1. science
2. ecology

Journal of Microbiological Methods 55 (2003) 865 – 874
www.elsevier.com/locate/jmicmeth
Activity assessment of microorganisms eluted from sediments
using 5-cyano-2,3-ditolyl tetrazolium chloride: a quantitative
comparison of flow cytometry to epifluorescent microscopy
Cyndee L. Gruden *, Ann Khijniak, Peter Adriaens
Environmental and Water Resources Engineering, University of Michigan, Ann Arbor, MI 48104, USA
Abstract
Enhanced natural recovery may be successfully implemented at contaminated sediment sites, which are often characterized
by large volumes of sediments with low to moderate levels of contamination to cost-effectively reduce human and ecological
risks. In order to evaluate the potential for microbial contribution to remediation strategies, physiological assessment of
indigenous microorganisms is essential. We report here a method for rapid and accurate assessment of metabolically (5-cyano2,3-ditolyl tetrazolium chloride [CTC]) active microorganisms eluted from sediment, based on flow cytometry (FCM).
Microorganisms eluted from sediment and suspended in estuarine medium were stained with CTC and counterstained with the
DNA stain Picogreen (PG). Optimal stain concentrations and incubation times were employed. FCM quantification of the dualstained microorganisms was not statistically different (paired t test; a = 0.05; df = 10) from enumeration (total or active numbers)
by an established method (fluorescent microscopy) over two orders of magnitude (approximately 104 – 106/ml). This research
suggests that FCM, which allows the collection and analysis of multiple parameters (light scatter and fluorescence emission), is
a good candidate for microbial characterization in complex environmental matrices, such as sediments, across a broad range of
activity levels (f 2% to 84% of total). Potential applications for this FCM-based method include the rapid assessment of
changes in sediment microbial activity in response to enhanced bioremediation strategies.
D 2003 Elsevier B.V. All rights reserved.
Keywords: CTC; Sediments; Flow cytometry; Anaerobic
1. Introduction
Contaminated freshwater, estuarine, and marine
sediments are a priority of national concern, due to
potentially adverse impacts on aquatic ecosystems,
human health, and economic development. For the
vast majority of contaminated sediment sites, biore* Corresponding author. Department of Civil Engineering,
University of Toledo, 2801 West Bancroft Street, Toledo, OH
43606, USA. Tel.: +1-419-530-8120; fax: +1-419-530-8116.
0167-7012/$ - see front matter D 2003 Elsevier B.V. All rights reserved.
doi:10.1016/j.mimet.2003.08.005
mediation technologies aimed at degrading and sequestering contaminants may represent an effective
and affordable sediment management approach. For
the development of potential bioremediation strategies, an accurate appraisal of the physiological state
of indigenous subsurface bacteria is essential (Yu et
al., 1995). Direct optical detection methods such as
epifluorescent microscopy (EPM) and flow cytometry
(FCM), which utilize stains specific to biological
molecules (e.g., DNA stains), are increasingly being
employed for microbial characterization due to the
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C.L. Gruden et al. / Journal of Microbiological Methods 55 (2003) 865–874
awareness of problems associated with culture-based
assays (Vives-Rego et al., 2000). FCM is advantageous because fluorescently stained microbial samples can be simultaneously analyzed for multiple
parameters (e.g., red, yellow, green fluorescence
emission and light scatter) (Kalmbach et al., 1997;
Nebe-von-Caron et al., 2000). In addition, FCM
detectors (photomultiplier tubes [PMTs]) provide superior sensitivity as compared to EPM. Despite the
speed, sensitivity, and reproducibility of FCM, applications have been limited in subsurface microbial
characterization.
