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Applied and Environmental Microbiology, October 2001, p. 4662-4670, Vol. 67, No. 10
0099-2240/01/$04.00+0 DOI: 10.1128/AEM.67.10.4662-4670.2001
Copyright © 2001, American Society for Microbiology. All rights reserved.
Resolution of Viable and Membrane-Compromised
Bacteria in Freshwater and Marine Waters Based on Analytical Flow
Cytometry and Nucleic Acid Double Staining
Gérald
Grégori,1,*
Sandra
Citterio,2
Alessandra
Ghiani,2
Massimo
Labra,2
Sergio
Sgorbati,2
Spencer
Brown,3 and
Michel
Denis1
Laboratoire d'Océanographie et de
Biogéochimie, Université de la Méditerranée,
CNRS UMR 6535, 13288 Marseille,1 and
Institut des Sciences du Végétal, CNRS UPR 2355,
91198 Gif-sur-Yvette,3 France, and
Dipartimento di Scienze dell'Ambiente e del Territorio,
Università di Milano-Bicocca, 20126 Milan,
Italy2
Received 20 March 2001/Accepted 29 June 2001
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ABSTRACT |
The membrane integrity of a cell is a well-accepted criterion for
characterizing viable (active or inactive) cells and distinguishing them from damaged and membrane-compromised cells. This information is
of major importance in studies of the function of microbial assemblages
in natural environments, in order to assign bulk activities measured by
various methods to the very active cells that are effectively
responsible for the observations. To achieve this task for bacteria in
freshwater and marine waters, we propose a nucleic acid
double-staining assay based on analytical flow cytometry, which
allows us to distinguish viable from damaged and membrane-compromised
bacteria and to sort out noise and detritus. This method is derived
from the work of S. Barbesti et al. (Cytometry 40:214-218, 2000) which
was conducted on cultured bacteria. The principle of this approach is
to use simultaneously a permeant (SYBR Green; Molecular Probes) and an
impermeant (propidium iodide) probe and to take advantage of the energy
transfer which occurs between them when both probes are staining
nucleic acids. A full quenching of the permeant probe fluorescence by
the impermeant probe will point to cells with a compromised membrane, a
partial quenching will indicate cells with a slightly damaged membrane, and a lack of quenching will characterize intact membrane cells identified as viable. In the present study, this approach has been
adapted to bacteria in freshwater and marine waters of the Mediterranean region. It is fast and easy to use and shows that a large
fraction of bacteria with low DNA content can be composed of viable
cells. Admittedly, limitations stem from the unknown behavior of
unidentified species present in natural environments which may depart
from the established permeability properties with respect to the
fluorescing dyes.
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INTRODUCTION |
Bacteria are a key component in
aquatic and terrestrial ecosystems due to their wide biodiversity,
their capacity to survive extreme environments, and their large variety
of metabolic activities. In aquatic environments, with elevated
abundances in the range of 105 to
106 cells cm
3
(39), bacteria play a major role in recycling organic
matter and subsequently in sustaining the nutrient turnover. The
understanding of bacterial activity with respect to the fate of organic
matter in marine water and freshwater is ecologically important, and many studies are dedicated to this objective (7, 9, 10, 37, 38,
40, 51). Bacteria are able to metabolize part of the dissolved
organic matter (DOM) present in micro- or nanomolar concentrations in
marine environments (5, 25) and to convert it into
particulate organic matter through bacterial biomass production, making
it available to higher trophic levels (38). The rate at
which the biological oxidation of DOM by bacteria occurs is dependent
on DOM biodegradability (2, 18). Conversely, viral lysis
of bacteria releases cellular constituents into the environment and
supplies a new source of carbon and nitrogen for primary producers and
consumers (1, 40).
The presence of bacteria in aquatic environments is also of special
concern with respect to human health problems and water quality.
