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Applied and Environmental Microbiology, March 2001, p. 1384-1387, Vol. 67, No. 3
0099-2240/01/$04.00+0 DOI: 10.1128/AEM.67.3.1384-1387.2001
Copyright © 2001, American Society for Microbiology. All rights reserved.
Determination of Virus Abundance in Marine
Sediments
R.
Danovaro,1,2,*
A.
Dell'Anno,1
A.
Trucco,1
M.
Serresi,3 and
S.
Vanucci4
Institute of Marine Science, University of
Ancona, 60131 Ancona,1 Department of
Zoology, University of Bari, 70125 Bari,2
Faculty of Medicine and Surgery, University of Ancona, 60020 Ancona,3 and Department of Animal
Biology and Marine Ecology, University of Messina, Salita Sperone,
90100 Messina,4 Italy
Received 28 August 2000/Accepted 13 December 2000
 |
ABSTRACT |
In this study, we optimized procedures to enumerate viruses from
marine sediments by epifluorescence microscopy using SYBR Green I as a
stain. The highest virus yields from the bulk of the sediments were
obtained by utilizing pyrophosphate and 3 min of sonication. The
efficiency of extraction benthic viruses by pyrophosphate-ultrasound
treatment was about 60% of the extractable virus particles. Samples
treated with nucleases had increased virus counts, suggesting a masking
effect of extracellular DNA. No significant differences were observed
between virus counts obtained by epifluorescence microscopy and
transmission electron microscopy. Both formaldehyde and glutaraldehyde
gave significant reductions of virus counts after only 24 h of sediment
storage, but no further loss occurred after 7 days.
| |
TEXT |
Viruses are now considered to be an
important component of all aquatic microbial communities. The
reevaluation of the role of viruses in marine ecosystems is due to the
discovery of very high virus abundance (see reference 13
for a review). Recent studies have stressed the ecological implication
of viruses in the release of dissolved organic matter, nutrient
recycling (18), and the pathways of organic carbon
utilization, with cascade effects on marine microbial food webs and
organic-matter cycling (12). The available methods for the
determination of virus abundance in aquatic environments include
counting by transmission electron microscopy (TEM) (1, 2, 16,
21), by flow cytometry (17), and by epifluorescence
microscopy (EFM) (9, 14, 20, 26, 29). The last technique
allows an accurate and easily performed enumeration, avoiding the use
of expensive and bulky equipment (13). In addition, EFM is
reported to be up to seven times more efficient than TEM for counting
viruses (15, 28).
Available information dealing with benthic virus ecology is scant. This
is due to the lack of adequate protocols for easily determining their
abundance and distribution in marine sediments (6). The
main objective of this work was the optimization of procedures to
enumerate viruses in different marine sediments. We focused our
attention on EFM counting using SYBR Green I as a stain
(20) in order to address the following issues: (i) virus dislodgment from sediment particles (using surfactant and ultrasound treatments), (ii) the efficiency of virus extraction from bulk sediment
(by the number of postsonication washings), and (iii) stain-counting
accuracy and efficiency (by removing possible interferences due to
extracellular DNA in virus counting and by comparison with TEM counts).
In addition, we tested the effects of preservatives on virus abundance
in long-time-course experiments carried out on fixed sediment samples.
Sediment sampling and selection of sediment.