Numerous stains have been developed for the
assessment of cell function including reproductive
ability, metabolic activity, membrane activity, and
membrane potential. These methods have been
employed to distinguish viable, but nonculturable,
microorganisms that would be discounted in traditional, growth-based assays. Several of these probes are
compatible with flow cytometry including: (i) lipophilic cationic dyes (e.g., Rhodamine 123-accumulated on inner side of energized membranes), (ii) DNA
intercalators (e.g., propidium iodide-crossed only altered cell membranes), (iii) redox dyes (e.g., 5-cyano2,3-ditolyl tetrazolium chloride [CTC]-reduced during
respiration to form fluorescent intracellular precipitate) (Lopez-Amoros et al., 1997; Nebe-von-Caron et
al., 2000). By simultaneously staining with selected
multiple fluorochromes (e.g., non-specific stains and
activity stains), FCM analysis can reveal the functional heterogeneity of the microbial sample (Kalmbach et
al., 1997; Nebe-von-Caron et al., 2000). Rapid, highthroughput microbial techniques, such as FCM, that
accurately quantify metabolic activity are of interest in
the bioremediation field due to the need for spatially
diverse analysis of ecological parameters in a wide
range of applications.
For indicating metabolically active cells, the use of
tetrazolium salts, such as CTC, is beneficial due to
rapid reduction (minutes to hours) to formazan by
intracellular reductase enzymes. The mechanisms for
biotic CTC reduction under aerobic conditions have
been researched extensively (Lopez-Amoros et al.,
1995; Smith and McFeters, 1997). Although CTC
application under anaerobic conditions has been
documented in the literature (Smith and McFeters,
1997; Bhupathiraju et al., 1999a; Proctor and Souza,
2001; Gruden et al., 2003), a comprehensive under-
standing of the anaerobic processes involved in CTC
reduction has yet to be disclosed (Bhupathiraju et al.,
1999a; Gruden et al., 2003). To date, CTC has been
used to assess microbiological activity in complex,
anaerobic environmental matrices including wastewater, sludge, soil, and sediments (Rodriguez et al.,
1992; Winding et al., 1994; Yu et al., 1995, Proctor
and Souza, 2001; Gruden et al., 2003; Ziglio et al.,
2002). For example, CTC was used to quantify
fluctuations in microbial activity in response to varying environmental conditions (e.g., nutrients) in
wastewater applications including activated sludge
and anaerobic digester sludge (Gruden et al., 2003;
Ziglio et al., 2002).
The fluorescent precipitate formed by CTC reduction has been quantified in whole cells using optical
detection methods such as direct microscopy (Rodriguez et al., 1992, Pyle et al., 1995; Gruden et al.,
2003). Subsurface microbial samples present challenges to optical detection methods due to an abundance of non-living bacterial sized particles and
inherent background fluorescence (Yu et al., 1995;
Proctor and Souza, 2001). As compared to other redox
dyes, CTC works well in complex matrices because
the fluorescent red intracellular precipitate is easily
detected and retains its fluorescence upon storage for
up to 2 days (Yu et al., 1995). Potential applications
for this method include the rapid determination of
microbial contribution to remediation of subsurface
contaminants through the comparison of CTC-active
bacterial counts from samples at contaminated sites to
samples from geologically similar but uncontaminated
background sites (Bhupathiraju et al., 1999b). This
manuscript details a CTC-based flow cytometric
method for assessing the metabolic activity of sediment microorganisms and provides a comparison to
an established direct count protocol.
2. Materials and methods
2.1. Sediment sample preparation
Microorganisms were eluted, using an established
protocol (Barkovskii and Adriaens, 1996, 1998) from
Passaic River, NJ, Pearl Harbor, HI, and San Diego
Bay, CA, sediments (VS = 10 – 20%). The eluted
microorganisms were suspended in estuarine medium
C.L. Gruden et al. / Journal of Microbiological Methods 55 (2003) 865–874
(40 mM phosphate buffer (KH 2 PO 4 /Na 2 HPO 4 ;
pH = 7.2); 2.0 g/l NaHCO3; 0.9 g/l (NH4)2Cl; 0.07
g/l CaCl26H2O; 0.13 g/l MgSO47H2O; 0.02 g/
l FeCl24H2O; cysteine – HCl; trace minerals; vitamin
stock; resazurin). In order to evaluate the impact of
organic acid (OA) amendment and incubation time on
microbial activity, some samples were enriched with
organic acid cocktail (45 mg/l acetate; 30 mg/l propionate; 15 mg/l butyrate; 10 mg/l benzoate) and
incubated for up to 7 days while constantly stirring
at room temperature prior to analysis. All transfers,
enrichments, amendments, and incubations were carried out in an anaerobic chamber. The percentages of
CTC-active microorganisms in OA amended samples
and estuarine medium only samples were compared as
a function of incubation time.