Indeed, several bacterial strains in freshwater and marine water have
been demonstrated to be pathogenic. For instance, Vibrio vulnificus in estuarine and marine waters is responsible for fatal infections following ingestion (typically of wild oysters) or contamination of a wound (28, 36, 52). Other species, such as Escherichia coli and Salmonella enterica
serovar Enteritidis have been shown to be indicators of human
pollution through fecal contamination (14). In response to
their environmental conditions, bacteria may be present in a viable but
nonculturable state by classical microbiological methods (6,
26, 32). When bacteria are monitored, it is very important to
determine the viability of cells in addition to their abundance, since
this is a critical issue in assessing water quality and preventing
sanitation problems (44). New approaches have been
developed to assess bacterial viability (29, 33, 39). Most
are based on the ability to detect metabolic or functional activity by
using different fluorochromes: (i) detection of cell divisions by
counts of CFU on plates (34), (ii) assessment of membrane
potential with dyes of the carbocyanine and oxonol families
(31), (iii) dye (propidium iodide [PI] or ethidium
bromide) exclusion due to membrane integrity (19, 43), (iv) intracellular reduction of 5-cyano-2,3-ditolyl tetrazolium chloride (CTC) and
2-(p-iodophenyl)-3-(p-nitrophenyl)-5-phenyl tetrazolium chloride (INT) by dehydrogenases (8, 11, 41, 46, 47,
49, 50), (v) reduction of fluorescein diacetate (FDA) by
esterases (17, 22), (vi) radiolabeled substrate uptake analyzed by microautoradiography (24, 47), (vii) the
presence of nucleoids (55), and (viii) nucleic acid
content heterogeneity (12). Molecular analysis of
components such as ribosomes can also reveal the state of growth
(13). The 16S rRNA probe count (24, 42, 48,
53) is one example of molecular approaches.
In investigations of bacteria in natural environments, it is necessary
to distinguish them from other particles and to characterize them by
their properties, such as morphology, physiology, and taxonomy. All of
these aspects imply analyses at the individual cell level, but methods
involving cell enumeration by optical microscopy are both
time-consuming and subjective (39). The development of
analytical flow cytometry (45), together with advances in
the field of fluorescent probes (21, 39), have enabled the
application of this rapid and semiautomatic technique to the study of
bacteria in natural aquatic environments. Barbesti et al.
(3) took advantage of flow cytometry to devise a DNA double-staining protocol based on energy transfer between specific probes to distinguish viable, dead, and compromised cells in
bacterial cultures. In the present study, we report the use of this
nucleic acid double-staining (NADS) protocol with natural freshwater
and marine water to count bacteria and to rapidly determine clusters according to their cell membrane permeability by using the same assay.
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MATERIALS AND METHODS |
Sampling sites.
Freshwater samples were collected from three
rivers in northern Italy: the Adda, the Lambro, and the Seveso. A
monitoring experiment was conducted along the Adda River, and samples
were collected ca. 1 km upstream from a sewage outflow, at the site of
sewage outflow, and at ca. 1 and 6 km downstream from the sewage outflow. Sampling was carried out once a week for 1 month.
Seawater samples were collected in surface waters from two different
environments: at site 1 inside Pointe-Rouge, a pleasure-boat harbor in
Marseilles, France, and site 2 facing Maire Island, at the southern
edge of the city and not under its direct influence. Samples were
prefiltered on a 100-µm-mesh-size net to avoid clogging of the instrument.
Instruments.
Different flow cytometers were used in
this study as follows. (i) We used a Bryte-HS (Bio-Rad,
Hercules, Calif.) flow cytometer equipped with a 75-W Xenon lamp, and a
470- to 490-nm excitation filter was used for the analysis of
freshwater samples. The emitted fluorescence was split into two
different channels: FL1 (green) at 515 to 565 nm and FL2 (orange-red)
at 590 to 650 nm. All of the data related to freshwater samples were
analyzed by using Bio-Rad WinBryte software. (ii) Wed also used a
Cytoron Absolute (Ortho Diagnostic Systems) flow cytometer, equipped
with an air-cooled argon laser (excitation wavelength at 488 nm) to
analyze seawater samples. Each cell was characterized by five optical
parameters: two diffraction parameters, namely, forward-angle scatter
(related to the particle size) and right-angle scatter (related to the cell structure), and three fluorescence parameters measuring emissions in the red (>620-nm), orange (565- to 592-nm), and green (515- to
530-nm) wavelength ranges. Data were collected and stored in list mode
with Immunocount software (Ortho Diagnostic Systems). This software
provides directly the cell concentration (as cells millimeter3) of the resolved populations. (iii)
Finally, we used an Elite EPS (Beckman-Coulter) equipped with a 488-nm
argon laser for cell sorting in Gif-sur-Yvette, France. The emitted
fluorescence was split into three different channels: PMT3 (green) at
520 to 530 nm, PMT4 (orange) at 615 to 640 nm, and PMT5 (red) at 645 to
795 nm. Sorting was done with a sample rate of ca. 1,000 events
s1 into Eppendorf tubes. Visual checks for the
identification of bacteria from detritus, as well as sorting and
staining, were made on a Polyvar (Reichert, Vienna, Austria) microscope
with epifluorescence and interference contrast.