In order to make
the protocol for virus counting suitable for the widest variety of
sediment samples, two different sediment types were selected in this
study: shallow sands and deep-sea muds. As deep-sea samples are not
generally analyzed immediately, the effects of long-term storage with
preservatives were also investigated. Sandy-sediment samples (modal
grain size, between 125 and 250 µm) were collected by hand coring
(using Plexiglas tubes [4.7 cm inside diameter]) in June and in
September 1999 near the low tidal line of a quiet tidal flat in
Falconara Beach (43°6'N, 13°5'E; northern Adriatic Sea). Deep-sea
sediment samples were collected in March 1998 in the Porcupine Abyssal
Plain (northeastern Atlantic Ocean at 4,800-m depth; 48°50'N,
16°29'W). This area, characterized by strong seasonality and a high
interannual variability in organic-matter inputs (8, 19),
can be considered representative of typical deep-sea conditions
(23). Undisturbed sediment samples were collected with a
multicorer (Maxicorer; inside diameter, 9.0 cm; depth penetration, >20
cm). For virus analysis, immediately after sampling, subsamples of
about 0.5 ml of the top 5 mm of both sediment types were taken from
different cores and deployments, added to 3 ml of prefiltered
(0.02-µm pore size) seawater containing 2% formalin, and stored at
4°C. All virus analyses of fixed material were performed within 3 to
4 weeks after fixation. Long time course experiments to study the
effects of formaldehyde and glutaraldehyde storage were carried out on
sandy sediments. The glassware utilized for virus counts was carefully
cleaned by soaking it in 10% HCl overnight, rinsed with MilliQ water
and subsequently autoclaved. All the solutions were prepared with
MilliQ water, filtered through 0.02-µm-pore-size filters, and then autoclaved.
Treatments for virus dislodgment.
Sediment samples collected
in the northeastern Atlantic (triplicate 0.5-ml samples) were added to
4.0 ml of MilliQ water and 1.0 ml of sodium pyrophosphate solution
(10-mM final concentration) and incubated for 15 min. Additional
sediment samples (n = 3; 0.5 ml) without pyrophosphate
were added to 5 ml of MilliQ water and served as controls. After
incubation, all samples were shaken manually for 1 min and then
centrifuged (800 × g; 1 min) to reduce interference
due to suspended particles. Aliquots of the supernatant were diluted
500 to 1,000 times and filtered through 0.02-µm-pore-size Anodisc 25 membrane filters (pressure, <100 mm of Hg). The filters were then
stained with 20 µl of SYBR Green I (Lot no. 4967-30; diluted 20-fold
in MilliQ water; optical density at 495 nm = 1.357) for 15 min in
the dark, rinsed twice with 1 ml of MilliQ water (in order to eliminate
fluorescence background noise), and analyzed by EFM using a Zeiss
Axioplan microscope equipped with a 50-W lamp. Ten to 50 fields were
viewed at ×1,000 magnification, and a minimum of 400 viruses were
counted. Viruslike particles (VLP) were discriminated from bacteria
(0.2- to 2-µm diameter) by their dimensions (0.015- to 0.2-µm
diameter [20]). As sodium pyrophosphate enhanced virus
extraction efficiency, any further steps were carried out with this
surfactant. In order to test the combined effects of pyrophosphate and
ultrasound treatments on virus extraction, muddy and sandy sediments
(n = 3; 0.5 ml for both sediment types) were added to
pyrophosphate and sonicated (Branson 2200 sonifier; 100 W; 47 kHz) for
0, 1, 3, 8, and 15 min in an ice bath to prevent overheating. The
sonication was interrupted for 30 s every minute, during which
time the samples were shaken manually. After each treatment, the sample
was centrifuged, and aliquots of the supernatant were processed as
described above.
Postsonication extraction efficiency.
Once the optimal
sonication time was identified, the efficiency of virus detachment from
sediment particles was checked by estimating the ratio of virus
abundance after the first extraction with ultrasound and pyrophosphate
treatment versus the cumulative virus abundance obtained by this
procedure plus three further washing steps. The added steps were the
following: (i) an aliquot of supernatant obtained from deep-sea
sediment samples (0.5 ml of sediment plus 4.0 ml of MilliQ water and
1.0 ml of sodium pyrophosphate) after sonication (3 min) and
centrifugation was withdrawn and treated for counting as described
above; (ii) the remaining supernatant was carefully discharged, the
pellet was resuspended with 5 ml of MilliQ water, shaken for 1 min, and
centrifuged again, an aliquot of the supernatant was withdrawn, and
viruses were counted as described above; and (iii) this procedure was
repeated three times (since after the third washing, less than 5% of
the total virus abundance was encountered).