2.2. DNA stain
Picogreen (PG) (P-7589; Molecular Probes,
Eugene, OR), a proprietary, high-affinity, green,
nucleic acid dye (ex/em k = 502/523 nm), was stored
at 20 jC for up to 30 days. A range of PG
concentrations (5 10 4 to 1 10 2 of stock) and
incubation times (2 min to 4 h) were investigated for
determining optimum conditions (5 10 3 of stock
and 30 min) for enumeration of sediment microorganisms. After PG incubation, samples were split for
independent FCM and fluorescent microscope (EPM)
counts (Lange et al., 1997). In order to maximize
fluorescence emission, samples were not fixed prior to
analysis. In all instances, they were analyzed within
4 h of preparation.
2.3. Metabolic activity stain
The metabolic activity of the sediment microorganisms was determined using the tetrazolium redox dye
CTC (Polyscience, Warrington, PA) (ex/em k = 475/
600 nm). The CTC assays were performed using
filter-sterilized (0.2 Am) CTC stock solution (20
mM) prepared in ultrapure water and stored at 4 jC
for a maximum of 2 days (Rodriguez et al., 1992).
The appropriate final CTC concentration and incubation time (5 mM and 1 h) for flow cytometric analysis
were determined in optimization experiments (Smith
et al., 1994; Bhupathiraju et al., 1999b; Proctor and
Souza, 2001; Gruden et al., 2003).
867
2.4. Flow cytometry
The samples were analyzed using a FACSCaliburk
dual laser flow cytometer, with six parameters of
detection: forward scatter (FSC), sidescatter (SSC),
and four fluorescence intensity detectors (FL1, FL2,
FL3, FL4) (Becton Dickinson, San Jose, CA). The
specifications of the detector channels were as follows:
FL1 (green light: k = 515 – 545 nm); FL2 (yellow/
orange: k = 563 –607 nm); FL3 (dark red: k = 670 nm
LP); and FL4 (red: k = 653 –669 nm). An air-cooled
argon laser (488 nm) and red diode laser (635 nm)
were used for excitation. All parameters were collected
as logarithmic signals.
2.5. Calibration and instrument settings
Daily calibration was completed by analyzing unlabeled beads (to simulate unstained microorganisms)
and fluorescein isothiocyanate (FITC: yellow-green),
phycoerythrin (PE: red-orange), and peridinin chlorophyll protein (PerCP: red) beads (to simulate fluorochrome labeled microorganisms) (CaliBRITEk beads
[Cat. No. 349502], Becton Dickinson). The beads have
expected values for FSC, SSC, and fluorescence, and
these values are used to adjust instrument settings, to
set fluorescence compensation, and to check instrument sensitivity (BD FACSCompk software) before
any samples were processed. The unlabeled beads
were used to discriminate actual events from background debris using the FSC detector. The photomultiplier tube voltages were adjusted so that the
mean channel values of the beads corresponded to
the programmed target values. The threshold was
adjusted to minimize background noise.
Each fluorochrome emits light over a range of
wavelengths when excited by the laser. This spectral
overlap was corrected using electronic compensation.
Using a mixed suspension of equal amounts of
unlabeled, green, yellow, and red beads, the software
differentiated green-yellow signal from orange signal
and orange from red signals by aligning labeled bead
populations with the corresponding unlabeled bead
population. After the instrument settings were established, FL1, FL2, and FL3 sensitivity (mean channel
separation between the signal produced by labeled and
unlabeled beads) and FSC and SSC sensitivity (mean
channel difference between light scatter, beads signal,
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C.L. Gruden et al. / Journal of Microbiological Methods 55 (2003) 865–874
and background noise) were tested using the mixedbead suspension.
