APPLIED AND ENVIRONMENTAL MICROBIOLOGY, May 2003, p. 2994–2998
Copyright © 2003, American Society for Microbiology. All Rights Reserved.
Vol. 69, No. 5
Community-Level Physiological Profiling Performed with an
Oxygen-Sensitive Fluorophore in a Microtiter Plate
Jay L. Garland,1* Michael S. Roberts,1Lanfang H. Levine,1and Aaron L. Mills2
Dynamac Corporation, Kennedy Space Center, Florida 32899,1and Laboratory of Microbial Ecology,
Department of Environmental Sciences, University of Virginia, Charlottesville, Virginia 229042
Received 15 July 2002/Accepted 7 February 2003
Community-level physiological profiling based upon fluorometric detection of oxygen consumption was
performed on hydroponic rhizosphere and salt marsh litter samples by using substrate levels as low as 50 ppm
with incubation times between 5 and 24 h. The rate and extent of response were increased in samples
acclimated to specific substrates and were reduced by limiting nitrogen availability in the wells.
Characterization and classification of heterotrophic micro-
bial communities based on rapid assessment of multiple sole-
carbon-source use, termed community-level physiological pro-
filing (CLPP) by Lehman et al. (9), were developed over 10
years ago (6). Although CLPP is a valuable method for assess-
ing relative change in microbial communities when properly
applied (3, 12), limitations due to selective enrichment result-
ing from the high substrate concentrations (i.e., ?100 mM)
and long incubation periods (typically 1 to 4 days) restrict its
present use for assessing actual differences in the physiological
capabilities of microbial communities (2, 8, 14, 16). Develop-
ment of more functionally relevant CLPP will depend on the
application of new detection methods that allow for rapid and
direct (e.g., O2consumption or CO2production) rather than
indirect (e.g., tetrazolium dye reduction) assessment of respi-
ration, with decreased requisite incubation times and substrate
concentrations. In addition, the use of undefined, proprietary
reagents should be minimized so that incubation conditions
can be defined and manipulated by the investigator.14C respi-
rometric approaches limit enrichment effects by allowing for
detection of physiological response with addition of very low
substrate concentrations (i.e., nanomolar levels), but even sim-
plified assay systems (13) still have the expense associated with
disposal of radioactive wastes. CLPP approaches based on CO2
monitoring that limit incubation time have been developed (1)
but require high substrate concentrations (i.e., 10 to 100 mM).
Tracking the rate of O2consumption in the test medium may
improve sensitivity, given the relatively low concentration and
solubility of O2in water. A recently developed, fluorescence-
based microplate platform for assessing dissolved oxygen (BD
Oxygen Biosensor System; BD Biosciences, Bedford, Mass.)
(17) could enable rapid testing of multiple substrates; the ef-
fectiveness of this system to detect known shifts in substrate
utilization by mixed microbial communities is the subject of
this work. The BD Oxygen Biosensor System is based on an
O2-sensitive fluorophore, 4,7-diphenyl-1,10-phenathroline ru-
thenium (II) chloride, absorbed into a silicone matrix that is
permeable to O2(17). The fluorescence of the ruthenium dye
is quenched by the presence of O2, so the signal from the
fluorophore-gel complex loaded on the bottom of the micro-
plate wells increases in response to respiration in the overlying
sample. In this work, samples of environmental systems sus-
pended in sterile phosphate-buffered mineral salts (PBMS) (7
g of K2HPO4liter?1, 3 g of KH2PO4liter?1, 0.1 g of MgSO4
liter?1, 0.5 g of [NH4]2SO4liter?1, 0.01 g of CaCl2liter?1,
0.005 g of FeSO4liter?1, 0.0025 g of MnSO4liter?1, and 0.0025
g of Na2MoO4liter?1) were inoculated into the wells.
In the first part of the work, the fluorescence responses of
rhizosphere communities to two different types of surfactants
were related to their previous exposure to these compounds
and to independently collected data on surfactant degradation.
