of empirical yield values derived from growing a
preconditioned pure culture on a single pure carbon source,
to convert growth on natural complex AOC to carbon
equivalents. Alternative to the above experimental approach,
a theoretical argument could have its merit. Batte et al. (13)
suggested the average carbon content of a bacterial cell to
be 2.0 × 10-14 g. From this value it can be deduced that 1 µg
of carbon equals 5.0 × 107 cells. Therefore, if 1 µg of AOC is
utilized, and for example 50% thereof is actually assimilated,
then the theoretical yield would be 2.5 × 107 cfu (µg of AOC)-1.
However, also this argument is fundamentally flawed, since
microbial cells sizes and carbon content differ tremendously,
as does the percentage of carbon being assimilated by the
cells (14). Hence, for the sake of simplicity, we recommend
the use of a constant value of 1 × 106 bacteria (µg of AOC)-1
as suggested by Van der Kooij (1) for the conversion of natural
microbial consortium growth to AOC values, representing a
value close to the maximum yield of bacteria on AOC (1). An
up to now untested argument which needs to be considered
is that cellular size might vary for different bacteria grown
on different AOC substrates. The implication thereof is that
total yield should be a function of both the cell numbers and
the cell size. However, first flow cytometric measurements
of cellular size of the natural microbial consortium used in
this study have revealed no significant differences in cell size
between the inoculum and those cells that have entered the
stationary phase after all AOC was consumed, irrespective
of the water that was tested (data not shown).
The proposed AOC method could easily be automated,
and for this purpose it has been patented (10). Yet, even
without automation, the simplicity with which the flowcytometry-based method can measure a large number of
samples enabled the recording of batch growth curves as
depicted in Figures 4 and 5A. This approach has considerable
additional benefits over the only end-point measurements
of the conventional method, as it provides a more complete
and realistic interpretation of microbial growth on AOC. For
example, growth inhibitory substances, AOC quality (type),
and AOC quantity all have profound effects on the growth
kinetics. With regard to the latter, it was initially explored
whether the kinetic approach could be used as a tool to
predict the AOC concentration of a sample, based on the
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ENVIRONMENTAL SCIENCE & TECHNOLOGY / VOL. 39, NO. 9, 2005
specific growth rate, even before the stationary phase is
reached. This notion is supported to some extent by the
results depicted in Figure 5A,B for growth on natural AOC.
It has, however, to be taken into account that numerous
factors apart from the carbon concentration could influence
the specific growth rate of a natural microbial population in
a water sample from the environment or from a technical
system. This kinetic approach adds value not only to the
field of AOC measurement but also to the general study of
natural microbial growth under carbon-limited conditions.
Acknowledgments
We appreciate the critical evaluation of the manuscript by
Dr. Martin Rieger and technical assistance from Verena Loser.
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Received for review November 5, 2004. Revised manuscript
received February 1, 2005. Accepted February 14, 2005.
ES048277C