Initial instrument settings for each experiment
were: threshold-SSC; FSC-E01, logarithmic amplification; SSC-375, logarithmic amplification; FL1-600,
logarithmic amplification; FL2-550, logarithmic amplification, FL3-800, logarithmic amplification. An
unstained sample was processed prior to each run to
optimize the instrument settings. Settings were
deemed acceptable when the entire unstained population was visible on an FSC vs. SSC dot plot and the
bell-shaped histogram showed a distinct gap from the
noise signal on a histogram of both FSC and SSC
events. In addition, the voltages of FL1, FL2, and FL3
were adjusted using fluorescence as a function of SSC
to contain the target population within the first order
of magnitude on the logarithmic y-axis (100 to 101) to
correct for background fluorescence in the unstained
sample. The system threshold was set on SSC signals
and all analyses were performed at the low setting (12
Al/min). Enumeration was carried out using standard
procedures involving a lipophilized bead pellet that
dissolved, releasing a known concentration of fluorescent beads (TruCountk Tubes: 340567; Becton
Dickinson) and in accordance with previously published protocols (Lange et al., 1997). Data analysis
was performed manually using CellQuestk software
(Becton Dickinson).
2.6. Gating and control samples
A negative control (unstained sediment bacteria)
was used to define the background or negative region
on fluorescence contour plots. A positive control (PGand CTC-stained pure culture of bacteria) was used to
determine the location of the fluorescent signal or the
region expected for a positive response. SSC, which is
often considered a measure of the internal complexity
of the labeled cell, and fluorescent (FL1/FL3) emission were used simultaneously to separate the target
population (fluorescently labeled microorganisms)
from background particles. Negative and positive
controls were compared to stained samples. Events
that generated a fluorescent signal above the negative
region (background) were gated as the population of
interest. In addition, each experiment included controls such as PG only, CTC only, and PG and CTC
after microorganisms were exposed to sodium azide in
an effort to determine background fluorescence associated with non-specific binding of the selected stains
and to account for potential exocellular and/or abiotic
reduction of CTC.
2.7. Microscopic analysis
The Zeiss Axioplan Universal Microscope (451888,
Carl Zeiss MicroImaging, Thornwood, NY) was equipped with an epifluorescence attachment (448006, Carl
Zeiss), a super high-pressure mercury lamp power
supply (Carl Zeiss) and 50-W mercury lamp (HBO
50W L2, Toby’s Instrument Shop, Saline, MI) for
epifluorescence illumination. Fluorescent turret and
filter sets (Optavar Fl, 446421, Carl Zeiss) were standard with the Axioplan.
2.8. Filtration and enumeration
The fixation, filtration, and enumeration methods
as well as the detection limit used in this research
were adapted from widely accepted protocols and
were previously reported (Gruden et al., 2003). After
incubation on a shaker table at room temperature
(f 25 jC) in the dark, samples were filtered through
a 0.2-Am black polycarbonate filter (Poretics, Cat. No.
K02BP02500) backed with a 25-mm glass-fiber prefilter (11-394-106C, Fisher Scientific, Fair Lawn, NJ).
Filters were dried and mounted on a slide with
immersion oil and a cover slip and stored in the dark
prior to microscopic analysis. Triplicates were
counted. Laboratory controls were prepared including
dead (sodium azide) samples and method blanks. A
minimum of seven fields and 200 microorganisms
were counted per slide. Only microorganisms that
were both DNA stained (green) and contained red
intracellular precipitate were considered metabolically
active.
3. Results
The results from all three sediments studied in this
work were similar. The data presented in this manuscript represent sediment samples from the Passaic
River, NJ (Volatile Solids = 15%). Prior to each experiment, an unstained sample, which served as a
negative control, was analyzed to determine back-
C.L. Gruden et al. / Journal of Microbiological Methods 55 (2003) 865–874
869
stained (PG and CTC) pure culture was analyzed,
and both FL1 (green fluorescence) and FL3 (dark red
fluorescence) were plotted as a function of SSC. This
positive control sample was used to determine the
region (R1) of expected response from PG-stained
microorganisms (Fig. 2A) and region (R2) of expected
response from CTC-active microorganisms as a subset
of R1 (Fig. 2B).