These experiments were part of an ongoing project evaluating
direct addition of human-hygiene water into hydroponic plant
systems as an approach for water recycling on extended space
missions, and specific description of the plant growth system is
presented elsewhere (4). Briefly, wheat (Triticum aestivum L.
cv. USU-Apogee) was grown by using nutrient film technique
hydroponic culture inside a controlled environmental growth
chamber. One hydroponic system served as a control with no
continuous addition of surfactants, while the other systems
received either sodium laureth sulfate (SLES) as Rhodapex
ES-2 (25% SLES; Rhodia Inc., Cranbury, N.J.) or cocamido-
propyl betaine (CAPB) as Mirataine BET C-30 (30% CAPB;
Rhodia, Inc.), beginning 4 days after planting. Surfactant stock
solutions (200 ml of 2,000 ppm) were added to the tanks in a
continuous mode via a peristaltic pump over a 23-h period.
Twenty days after planting, suspensions of rhizosphere com-
munities were obtained by excising sections (?1 by 1 cm) of
root mat from the front of the trays and by hand shaking in 25
ml of sterile PBMS containing 2-mm glass beads for 2 min. A
single rhizosphere sample was obtained from each treatment,
and undiluted and diluted (1/5 and 1/25) rhizosphere suspen-
sions were inoculated into the BD microplates and were read
on a Dynex MFX Microplate Fluorometer at 485-nm excita-
tion and 604-nm emission wavelengths with the top-reading
mode. Plate contents were incubated at 30°C, and readings
were obtained every 15 min for 120 h. Plates were not shaken
during the incubation period. On day 21, a single pulse of 400
ml of a 1,000-ppm SLES and 1,000-ppm CAPB solution was
added to all treatments to determine surfactant decay rates
* Corresponding author. Mailing address: Mail Code DYN-3,
Kennedy Space Center, FL 32899. Phone: (321) 476-4276. Fax: (321)
853-4165. E-mail: firstname.lastname@example.org.
based on analysis of nutrient solution for SLES by using ion
pair chromatography with suppressed conductivity detection
(11) or for CAPB by using high-pressure liquid chromatogra-
phy linked to electrospray ionization-mass spectrometry (10).
Samples from acclimated systems produced a more pro-
nounced peak in fluorescence more rapidly when incubated in
the BD systems at the 50-ppm level of surfactant (Fig. 1A and
C; Table 1), corresponding to the higher rates of degradation
in the same hydroponic systems as independently assessed by
chemical analysis (Table 2). Both the extent and rate of fluo-
rescence were dependent on inoculum density (Table 1), so
comparisons among treatments were made with samples of
approximately equivalent density (5.38 to 5.58 log cells ml?1
based on acridine orange direct counts  [Fig. 1]). For exam-
ple, the extent of response (i.e., peak response) to 50 parts of
SLES per million was greater in rhizosphere samples from
SLES-acclimated systems (2.58 normalized relative fluorescent
units [NRFU]) than in samples from the unacclimated (1.61
NRFU) or CAPB-acclimated (1.71 NRFU) systems and the
rate of response (i.e., time to peak) was faster in the SLES-
acclimated samples (12.5 h) than in the unacclimated (20.75 h)
or CAPB-acclimated (19.75 h) samples. The greater SLES
utilization in the BD microplate assay corresponds to a higher
rate of SLES disappearance within the SLES-acclimated sys-
tem (3.61 ppm h?1) than in the unacclimated or CAPB-accli-
mated systems (0.11 to 0.12 ppm h?1). Similarly, samples from
the CAPB-acclimated system showed a greater response to-
ward 50 parts of CAPB per million in the BD microplate assay
than did samples from either the unacclimated or SLES-accli-
mated systems (Fig. 1B and D), corresponding to a higher rate
of CAPB disappearance from the CAPB-acclimated systems.
Biofilm samples removed from polyvinyl chloride coupons in-
cubated in the nutrient delivery tanks showed a fluorescent
response to acclimation similar to that seen with the rhizo-
sphere samples (data not shown).