Fluorescent probes.
SYBR Green I or II (Molecular Probes,
Eugene, Oreg.) and PI (Sigma Chemical Co.) were used for the double
staining of nucleic acids. SYBR Green and PI fluoresce in the green
(maximum at 521 nm) and the red (maximum at 617 nm) wavelength ranges,
respectively, when excited with a 488-nm argon laser. They both stain
RNA and DNA (15). 5 (and 6)-carboxyfluorescein diacetate
(CFDA; Molecular Probes) was employed to detect esterase activity in
living cells (20, 22). A stock solution of 5 mg
cm3 was made in dimethyl sulfoxide (Sigma) and
stored at 4°C. This stock solution was thawed and diluted 100-fold in
distilled water to prepare daily working solutions kept for a maximum
of 3 h to avoid CFDA degradation. Samples (3 cm3) were supplemented with
100-mm3 CFDA working solution and were analyzed
after 5 min of incubation in the dark at room temperature.
Flow cytometric analysis.
For each freshwater sample, three
dilutions were prepared and stained with 1:10,000 (vol/vol) SYBR Green
I (neither the molecular weight nor the chemical formula are provided
by the manufacturer) and a 10-µg cm3 PI
commercial solution. The final fluorochrome concentrations were as
defined by Barbesti et al. (3) for bacterial cultures. Seawater samples were stained without dilution with 1:1,000 (vol/vol) SYBR Green II (neither the molecular weight nor the chemical formula are provided by the manufacturer) and a 10-µg
cm3 PI commercial solution. For both types of
samples, flow cytometric analyses were processed after 30 min of
incubation in the dark.
CFU.
The culturability of bacteria from freshwater was
assessed by mixing 0.1 cm3 of the sample with
ca. 25-cm3 of plate count agar medium [PCA;
peptone from 5 g of casein dm3, 2.5 g
of yeast extract, 1 g of D-(+)-glucose
dm3, 14 g of agar
dm3 (pH 7.0 at 25°C)] at 45°C.
After medium solidification, plates were incubated for 72 h at
22°C. The total bacterial charge was estimated as the CFU per cubic
centimeter of sample.
Ozone and heat treatments.
One liter of freshwater was
treated with ozone (ca. 30 mg dm3) at a
laboratory scale for 90 min in order to kill the sample cells. After
ozone treatment, the bacterial viability and culturability was checked
by analyzing 1 cm3 of ozonized sample by flow
cytometry and plating 0.1 cm3 on PCA, respectively.
The flow cytometric characterization of membrane-compromised bacteria
was also done by exposing cells of seawater samples
to heat treatment
(40 min in a 60 to 70°C water
bath).
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RESULTS |
NADS protocol.
The live-dead protocol developed by
Barbesti et al. (3) for cultures of E. coli
is based on the simultaneous staining of bacteria nucleic acids by a
permeant (SYBR Green I) and an impermeant (PI) fluorescent probe and
the interpretation of the green versus the red fluorescence cytograms.
Green, green plus orange-red, and orange-red E. coli cells
were identified as live, damaged, and dead cells, respectively
(3). In order to extend this approach to natural samples
from aquatic environments, several experiments were conducted with
bacteria from freshwater and seawater.
Figure
1 shows the green versus the red
fluorescence cytograms from marine samples under different conditions.
Cytogram A1
is representative of marine samples in the absence of
staining.
The weak signals close to the origin are due to the
instrument's
electronic noise and the background signals of
nonfluorescent
heterotrophic bacteria. Stained cells will therefore
yield fluorescence
signals above this background. Cytograms B1 and B2
were obtained
with samples of 0.2-µm-pore-size-filtered seawater
supplemented
with PI (cytogram B1) or SYBR Green II (cytogram B2). With
these
cell-free samples, signals are produced by the instrument noise
and the free dyes in solution which may generate a large number
of
events and quickly saturate the data acquisition. Consequently,
a
fluorescence threshold (dotted square) was systematically introduced.