Fig. 1. Control samples used to establish gating protocol. (A)
Contour plot of green fluorescence emission (FL1) vs. SSC for
unstained sediment microorganisms. R2 is defined as the gate or
region of interest where green fluorescent emission is expected to
occur. Contour plot of (B) green fluorescence emission (FL1) and
(C) dark red fluorescence emission (FL3) as a function of SSC for a
positive control sample (pure culture stained with both PG and
CTC). R1 delineates DNA containing (stained green) microorganisms. R2 contains metabolically active microorganisms (dark
red) that are a subset of the population defined by R1 only.
ground fluorescent response due to the sample matrix.
Fig. 1 is a contour plot of the unstained sample, and it
displays green fluorescence (FL1) as a function of
SSC. Background fluorescence is shown below 101
on the logarithmic FL1 axis. In addition, a dual-
Fig. 2. Contour plot of green fluorescence emission (FL1) as a
function of SSC for PG-stained sediment microorganisms. R1 is
defined as the gate containing sediment microorganisms (82.5% of
total events). Contour plot of dark red fluorescence emission (FL3)
as a function of SSC for CTC- and PG-stained sediment
microorganisms. R2 is defined as the gate containing metabolically
active sediment microorganisms (83.9% of events in R1).
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C.L. Gruden et al. / Journal of Microbiological Methods 55 (2003) 865–874
Green fluorescence (FL1) was plotted as a function
of SSC to discriminate the PG-stained sediment
microorganisms (Fig. 2A). In Fig. 3, the gated population (R1), which composed 82.5% of the total
events, had a uniform fluorescent intensity (FL1)
and SSC and was readily distinguished from background events (below 101 on the logarithmic scale of
FL1 and SSC axes). Metabolically active sediment
microorganisms fluoresced both dark red and green
when stained with CTC and PG. The dark red
fluorescent (FL3) events, a subset of those in R1
(green fluorescent events from Fig. 2A), were plotted
as a function of SSC (Fig. 2B). The subpopulation of
microorganisms that were stained by both PG and
CTC are circumscribed in R2 (83.9% of R1).
Optimization experiments were performed for
FCM analysis of CTC-stained sediment bacteria
(Fig. 3). No statistically significant change (paired t
test; a = 0.05) in % CTC activity was observed at
Fig. 4. The difference between the concentration of bacteria
determined by EPM and FCM plotted as a function of sample.
Samples are ordered from lowest to highest concentration (no./ml).
Total (5) and active (n) numbers of sediment bacteria are
represented. Each data point represents a set of triplicates. There
was no statistically significant difference between the total (5)
(paired t test; a = 0.05; df = 10; jt Statj = 0.33) or active (n) (paired t
test; a = 0.05; df = 10; jt Statj = 0.46) numbers of sediment microorganisms quantified using EPM and FCM.
concentrations greater than 5 mM CTC (Fig. 3A) or
after more than 1 h of incubation (Fig. 3B). Optimal
incubation conditions for microscopic analysis of
sediment bacteria were 5 mM CTC and 4 h of incubation (data not shown).
Sediment microorganisms stained with CTC and
counterstained with PG were enumerated using FCM
and compared to values estimated by an established
direct count method (EPM). PG-stained (total) sedi-
Fig. 3. Optimization of CTC concentration (A) and incubation time
(B) for sediment microorganisms analyzed using a benchtop flow
cytometer. These figures represent two independent experiments.
Error bars representing standard deviation of the mean are smaller
than symbols in some instances.
Fig. 5. Enumeration of sediment microorganisms using flow
cytometry as compared to epifluorescent microscopy. All samples
were stained with PG (5 10 3 of stock for 30 min) and
counterstained with CTC (5 mM for 1 h). Error bars represent
standard deviation of the mean. In some instances, error bars are too
small to be seen.