Interference with virus enumeration due to extracellular DNA.
In order to eliminate uncertainties in virus counting due to
extracellular DNA interference, we tested the effect of nuclease treatment on sediment samples. Twenty-five microliters of DNase I from
bovine pancreas (1.9 U ml
1), 10 µl of nuclease P1 from
Penicillium citrinum (4 U ml1), 10 µl of
nuclease S1 from Aspergillus orizae (2.3 U
ml1), and 10 µl of esonuclease 3 from Escherichia
coli (1.9 U ml1) were added to 1.0-ml aliquots of
the supernatant obtained from fresh sandy sediments and incubated for
15 min at room temperature. Additional aliquots of the supernatant (1.0 ml) without enzymes were incubated under the same conditions and served
as controls.
Comparison of EFM and TEM counts.
Virus enumeration performed
by EFM from fixed sandy sediments was compared to TEM virus counting.
For TEM analyses, we omitted the preconcentration procedure (i.e.,
ultracentrifugation) before mounting the sample on a grid
(14), since the number of virus particles in sediment
samples is expected to be very high (6, 21, 25).
Subaliquots of 10 µl of the supernatant obtained as described for EFM
counting were put onto 400-mesh Formvar-coated Cu grids, stained with
2% uranyl acetate, and dried under silica gel (14). The
grids were examined by a Philips CM 200 TEM at a magnification of
×38,000, and view fields were counted until the total counts exceeded 200.
Effect of preservatives in long-term experiments.
After
sampling was done, about 50 ml of fresh sandy sediment was gently mixed
and carefully split into three subaliquots: one was unfixed and
immediately analyzed, the second was fixed with formalin (2% final
concentration), and the last was fixed with glutaraldehyde (2% final
concentration). The sediment samples were processed in triplicate as
described above, but the pyrophosphate final concentration was reduced
to 5 mM as suggested by Epstein and Rossel (11) for
benthic bacterial extraction from coarser sediments. Fixed samples
stored at 4°C were analyzed after 1, 7, 30, and 90 days.
Statistical analyses.
To test differences between virus
counts, the t test was employed throughout the study. Its
use was justified, as (i) the density data were normally distributed
(they were checked by a standard graphical test [24])
and (ii) the test for homogeneity of variances (the
Fmax test [24]) showed that our
samples were homoscedastic. VLP dispersion was evaluated to test the
homogeneity of the virus distribution by calculating the coefficient of
variation (CV) (CV = standard deviation/mean ×100).
Deep-sea sediment samples incubated with sodium pyrophosphate displayed
higher virus counts than untreated samples [(14.6 ± 2.79) × 109 and (9.94 ± 6.12) × 109
viruses g1 (dry weight) of sediment, respectively], but
the differences were not statistically significant (t test;
P = 0.296). However, pyrophosphate-treated samples were
characterized by significantly lower CVs (19.1 versus 61.6%;
t test; P < 0.05). The effects of sonication on virus counts carried out on both abyssal and coastal sediments are reported in Fig. 1. The
highest virus counts for both sediment types (11.1 × 109 and 1.13 × 109 viruses
g1 for muddy and sandy sediments, respectively) were
obtained after 3 min of sonication and were significantly higher (two-
to four fold) (t test; P < 0.01) than
values obtained without sonication (i.e., with simple shaking). Further
increases in the sonication time decreased virus counts, and a
sonication lasting 15 min reduced virus counts by about 1 order of
magnitude (t test; P < 0.01 for both
sediment types).
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FIG. 1.
Effect of sonication on virus abundance in deep-sea (a)
and sandy (b) sediments. The standard deviations are shown.
|
|
The efficiency of virus extraction by pyrophosphate-ultrasound
treatment was ca. 60% (Fig.