Cytograms C1 and C2 correspond to fresh marine samples stained
with PI
and SYBR Green II, respectively. Few cells appeared to
be stained by PI
(red fluorescence signals) in cytogram C1, and
two populations
were distinguished upon staining by SYBR Green
II (green fluorescence
signals) in cytogram C2.
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FIG. 1.
Green versus red fluorescence cytograms corresponding to
different test experiments. Panels: A1, unstained seawater sample; B1
and B2: 0.2-µm-filtered seawater sample supplemented with PI and SYBR
Green II, respectively; C1 and C2, seawater samples supplemented with
PI and SYBR Green, respectively. The final probe concentrations were 10 µg of PI cm3 and 1:1,000 (vol/vol) SYBR Green II.
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In Fig.
2, cytograms D1 and D2 concern a
seawater sample subjected to the heat treatment described in Materials
and Methods
to kill all cells, including bacteria. Membrane-compromised
bacteria
are identified by their PI staining in cytogram D1, whereas
staining
with SYBR Green II (cytogram D2) resulted in a signature
similar
to that of the fresh sample in cytogram C2. When both stains
were
simultaneously incubated in the heat-treated sample, cells
essentially
exhibited red fluorescence as expected (cytogram D3).
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FIG. 2.
Single and dual staining of heat-treated seawater
samples. Panels: D1 and D2, heat-treated seawater samples supplemented
with PI and SYBR Green II, respectively; D3, heat-treated
(40 min at 60 to 70°C) seawater sample simultaneously stained with PI
and SYBR Green. Killed bacteria essentially appeared as red fluorescent
cells. The final probe concentrations were 10 µg of PI
cm3 and 1:1,000 (vol/vol) SYBR Green II.
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In an experiment in which a seawater sample was double stained with
SYBR Green II and PI, cells fluorescing preferentially
in the green
(gated cells in the left cytogram of Fig.
3) were
sorted as described in Materials
and Methods and stained with
CFDA. This additional staining resulted in
a significant increase
in the green fluorescence of all sorted cells.
In contrast, heat-treated
cells, which only yielded red fluorescence
after double staining
by SYBR Green II and PI, did not emit green
fluorescence after
being stained with CFDA (not shown).
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FIG. 3.
Cell sorting and esterase activity assay. Green
fluorescent bacteria resolved by the NADS protocol (gated cells on the
fresh-marine-sample cytogram) were sorted and further stained with CFDA
as described in Materials and Methods. The cells exhibited an increase
in green fluorescence due to esterase activity and carboxyfluorescein
retention (gated cells on the sorted cells-plus-CFDA cytogram).
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Ozone treatment of freshwater samples led to complementary results. In
Fig.
4, a freshwater sample collected
from Seveso River
and double stained with SYBR Green I and PI
essentially gave cells
with green fluorescence (untreated-sample
cytogram), whereas all
cells fluoresced red after ozone treatment
(ozone-treated-sample
cytogram). After 0.1 cm
3 of
the fresh sample was plated on PCA, the formation of numerous
colonies
was observed (Fig.
4, left plate). In contrast, no colonies
were
observed after being plated on PCA with 0.1 cm
3
of the fraction submitted to ozone treatment (Fig.
4, right plate).
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FIG. 4.
Application of the NADS protocol to a Seveso River
(Italy) sample before and after ozone treatment. The untreated-sample
cytogram corresponds to a freshwater subsample mainly containing green
bacteria. The ozone-treated-sample cytogram was obtained after ozone
treatment to kill cells, and it shows essentially red bacteria.
Quadrants are identified as A to D. The plate count obtained with green
bacteria from the untreated sample shows abundant colonies, whereas the
plate count made with red cells from the ozone-treated sample yielded
no colonies.
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Freshwater. (i) Protocol optimization.
Bacterial abundance in
freshwaters varies significantly, depending upon the level of
eutrophication and various discharges produced by human activity. We
have therefore tested the adequacy of NADS, devised for bacterial
cultures (1:10,000 [vol/vol] SYBR Green I final dilution and a
10-µg cm3 final concentration of PI )
(3), when applied to freshwater samples. Adda River
samples were stained as collected and after a 10-fold dilution. For the
diluted subsample, the background fluorescence of SYBR Green I in
solution was too high to be eliminated in the orange-red channel by
electronic compensation (not shown). In addition, an increase in
background fluorescence, affecting signal resolution, was observed for
high PI concentration. To avoid these problems, it was necessary to
adapt the concentrations of the fluorescent probes to that of bacteria.