C.L. Gruden et al. / Journal of Microbiological Methods 55 (2003) 865–874
ment microorganisms are bright green. Microorganisms that are considered metabolically active as
judged by CTC contain a red fluorescent intracellular
precipitate. In order to determine if the two methods
(EPM and FCM) for enumerating both total and active
sediment bacteria produced different results, the average of the differences was assessed (Fig. 4) (Berthoux and Brown, 1994). In Fig. 4, samples were
plotted in ascending order (1 – 6) based on total
concentration (EPM). At highest and lowest concentrations (samples 1 and 6), FCM appears to overestimate the number of bacteria, and the lowest
concentration demonstrates the greatest difference
between values (0.53). There was no statistically
significant difference (paired t test; a = 0.05; df = 10;
jt Statj = 0.33) between the total number of microorganisms quantified using EPM and FCM. In addition, the number of CTC-active organisms determined
using FCM was not statistically different than the
number determined using EPM (paired t test; a = 0.05;
df = 10; jt Statj = 0.46). Flow cytometry and epifluorescent microscopy provided comparable quantification of CTC-active microorganisms across two orders
of magnitude (104 –106 cells/ml) (Fig. 5).
The percent of CTC-active microorganisms eluted
from sediment with and without OA amendment was
plotted as a function of incubation time (Fig. 6). OA
addition resulted in a statistically significant increase
in percent CTC active from approximately 16– 72%
after 12 h of incubation (Fig. 6). No statistically
Fig. 6. Comparison of the percent of CTC active sediment
microorganisms with and without OA amendment (45 mg/l acetate;
30 mg/l propionate; 15 mg/l butyrate; 10 mg/l benzoate) as a
function of incubation time.
871
significant difference was noted in total numbers of
sediment microorganisms for either treatment at any
incubation time (data not shown). After several days
of incubation (7 days), the percent of CTC-active
microorganisms of the OA amended sample returned
to a baseline value of approximately 4% CTC active.
4. Discussion
PG distinguished the microbial population from
sediment background fluorescence, as compared to an
unstained sample. Other stains tested (e.g., SYTO BC,
Molecular Probes) resulted in non-specific staining of
the background and an inability to distinguish the
microorganisms (data not shown). As cited in previously reported sediment studies using direct count
methods (Weinbauer et al., 1998; Griebler et al.,
2001), high-affinity green nucleic acid dyes, such as
PG, effectively achieved separation of the community.
Optimum PG incubation conditions (3 10 5 of
stock and 30 min) for enumeration of microorganisms
eluted from sediment (data not shown) were in the
range of those previously reported (Weinbauer et al.,
1998).
Several authors have successfully distinguished
CTC-active microorganisms in aqueous environmental samples using FCM (Kaprelyants and Kell, 1993;
del Giorgio et al., 1997; Sieracki et al., 1999). Proctor
and Souza (2001) employed epifluorescent microscopy to enumerate CTC-active sediment microorganisms. However, FCM analysis of subsurface
microorganisms has been considered difficult due to
red autofluorescence (Davey and Kell, 1996). In order
to eliminate interferences associated with red background fluorescence, the detection of sediment microorganisms was carried out using a green stain (PG) for
microbial identification in this work. Gating protocols
were adapted from Sieracki et al. (1999). CTC-active
sediment microorganisms were readily separated from
red autofluorescence (background) since only green
events were further analyzed for CTC (dark red)
activity. In addition, fluorescence intensity of labeled
microorganisms was maximized because samples
were not fixed prior to analysis, which occurred
directly following ( < 4 h after) preparation. Preliminary tests indicated a 50% decrease in fluorescent
signal following sample fixation (1% formaldehyde
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C.L. Gruden et al. / Journal of Microbiological Methods 55 (2003) 865–874
for 30 min) (data not shown) as previously reported
(Lebaron et al., 1998).
The CTC optimization values determined for this
work (5 mM CTC and 4 h of incubation), used for
both FCM and EPM analyses, are in agreement with
previously published protocols for anaerobic microorganisms in soils and sediments (Proctor and Souza,
2001; Bhupathiraju et al., 1999b; Yu et al., 1995). As
expected, the optimum incubation time for FCM
analysis is significantly shorter (1 h) than for epifluorescent microscopy (4 h) because the FCM detectors
(photomultiplier tubes), which are considerably more
sensitive than EPM detectors (e.g., the human eye)
(Sieracki et al., 1999), can identify lesser amounts of
reduced formazan without difficulty.