2). The
abundances of viruses extracted
by this procedure were significantly
lower than the total cumulative
virus abundance (
t test;
P < 0.01). The subsequent first and second
washings
recovered 27.5 and 9.0% of the total virus abundance,
and after the
third wash step, <5% was recovered.
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|
FIG. 2.
Postsonication extraction efficiency of virus recovery
from deep-sea sediments. The standard deviations are shown.
|
|
The virus abundances obtained from nuclease-treated samples were
significantly higher than those in untreated samples [(5.11
± 0.15) × 10
8 and (4.62 ± 0.19) × 10
8 viruses g
1 (dry weight) of sediment,
respectively (
t test;
P < 0.05)]. Counts
obtained by TEM and EFM (from fixed sediment samples collected
at
Falconara) displayed no significant differences [(1.54 ± 0.15)
× 10
9 and (1.46 ± 0.13) × 10
9 viruses g
1 (dry weight) of sediment,
respectively (
t test;
P = 0.513)].
Sediments fixed with both formaldehyde and glutaraldehyde showed a
significant decrease in virus numbers after only 24 h of
preservation (21 and 23%, respectively [
t test;
P < 0.01]) (Fig.
3).
After 7 days of formaldehyde and glutaraldehyde storage, virus
abundance decreased from 30 to 40%, respectively, compared to
that in
fresh sediment samples and then remained constant for
up to 90 days in
storage.
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|
FIG. 3.
Effects of preservatives on virus abundance in sandy
sediments. Reported are formaldehyde (a) and glutaraldehyde (b)
long-term storage. The standard deviations are shown.
|
|
Virus identification by EFM has been attempted since the early 1990s
(
14,
22,
26), and the uncertainty in recognizing
viruses
has induced scientists to refer to the products of these
counts as VLP
(
2,
3). In this study, we employed SYBR Green
I, which has
the highest staining efficiency under EFM and flow
cytometry
(
17) compared to other fluorochromes (such as DAPI
[4',6'-diamidino-2-phenylindole] and Yo-Pro I [
20]).
Since all
these fluorochromes bind to nucleic acids, the possibility
that
some small bacteria (<0.3-µm diameter) may be counted as
viruses
cannot be excluded. Noble and Fuhrman (
20)
reported that even
if all small bacteria were counted as viruses, the
overestimation
of the total virus counts would be negligible. In this
study,
utilizing SYBR Green I at a concentration higher than that
previously
reported for virus counts in water samples (
17,
20), we noticed
that virus-size fluorescent particles displayed
a higher brightness
than bacterial fluorescence. Also, the confidence
in virus counts
reported in this study is based on two factors: (i) TEM
and EFM
provided almost identical estimates of virus density and (ii)
extracellular-DNA removal increased virus counts. In this regard,
Drake
et al. (
9) reported no significant differences between
the
VLP densities of DNase-treated pore water samples and control
samples.
In this study, surprisingly, we found significantly higher
virus
densities in nuclease-treated samples. As sediments might
contain large
amounts of extracellular DNA (
5,
7,
8),
such increased
yields due to nuclease treatment could be related
to reduction of the
fluorescence noise (i.e., reducing the masking
effect) due to SYBR
staining of extracellular DNA, facilitating
virus
counting.
Previous studies proved that the use of 10 mM pyrophosphate allowed the
highest virus yields from formaldehyde-preserved freshwater
sediments
(
16). In accordance with this, we found that the use
of
higher pyrophosphate concentrations made the optical field
opalescent
under EFM, thus making virus counting difficult. Pyrophosphate
treatment did not significantly increase virus extraction, but
the use
of this surfactant at a final concentration not exceeding
10 mM
significantly lowered CVs (about threefold lower than in
untreated
samples). Similar results have been reported for benthic
bacteria
(
11), indicating that the use of pyrophosphate increases
counting accuracy under EFM so that a smaller number of replicate
analyses is required to obtain acceptable CV
values.