At the 500-ppm level, interpretation of the fluorescence
response was less clear. The lag in SLES response remained
shorter in acclimated samples as estimated by various means,
but little difference existed in the peak value due to the as-
ymptotic nature of the response, apparently due to the main-
tenance of low dissolved O2as a result of sustained respiration
(Fig. 1B). The asymptotic level of fluorescence (?8 NRFU)
was the maximal fluorescence response as determined from
sodium sulfite (100 mM) controls. Differences among samples
were ambiguous at the 500-ppm level of CAPB due to pro-
nounced secondary peaks and overall complexity in the signal
(Fig. 1D). While the degradation pathway of the CAPB mol-
ecule is not well defined, multiple steps (with different reaction
rates) are likely required for complete oxidation of the entire
molecule and probably cause the multiple peaks (15).
FIG. 1. Fluorescence response of rhizosphere samples to 50 parts of SLES per million, 500 parts of SLES per million, 50 parts of CAPB per
million, and 500 parts of CAPB per million. Data are presented for samples from hydroponic systems previously exposed to no surfactant (control),
SLES, and CAPB. Inoculum density is approximately equivalent (5.38 to 5.58 log cells ml?1) for all samples. Origin of y axis equals 0.75 NRFU.
VOL. 69, 2003CLPP PERFORMED WITH AN OXYGEN-SENSITIVE FLUOROPHORE2995
Various response parameters describing either the delay
(i.e., minimum response time, time to peak, or lag estimated
from the log-linear model) or peak in fluorescence appear
suitable for comparison of samples (details in Table 1). Certain
models (i.e., logistic model) are only applicable to the higher
substrate concentration (500 ppm) in which an asymptotic level
of fluorescence is observed, while definition of both the extent
and rate of peak response is more readily applied to the lower
substrate concentrations (50 ppm) due to the clear definition
of a peak (visualization of response curves in Fig. 1). Further
studies are needed to assess the most suitable variables for
consistently discriminating among communities; the various
parameters reported in Table 1 reflect a number of the differ-
ent analytical approaches to evaluating the microplate data.
Lower substrate concentrations are preferred in physiologi-
cal assays, given the lower potential for selective enrichment,
and in this study the 50-ppm level provided more easily inter-
pretable data. Substrate concentrations below 50 ppm (i.e., 5
and 10 ppm) were tested but did not produce a detectable
response (data not shown).
A stimulatory effect of acclimation was not observed when
rhizosphere communities were incubated in Biolog plates con-
taining SLES or CAPB as a sole carbon source. Rhizosphere
samples were obtained during three replicate studies and were
prepared as described above, with the exception that sterile
deionized water rather than PBMS was used as a diluent, given
the concentration of nutrients already present in Biolog plates,
and was inoculated into Biolog MT plates. The absorbance was
negligible after 72 h of incubation for both surfactants at 5,000
and 50 ppm for all treatments (Table 3). The response was
greater at 500 ppm for both surfactants, but no consistent
differences were apparent between the acclimated and unac-
climated systems (Table 3). The lack of correlation between
the response in the Biolog CLPP approach and known differ-
ences in specific carbon source utilization has been previously
Follow-up tests were performed with salt marsh plant litter
decomposition communities to determine (i) if detectable pat-
terns of response could be produced for a variety of readily
assimilated substrates (i.e., amino acids, monosaccharides, and
organic acids) and (ii) if the extent of response was sensitive to
relevant biochemical factors that affect substrate utilization
(i.e., N availability). Litter bags (20 by 20 cm) filled with ?20
g of locally collected, air-dried black needle rush, Juncus ro-
emarianus, were placed in a restored marsh within the Merritt
Island National Wildlife Refuge in July 2001 and were sampled
after 42 days of incubation. Suspensions produced by blending
in 0.85% NaCl were diluted 1:10 into PBMS (to reduce the
particulate content, yielding an inoculum density of ?1.5 ? 106
TABLE 1. Response of samples from the rhizosphere of hydroponically grown wheat to SLES in the BD Oxygen Biosensor System
(log cells ml?1)c
CAPB acclimated 50
aSamples obtained from systems receiving no surfactant (unacclimated) or either SLES or CAPB.
bConcentration of SLES added to the wells within the BD Oxygen Biosensor plates.
cSingle samples removed from each system were inoculated into microplates after no, 1/5, or 1/25 dilution, resulting in three different inoculum densities.
dTime for normalized relative fluorescence to increase by 10% (i.e., to 1.1).
eMaximum normalized relative fluorescence.
fPeak area calculated with Sigma Plot function.
gResults from linear regression analysis of log-transformed normalized relative fluorescence versus time; lag is the x intercept; ?maxis the slope (retransformed into
linear space). The linear portion of the data was selected for analysis based on visual inspection.
hResults from curve fitting to four-parameter logistic curve Y ? Y0? [a/1 ? (x/x0)b]. Lag estimate equals x0; ?maxestimate equals 1/b; a is a fitting parameter.