A final concentration of 1 µg of PI cm3 and a
final dilution of 1:100,000 for SYBR Green I were found to be optimal
for freshwater samples with less than 0.5 × 106 bacteria cm3. For
more concentrated freshwater samples (up to 20 × 106 bacteria cm3), the
fluorochrome concentrations (1:10,000 [vol/vol] SYBR Green I final
dilution and 10-µg cm3 final concentration of
PI) determined for bacterial cultures remain the best. The
staining efficiency was also checked with respect to incubation
time for 60 min. Staining was stable for each class after 30 min of incubation.
(ii) Adda River monitoring.
The Adda River (Italy) was sampled
as described in Materials and Methods. The different bacterial states
were resolved using the NADS protocol, and a typical distribution is
displayed in Fig. 5. At 1 km upstream
from the sewage outflow, the green and green-plus-red bacterial
fractions corresponded to 61% of the whole bacterial population,
whereas the red fluorescent bacteria amounted to 39% (Fig. 5). The
strong bacterial input at the site of the sewage outflow (5 × 107 cells cm3) was
reduced by ca. 80% 1 km downstream and by ca. 90% 5 km farther downstream. The sewage outflow also corresponded to an inversion of the
ratio of green and green-plus-red cells to red cells, leading to 60%
red fluorescent cells. This effect was reduced by half 1 km
downstream and fully disappeared 6 km downstream, where the percentage of red fluorescent cells decreased to 35% as upstream.
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FIG. 5.
Bacteria distribution in the Adda River (Italy) at four
different stations: 1 km upstream from a sewage outflow, at the sewage
outflow site, and at 1 and 6 km downstream from the sewage outflow
point. Bacteria were resolved according to their fluorescence (green
and green-plus-red bacteria and red fluorescent bacteria) generated by
the NADS protocol. The values in brackets are the percentages of red
fluorescent bacteria relative to the total count.
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Marine water. (i) Protocol optimization.
In the SYBR Green dye
family, SYBR Green II expresses a higher selectivity for RNA, with a
quantum yield of 0.54, while keeping a strong affinity for
double-stranded DNA with a quantum yield of 0.36, about half that of
SYBR Green I (15). SYBR Green II was therefore tested on
marine samples to better use the energy transfer phenomenon when both
dyes (SYBR Green II and PI) bind to nucleic acids. The optimum
concentration of SYBR Green II to detect bacteria in seawater was found
to be a 1:1,000 (vol/vol) SYBR Green II final dilution after we tested
the concentration range from 0.025:1,000 to 2:1,000 (vol/vol).
This threshold for a maximum detection of bacteria is effective after
only 10 min of incubation. The concentration of PI was set at 10 µg
cm3. The appropriate incubation time for the
NADS procedure when SYBR Green II and PI were used was also found to be
30 min by monitoring the time course of the different bacterial states
resolved by NADS during 1 h of incubation.
The flow cytometric analysis of bacteria in seawater by the NADS
protocol yielded typical cytograms characterizing cells by
their
fluorescence in panels A, B, and D (Fig.
4) as green,
green-plus-red,
and red, respectively, as for freshwater
bacteria.
(ii) Northwestern Mediterranean coast.
The comparison of the
bacterial abundance at both marine sampling sites and in the different
fluorescent states is reported in Fig. 6.
The overall abundance was higher by 80% at the Pointe-Rouge site (site
1; 8.87 × 105 cells
cm3) than at the site facing Maire Island (site
2; 4.94 × 105 cells
cm3). The amount of red fluorescent
bacteria was similar at both sites (ca. 3 × 105 cells cm3), which led
to a lower percentage of red fluorescent bacteria at site 1 than at
site 2 (33 versus 53%). Consistently, there were more green bacteria
at site 1 than at site 2, both in absolute and in relative terms (62 and 41%, respectively). The levels of green-plus-red bacteria were
similar at both sites (5 and 6%, respectively).
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FIG. 6.
Staining classes for bacteria collected from surface
water inside Pointe-Rouge harbor (Marseilles, France) and a few
kilometers away from the town, at the site facing Maire Island.
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The relationship between the DNA content and the bacterial response to
the NADS assay is illustrated in Fig.
7
with different
staining protocols applied to a seawater sample
collected at site
2. As previously reported (
12), two
different populations are
distinguished by their DNA content (Fig.