Maranger and Bird (
16) utilized 45s of sonication to
dislodge viruses from sediment samples. The results of the present
study confirm that sonication is crucial for maximizing virus
recovery
from marine sediments. However, the optimal time of sonication
to
achieve the highest virus yields was never tested before. In
this
study, the highest extraction efficiency from both sediment
types was
obtained with 3 min of sonication. Similar results have
been reported
for bacterial extraction from different sediment
types (
4,
11). As observed for benthic bacteria, longer sonication
times
(up to 15 min) reduced virus abundance to 1/10.
The extraction efficiency of benthic viruses after pyrophosphate and
sonication treatment was approximately 60% of the total
cumulative
virus number. Similarly low extraction efficiencies
were reported for
benthic bacteria by Ellery and Schleyer (
10),
which
suggested the need for a correction factor (of 1.44) for
inadequacy in
bacterial
extraction.
Xenopoulos and Bird (
29) observed a dramatic decrease in
virus abundance (up to 75%) in water samples during the first 4
weeks
of formaldehyde storage. A significant decrease in virus
counts after
glutaraldehyde preservation has also been observed
in samples of
Phaeocystis infected by viruses (
17).
Similarly,
Turley and Hughes (
27) reported a bacterial
decrease of 39%
during the first 40 days of seawater sample storage
with glutaraldehyde.
In accordance with these findings, we found that
both formaldehyde
and glutaraldehyde storage caused significant
reductions in virus
counts after only 24 h of preservation. After
7 to 90 days of
preservation, no further virus loss was
observed.
| |
ACKNOWLEDGMENTS |
We acknowledge the support from the European Commission's Marine
Science and Technology Program (MAST III) under Mass Transfer and
Ecosystem Response (MATER; MAS3-CT-960051) and High-Resolution Spatial
and Temporal Study of the Benthic Biology and Geochemistry in a
NorthEastern Atlantic Abyssal Locality (BENGAL; MAS3-CT-950018).
We thank Monica Armeni (University of Ancona) for her technical
assistance and Tanya Hall (University of Athens) for improving the
English form.
| |
FOOTNOTES |
*
Corresponding author. Mailing address: Institute of
Marine Science, University of Ancona, Via Brecce Bianche, 60131 Ancona, Italy. Phone: 39 71 220 4654. Fax: 39 71 220 4650. E-mail:
danovaro{at}popcsi.unian.it.
| |
REFERENCES |
| 1.
|
Berg, Ø.,
K. Y. Borsheim,
G. Bratbak, and M. Heldal.
1989.
High abundance of viruses found in aquatic environments.
Nature
340:467-468[CrossRef][Medline].
|
| 2.
|
Borsheim, K. Y.,
G. Bratbak, and M. Heldal.
1990.
Enumeration and biomass estimation of planktonic bacteria and viruses by transmission electron microscopy.
Appl. Environ. Microbiol.
56:352-356[Abstract/Free Full Text].
|
| 3.
|
Brussaard, C. P. D.,
R. S. Kempers,
A. J. Kop,
R. Riegman, and M. Heldal.
1996.
Virus-like particles in a summer bloom of Emiliana huleyi in the North Sea.
Aquat. Microb. Ecol.
10:105-113.
|
| 4.
|
Danovaro, R.,
A. Feminò, and M. Fabiano.
1994.
Comparison between different methods for bacterial counting in marine sandy sediments.
Boll. Mus. Ist. Biol. Univ. Genova
58:142-152.
|
| 5.
|
Danovaro, R.,
A. Dell'Anno,
A. Pusceddu, and M. Fabiano.
1999.
Nucleic acid concentrations (DNA, RNA) in the continental and deep-sea sediments of the Eastern Mediterranean: relationships with seasonally varying organic inputs and bacterial dynamics.
Deep-Sea Res.