TABLE 2. Results of linear regression analysis of surfactant decay
data from pulse addition studiesa
SurfactantTreatmentSlope (ppm h?1)
aAnalysis was performed on nonzero surfactant concentration (log trans-
formed) versus time after addition. Treatment refers to the previous exposure of
2996GARLAND ET AL.APPL. ENVIRON. MICROBIOL.
cells ml?1) either with or without (NH4)2SO4to evaluate N
effects on substrate response.
The fluorescence response to all of the substrates with the
addition of nitrogen followed the same logarithmic increase to
a peak succeeded by a rapid decrease to baseline levels, as
observed in the previous rhizosphere testing at the 50-ppm
concentrations of surfactants. The time to peak ranged from 8
to 18 h for all of the substrates tested, except phenylalanine,
which showed a lag in maximum response of 45 h, and peak
response ranged from 1.4 to 8.2 NRFU (Table 4). The removal
of nitrogen from the test media eliminated the response to all
the non-N-containing substrates but caused no consistent
change in response to the amino acids, as would be predicted
under nitrogen-limiting conditions.
These initial data indicate that the BD microplates offer an
approach for CLPP with lower potential for selective enrich-
ment than presently employed technology possesses. Detect-
able responses were observed at substrate addition levels 10 to
100 times less than those found in CLPP approaches based on
either CO2monitoring (1) or redox dyes (6). The time-to-peak
response was consistently less than 20 h and as short as 5 to 6 h,
indicating that a standard incubation time of 24 h could be
employed. This incubation time is shorter than the 2- to 3-day
assay period typically used for CLPP based on Biolog plates
but is longer than the several-hour incubation used in the
CO2-monitoring approach. The BD plates allow for complete
definition and concomitant manipulation of chemical factors
such as N levels that may affect substrate utilization. Further
testing with bottom-reading fluorometry may allow for testing
of intact communities (e.g., biofilm coupons and small litter
bags) compared to suspensions of organisms as in the present
study. While such extensions of the assay will not allow for
estimation of in situ rates of substrate utilization feasible with
radiotracer approaches, they will provide a CLPP approach
with increasing physiological relevance. Depending on the re-
search question, microbial ecologists can then choose CLPP
approaches targeted at phenotypic potential and/or phenotypic
Funding for this project was provided by the National Aeronautics
and Space Administration Office of Physical and Biological Research
as part of its Advanced Life Support project.
1. Degens, B. P., L. A Schipper, G. P. Sparling, and L. C. Duncan. 2001. Is the
microbial community in a soil with reduced catabolic diversity less resistant
to stress or disturbance? Soil Biol. Biochem. 33:1143–1153.
2. Di Giovanni, G. D., L. S. Watrud, R. J. Seidler, and F. Widmer. 1999.
Fingerprinting of mixed bacterial strains and BIOLOG gram-negative (GN)
substrate communities by enterobacterial repetitive intergenic consensus
sequence-PCR (ERIC-PCR). Curr. Microbiol. 38:217–223.
3. Garland, J. L., and R. M. Lehman. 1999. Dilution/extinction of community
phenotypic characters to estimate relative structural diversity in mixed com-
munities. FEMS Microbiol. Ecol. 30:333–343.
4. Garland, J. L., L. H. Levine, N. C. Yorio, J. L. Adams, and K. L. Cook. 2000.
Graywater processing in recirculating hydroponic systems: phytotoxicity, sur-
factant degradation, and bacterial dynamics. Wat. Res. 12:3075–3086.