7) when bacteria were
stained with SYBR Green II alone: a population
with a low DNA
content (Fig.
7, LDNA) characterized by a low green
fluorescence
and a population with a high DNA content (HDNA)
characterized
by a higher green fluorescence (Fig.
7). The NADS
protocol applied
to a subsample resolved two populations within the
green fluorescent
cells (Fig.
7). The bacterial abundances yielded by
the two staining
protocols are summarized in Table
1. Both protocols gave the
same amount of
total bacteria (5.4 × 10
5 to 5.5 × 10
5 cells cm
3). In
addition, all HDNA bacteria characterized upon staining
with SYBR Green
II alone remained as green bacteria after the
NADS assay, whereas
LDNA bacteria were resolved as green (~68%),
green-plus-red (2%),
and red cells (~30%).
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FIG. 7.
Comparison of single (SYBR Green II; 1:1,000 [vol/vol]
final dilution) and double (NADS protocol) staining of bacteria in a
seawater sample collected off Marseilles at the site facing Maire
Island. The green versus the red fluorescence cytograms corresponding
to SYBR Green II staining reveal the presence of bacteria with high (H)
and low (L) DNA contents. The cytogram associated with the NADS
protocol shows a significant amount of LDNA bacteria appearing as
red-negative cells in quadrant A. At the same time, cells that were
negative with SYBR Green II were apparent with PI staining in quadrant
D. Refer to Table 1 for the counts.
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TABLE 1.
Comparison of single (SYBR Green II) and double (NADS
protocol) staining of bacteria in a seawater sample collected off
Marseilles at the site facing Maire Island
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DISCUSSION |
This NADS flow cytometric protocol is based on the simultaneous
use of permeant (SYBR Green dyes) (27) and impermeant (PI) fluorescent probes (22, 30, 43, 53). Both dyes can be readily excited with the blue light from a laser or arc lamp of relatively simple and portable flow cytometers, which makes the assay
suitable for field control. The efficiency of the combined staining is
magnified by the energy transfer from SYBR Green to PI when both are
bound to the nucleic acids as described by Barbesti et al.
(3). Consequently, membrane-compromised bacterial cells are expected to fluoresce in the red-wavelength range, whereas the
fluorescence emitted by membrane intact cells will be restricted to the
green wavelengths. Cells with a partially damaged membrane will enable
various amounts of PI to bind some nucleic acids that will result in a
corresponding increase of the red fluorescence and a decrease of the
green fluorescence depending on the extent of the energy transfer from
SYBR Green to PI. SYBR Green I performed adequately with freshwater,
but SYBR Green II has been shown to perform better in seawater
(27).
Analogous fluorescence quenching could be achieved by using Hoechst
stain and PI to test the membrane integrity (35) in a
double-staining protocol. However, Hoechst fluorochromes require UV
excitation, which generally necessitates the use of expensive flow cytometers.
Evidence for esterase activity and fluorochrome retention in all green
fluorescent cells sorted after the NADS assay (Fig. 3), as well as the
plate count experiment (Fig. 4), lend independent support to the
identification of the green cells as viable. In addition, the heat
(Fig. 2) and ozone (Fig. 3) treatments used to artificially kill cells
in the analyzed samples also provided independent support for the
identification of red cells as membrane-compromised cells, qualifying
our NADS protocol as a membrane integrity assay for bacteria in
freshwater and marine water. However, if dead (heat- or ozone-killed)
cells undoubtedly appear as red cells, the full identification of red
cells in natural samples as dead cells remains to be further
substantiated. Indeed, some unknown species might be able to accumulate
PI without the prerequisite of having a compromised membrane. The NADS
assay descibed here requires an incubation time of 30 min to achieve a
full staining, both for freshwater and seawater samples, which is
consistent with other published staining protocols (27),
wherein reported incubations last 15 and 30 min for fixed and unfixed
bacteria, respectively. In the present protocol, only bacteria
containing nucleic acids are stained either by both PI and SYBR Green I
or II in the case of membrane-compromised cells or by SYBR Green I or
II alone when the insertion of PI is prevented by the membrane integrity. Under these conditions, bacterium-like detritus without nucleic acids cannot be confused with real bacteria (55).