46:1077-1094.
|
| 6.
|
Danovaro, R., and M. Serresi.
2000.
Viral density and virus-to-bacterium ratio in deep-sea sediments of the Eastern Mediterranean.
Appl. Environ. Microbiol.
66:1857-1861[Abstract/Free Full Text].
|
| 7.
|
Dell'Anno, A.,
M. Fabiano,
G. C. A. Duineveld,
A. Kok, and R. Danovaro.
1998.
Nucleic acid (DNA, RNA) quantification and RNA/DNA ratio determination in marine sediments: comparison of spectrophotometric, fluorometric, and high-performance liquid chromatography methods and estimation of detrital DNA.
Appl. Environ. Microbiol.
64:3238-3245[Abstract/Free Full Text].
|
| 8.
|
Dell'Anno, A.,
M. Fabiano,
M. L. Mei, and R. Danovaro.
1999.
Pelagic-benthic coupling of nucleic acids in an abyssal location of the northeastern Atlantic Ocean.
Appl. Environ. Microbiol.
65:4451-4457[Abstract/Free Full Text].
|
| 9.
|
Drake, L. A.,
K. H. Choi,
A. G. E. Haskell, and F. C. Dobbs.
1998.
Vertical profiles of virus-like particles and bacteria in the water column and sediments of Chesapeake Bay, USA.
Aquat. Microb. Ecol.
16:17-25.
|
| 10.
|
Ellery, W. N., and M. H. Scheyer.
1984.
Comparison of homogenization and ultrasonication as techniques in extracting attached sedimentary bacteria.
Mar. Ecol. Prog. Ser.
15:247-250.
|
| 11.
|
Epstein, S. E., and J. Rossel.
1995.
Enumeration of sandy sediment bacteria: search for optimal protocol.
Mar. Ecol. Prog. Ser.
117:289-298.
|
| 12.
|
Fuhrman, J. A., and R. T. Noble.
1995.
Viruses and protists cause similar bacterial mortality in costal seawater.
Limnol. Oceanogr.
40:1236-1242.
|
| 13.
|
Fuhrman, J. A.
1999.
Marine viruses and their biogeochemical and ecological effects.
Nature
399:541-548[CrossRef][Medline].
|
| 14.
|
Hara, S.,
K. Terauchi, and I. Koike.
1991.
Abundance of viruses in marine waters: assessment by epifluorescence and transmission electron microscopy.
Appl. Environ. Microbiol.
57:2731-2734[Abstract/Free Full Text].
|
| 15.
|
Hennes, J. E., and C. A. Suttle.
1995.
Direct counts of viruses in natural waters and laboratory cultures by epifluorescence microscopy.
Limnol. Oceanogr.
40:1050-1055.
|
| 16.
|
Maranger, R., and D. E. Bird.
1996.
High concentrations of viruses in the sediments of Lac Gilbert, Quebec.
Microb. Ecol.
31:141-151.
|
| 17.
|
Marie, D.,
C. P. D. Brussaard,
R. Thyrhaug,
G. Bratbak, and D. Vaulot.
1999.
Enumeration of marine viruses in culture and natural samples by flow cytometry.
Appl. Environ. Microbiol.
65:45-52[Abstract/Free Full Text].
|
| 18.
|
Middelboe, M.,
N. O. G. Jørgensen, and N. Kroer.
1996.
Effects of viruses on nutrient turnover and growth efficiency of noninfected marine bacterioplankton.
Appl. Environ. Microbiol.
62:1991-1997[Abstract].
|
| 19.
|
Newton, P. R.,
R. S. Lampitt,
T. D. Jickell,
P. King, and C. Boutle.
1994.
Temporal and spatial variability of biogenic particle fluxes during the JGOFS NE-Atlantic process studies at 47°N 20°W.
Deep-Sea Res.
41:1617-1642.
|
| 20.
|
Noble, R. T., and J. A. Fuhrman.
1998.