FCM analysis (up to 103 cells/s) of dual-stained
sediment microorganisms was 5 – 10 times faster than
EPM analysis (approximately 1 cell/s) when the time
required for sample preparation and instrument maintenance was incorporated. The amount of time saved
per sample increased with increasing number of
samples. FCM analysis eliminated the need to prepare
slides over several dilutions for accurate counting, the
laborious process of counting individual fields for
each individual fluorescent stain, and the effort associated with manual data reduction.
Total and active microorganisms were accurately
quantified using standardized counting beads using a
previously published protocol (Lange et al., 1997).
The greatest difference between the two methods
occurred at the lowest concentration of active microorganisms (f 1 104/ml) indicating the possibility
for increased error. This concurs with preliminary
laboratory data that indicated poor reproducibility
and accuracy in instances where the concentration
was outside of the acceptable signal to noise range
as determined during instrument calibration with
unstained samples (data not shown). The quantification of dual-stained sediment microorganisms by
FCM corresponded to EPM numbers over two orders
of magnitude (approximately 1 104 to 1 106/ml).
In aqueous environmental samples, the concentration
for accurate quantification of microorganisms, as
compared to EPM, was in a similar range (5 105/
ml to 5 106/ml) (Sieracki et al., 1999; del Giorgio et
al., 1997).
A statistically significant increase in % CTC
activity occurred after elution (with or without OA
amendment), indicating metabolic stimulation (twoto fourfold) due to the elution protocol and subsequent suspension in estuarine medium only. In addition, OA amendment in the elution protocol
(enrichment) significantly increased the number of
CTC-active microorganisms only (total number of
organisms did not increase) after several hours (12
h) of incubation. The amendment of OA is a common
strategy to enhance metabolic activity in sediments
prior to experimentation or to stimulate in situ activity
for remediation purposes (Barkovskii and Adriaens,
1996). However, the implications of this protocol on
microbial viability and respiration have not been
quantified and are likely to be site specific. These
results underline the requirement for a consistent
elution protocol between samples and individual
experiments in order to compare resulting data. These
results also demonstrate that OA amendment is an
ineffective method for increasing microbial activity in
the short term ( < 12 h) for these samples and the
specific OA cocktail. This data indicates that the
FCM-based method can accurately measure percent
CTC activity of sediment microorganisms across a
broad range (2.1 –83.8% CTC active).
5. Conclusions
Samples with varying sediments and biogeochemistry may require individualized protocols. Specific
protocols should be developed including optimum
incubation conditions for each of the selected fluorescent stains, microbial elution/enrichment methods,
and appropriate FCM operating parameters. Several
controls are required to distinguish the target cells and
physiological stains from background fluorescence
including positive and negative controls. If possible,
sample fixation should be avoided to maximize the
fluorescent intensity of stained microorganisms, facilitating the distinction between microorganisms and
inert particles.
Although CTC use may be considered contentious
(due to opposing views on the operational definition
of active microorganisms) (Creach et al., 2003; Servais et al., 2001; Choi et al., 1999; Kaprelyants and
Kell, 1993), it has been effectively applied as an
indicator of variation in metabolic activity in complex
matrices (e.g., biosolids and sediments) (Gruden et al.,
C.L. Gruden et al. / Journal of Microbiological Methods 55 (2003) 865–874
2003; Proctor and Souza, 2001; Bhupathiraju et al.,
1999b). CTC has previously been employed to detect
enhanced microbial activity corresponding to zones of
organic contamination (Bhupathiraju et al., 1999b;
Lopez-Amoros et al., 1998). This research demonstrates the utility of CTC activity assessment of
sediment microorganisms using flow cytometry across
a broad range of activity levels. This rapid and
accurate method may be applied to evaluate the
potential for microbial contribution to bioremediation
in contaminated sediments.
Acknowledgements
This work was supported in part by the NOAACooperative Institute for Coastal and Estuarine
Environmental Technology (CICEET). Tom Komorowski in the School of Public Health at the University
of Michigan and Jordan Peccia in the Department of
Civil and Environmental Engineering at Arizona
State University assisted in the preparation of the
manuscript.
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