The assay can be processed in <1 h, and it yields both quantitative (i.e., cell count) and qualitative (i.e., for viable,
membrane-damaged, and membrane-compromised cells) information for each
analyzed sample. The assessment of viability by epifluorescence
microscopy after acridine orange or DAPI
(4',6'-diamidino-2-phenylindole) staining on membrane filters
(16, 54)
a more time-consuming, less accurate, and less
objective approach than flow cytometric assayis incompatible with
routine sampling and analyses.
The NADS protocol offers higher resolution than the HDNA-LDNA
classification resulting from staining DNA with a single fluorescent probe (12), in which cells with a high DNA content were
identified as the fraction containing active bacteria and those with a
low DNA content were representative of inactive and dead bacteria. Damaged cells were not resolved in this approach, whereas the NADS
protocol clearly separates inactive cells (i.e., membrane compromised,
in red) from damaged cells (in red-plus-green).
The above-described results related to the NADS protocol support the
hypothesis that HDNA bacteria are viable cells. However, the
assimilation of LDNA bacteria with dead cells should be revised, since
in the above example two-thirds of the LDNA bacteria were found to be
viable and only one-third compromised membrane cells were found to be
viable (Table 1). The remaining 2% were membrane-damaged cells, a
transient step between viable and membrane-compromised states. We
recall that the NADS protocol is based only on the cell membrane
integrity, which is a widely accepted criterion for viability
(23, 35). Indeed, in spite of the long debate about live
and dead bacteria (4), the loss of membrane integrity results in the collapse of the cell energetics and active transports, that is to say, the death of the cell (35). To the extent
that the red fluorescing (PI permeable) cells have irreversibly damaged membranes, this parameter can provide a rough approximation of nonviable cells. On the other hand, a cell with preserved
membrane integrity (PI impermeable and therefore only
stained by green fluorescent SYBR Green) can be considered as viable,
which covers active (enzyme activities, metabolism, compact DNA, and
high content in rRNA) or inactive states (e.g., the lack of enzyme
activity in nutrient-depleted environments) (35). In
addition, active cells include culturable and nonculturable
bacteria. The reported NADS assay is not, strictly speaking, a
live-dead assay but rather a proxy, whose efficiency depends on cell
types and physiologic states occurring in natural samples composed of
very diverse populations in physiologic and cytostructural terms.
Indeed, present limitations to the NADS assay stem from the large
biodiversity of bacteria in natural environments, where most of the
species are still unknown. Therefore, it is possible that some
as-yet-unknown species may not be consistent with observations made
thus far regarding the bacterial membrane permeability with respect to
the fluorescent probes used in the NADS assay (PI and SYBR
Green). This must be kept in mind when one is dealing with
natural samples. The NADS assay, which only addresses membrane
integrity, provides the resolution of a bacterial population into
viable (active, inactive, culturable, and nonculturable),
membrane-damaged, and membrane-compromised cells, but complementary
approaches are needed to independently demonstrate the presence of
specific activity or metabolism, as well as the capacity for a cell to
undergo division. Because of its simplicity, NADS should become an
efficient tool in field sample analysis and a valuable source of
information concerning the functioning of the microbial compartment.
Indeed, until now, bacterial activity and production have referred to
total bacteria counts, wherein dead cells may represent a large
fraction (12; this study), even though these cells
contribute neither to remineralization nor to bacterial biomass
production. The application of the flow cytometric NADS protocol in
field studies will bring quantitative and qualitative improvements to
the basic information characterizing the role of bacteria in the
environment. This should in turn clarify our understanding of the part
played by bacteria in biogeochemical cycles and their modeling.
| |
ACKNOWLEDGMENTS |
This study was supported by the Integrated Action Programme
"GALILEE" between France and Italy. G.G. is the recipient of a doctoral fellowship from the Council of the
Provence-Alpes-Côte-d'Azur Region.
| |
FOOTNOTES |
*
Corresponding author. Mailing address: Laboratoire
d'Océanographie et de Biogéochimie,
Université de la Méditerranée, CNRS UMR 6535, 163 avenue de Luminy, Case 901, 13288 Marseille Cedex 9, France.
Phone: 33-4-91-82-91-14. Fax: 33-4-91-82-65-48. E-mail:
gregori{at}com.univ-mrs.fr.
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Applied and Environmental Microbiology, October 2001, p. 4662-4670, Vol. 67, No. 10
0099-2240/01/$04.00+0 DOI: 10.1128/AEM.67.10.4662-4670.2001
Copyright © 2001, American Society for Microbiology. All rights reserved.
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Abstract
Introduction