Use of SYBR Green I for rapid epifluorescence counts of marine viruses and bacteria.
Aquat. Microb. Ecol.
14:113-118.
|
| 21.
|
Paul, J. H.,
J. B. Rose,
S. C. Jiang,
C. A. Kellogg, and L. Dickson.
1993.
Distribution of viral abundance in the reef environment of Key Largo, Florida.
Appl. Environ. Microbiol.
59:718-724[Abstract/Free Full Text].
|
| 22.
|
Proctor, L. M., and J. A. Fuhrman.
1992.
Mortality of marine bacteria in response to enrichments of the virus size fraction from seawater.
Mar. Ecol. Prog. Ser.
87:283-293.
|
| 23.
|
Rice, A. L.,
M. H. Thurston, and B. J. Bett.
1994.
The IOSDL DEEPSEAS programme: introduction and photographic evidence for the presence and absence of a seasonal input of phytodetritus at contrasting abyssal sites in the northern-eastern Atlantic.
Deep-Sea Res.
41:1305-1320.
|
| 24.
|
Sokal, R. R., and F. J. Rohlf.
1987.
Introduction to biostatistics, 2nd ed.
W. H. Freeman, San Francisco, Calif.
|
| 25.
|
Steward, G. F.,
D. C. Smith, and F. Azam.
1996.
Abundance and production of bacteria and viruses in the Bering and Chukchi Sea.
Mar. Ecol. Prog. Ser.
131:287-300.
|
| 26.
|
Suttle, C. A.,
A. M. Chan, and M. T. Cottrell.
1990.
Infection of phytoplankton by viruses and reduction of primary productivity.
Nature
387:467-469[CrossRef].
|
| 27.
|
Turley, C. M., and D. J. Hughes.
1992.
Effects of storage on direct estimates of bacterial numbers of preserved seawater samples.
Deep-Sea Res.
39:375-394.
|
| 28.
|
Weinbauer, M. G., and C. A. Suttle.
1997.
Comparison of epifluorescence and transmission electron microscopy for counting viruses in natural marine waters.
Aquat. Microb. Ecol.
13:225-232.
|
| 29.
|
Xenopoulos, M. A., and D. F. Bird.
1997.
Virus à la sauce Yo-Pro: microwave enhanced staining for counting viruses by epifluorescence microscopy.
Limnol. Oceanogr.
42:1648-1650.
|
Applied and Environmental Microbiology, March 2001, p. 1384-1387, Vol. 67, No. 3
0099-2240/01/$04.00+0 DOI: 10.1128/AEM.67.3.1384-1387.2001
Copyright © 2001, American Society for Microbiology. All rights reserved.
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-
Lam, P., Cowen, J. P.
(2004). Processing Deep-Sea Particle-Rich Water Samples for Fluorescence In Situ Hybridization: Consideration of Storage Effects, Preservation, and Sonication. Appl. Environ. Microbiol.
70: 25-33
[Abstract]
[Full Text]
-
Williamson, K. E., Wommack, K. E., Radosevich, M.
(2003). Sampling Natural Viral Communities from Soil for Culture-Independent Analyses. Appl. Environ. Microbiol.
69: 6628-6633
[Abstract]
[Full Text]
-
Ashelford, K. E., Day, M. J., Fry, J. C.
(2003). Elevated Abundance of Bacteriophage Infecting Bacteria in Soil. Appl. Environ. Microbiol.
69: 285-289
[Abstract]
[Full Text]
-
Rohwer, F., Edwards, R.
(2002). The Phage Proteomic Tree: a Genome-Based Taxonomy for Phage. J. Bacteriol.
184: 4529-4535
[Abstract]
[Full Text]
-
Danovaro, R., Manini, E., Dell'Anno, A.
(2002). Higher Abundance of Bacteria than of Viruses in Deep Mediterranean Sediments. Appl. Environ. Microbiol.
68: 1468-1472
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[Full Text]
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