5. Garland, J. L. K. L. Cook, C. A. Loader, and B. A. Hungate. 1997. The
influence of microbial community structure and function and community-
level physiological profiles, p. 171–183. In H. Insam and A. Rangger (ed.),
Microbial communities: functional versus structural approaches. Springer-
Verlag KG Berlin, Berlin, Germany.
6. Garland, J. L., and A. L. Mills. 1991. Classification and characterization of
heterotrophic microbial communities on the basis of patterns of community-
level sole-carbon-source utilization. Appl. Environ. Microbiol. 57:2351–2359.
7. Hobbie, J. E., R. J. Daley, and S. Jasper. 1977. Use of Nuclepore filters for
counting bacteria by fluorescence microscopy. Appl. Environ. Microbiol.
8. Konopka, A., L. Oliver, and R. F. Turco. 1998. The use of carbon substrate
utilization patterns in environmental and ecological microbiology. Microb.
9. Lehman, R. M., F. S. Colwell, D. B. Ringelberg, and D. C. White. 1995.
Combined microbial community-level analyses for quality assurance of ter-
restrial subsurface cores. J. Microbiol. Methods 22:263–281.
10. Levine, L. H., J. L. Garland, and J. V. Johnson. 2002. HPLC/ESI-quadrupole
ion trap mass spectrometry for characterization and direct quantification of
amphoteric and nonionic surfactants in aqueous samples. Anal. Chem. 74:
TABLE 3. Response of rhizosphere community samples from
hydroponic systems either unacclimated (no surfactant added) or
acclimated to SLES or CAPB in Biolog NT plates
Community SurfactantSubstrate concn (ppm)A590a
aA590with subtraction of blank, after 72 h of incubation. Data are presented
as mean and standard deviation of three replicates.
bSLES was added to wells at the designated concentration.
cCAPB was added to wells at the designated concentration.
TABLE 4. Fluorescent response from marsh litter samplesa
Type of carbon
(concn in mM)
With NitrogenWithout Nitrogen
Carbohydrate Fructose (1)3.64
Amino acidGlutamic acid (1)
Organic acidGlycolic acid (1)
aPeak values are reported in NRFU. Two replicates were exposed to each
carbon source both with and without the addition of 2.5 mg of [N]NH4per liter
in the media.
VOL. 69, 2003 CLPP PERFORMED WITH AN OXYGEN-SENSITIVE FLUOROPHORE2997
11. Levine, L. H., J. E. Judkins, and J. L. Garland. 2000. Determination of Download full-text
anionic surfactants during wastewater recycling process by ion pair chroma-
tography with suppressed conductivity detection. J. Chromatogr. 874:207–
12. Mills, A. L., and J. L. Garland. 2002. Application of physiological profiles to
assess community properties, p. 135–146. In C. J. Hurst, R. L. Crawford,
G. R. Knudsen, M. J. McInerney, and L. D. Stetzenbach (ed.), Manual of
environmental microbiology. ASM Press, Washington, D.C.
13. Reid, B. J., C. J. A. MacLeod, P. H. Lee, A. W. J. Morriss, J. D. Stokes, and
K. T. Semple. 2001. A simple14C-respirometric method for assessing micro-
bial catabolic potential and contaminant bioavailability. FEMS Microbiol.
14. Smalla, K., U. Wachtendorf, H. Heuer, W.-T. Liu, and L. Forney. 1998.
Analysis of BIOLOG GN substrate utilization patterns by microbial com-
munities. Appl. Environ. Microbiol. 64:1220–1225.
15. Swisher, R. D. 1987. Surfactant biodegradation, 2nd ed. Marcel Dekker, New
16. Winding, A., and N. B. Hendriksen. 1997. Biolog substrate utilization assay
for metabolic fingerprints of soil bacteria: incubation effects, p. 195–205. In
H. Insam and A. Rangger (ed.), Microbial communities: functional versus
structural approaches. Springer-Verlag KG Berlin, Berlin, Germany.
17. Wodnicka, M., R. D. Guarino, J. J. Hemperly, M. R. Thomas, D. Stitt, and
J. B. Pitner. 2000. Novel fluorescent technology platform for high through-
put cytotoxicity and proliferation assays. J. Biomol. Screen. 5:141–152.
2998GARLAND ET AL.APPL. ENVIRON. MICROBIOL.