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Applied and Environmental Microbiology, September 1999, p. 3820-3827, Vol. 65, No. 9
0099-2240/99/$04.00+0
Copyright © 1999, American Society for Microbiology. All rights reserved.
Marine Bacterial Isolates Display Diverse Responses
to UV-B Radiation
Fabien
Joux,1,2,*
Wade H.
Jeffrey,1
Philippe
Lebaron,2 and
David L.
Mitchell3
Center for Environmental Diagnostics and
Bioremediation, University of West Florida, Pensacola, Florida
325141; Observatoire
Océanologique, Centre National de la Recherche Scientifique
(CNRS-UMR 7621), Institut National des Sciences de l'Univers et
Université Paris VI, Banyuls-sur-Mer 66651, France2; and Department of
Carcinogenesis, The University of Texas M. D. Anderson Cancer
Center, Smithville, Texas 789573
Received 30 March 1999/Accepted 16 June 1999
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ABSTRACT |
The molecular and biological consequences of UV-B radiation were
investigated by studying five species of marine bacteria and one
enteric bacterium. Laboratory cultures were exposed to an artificial
UV-B source and subjected to various post-UV irradiation treatments.
Significant differences in survival subsequent to UV-B radiation were
observed among the isolates, as measured by culturable counts.
UV-B-induced DNA photodamage was investigated by using a highly
specific radioimmunoassay to measure cyclobutane pyrimidine dimers
(CPDs). The CPDs determined following UV-B exposure were comparable for
all of the organisms except Sphingomonas sp. strain RB2256,
a facultatively oligotrophic ultramicrobacterium. This organism
exhibited little DNA damage and a high level of UV-B resistance.
Physiological conditioning by growth phase and starvation did not
change the UV-B sensitivity of marine bacteria. The rates of
photoreactivation following exposure to UV-B were investigated by using
different light sources (UV-A and cool white light). The rates of
photoreactivation were greatest during UV-A exposure, although diverse
responses were observed. The differences in sensitivity to UV-B
radiation between strains were reduced after photoreactivation. The
survival and CPD data obtained for Vibrio natriegens when
we used two UV-B exposure periods interrupted by a repair period
(photoreactivation plus dark repair) suggested that photoadaptation
could occur. Our results revealed that there are wide variations in
marine bacteria in their responses to UV radiation and subsequent
repair strategies, suggesting that UV-B radiation may affect the
microbial community structure in surface water.
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INTRODUCTION |
Marine bacteria may account for up
to 90% of the cellular DNA in oceanic environments (11,
42). These organisms play a central role in the cycling of
nutrients in aquatic ecosystems and are a fundamental link in the
carbon transfer process (i.e., the microbial loop
[3]). Therefore, the study of factors that control
bacterial growth in the environment is of primary importance. Among the
different factors that affect bacterial growth, solar UV radiation
(UVR) (wavelengths, 290 to 400 nm) has only recently received
attention. UVR could be particularly deleterious for bacteria because
these organisms have simple haploid genomes with little or no
functional redundancy and are small, which precludes effective cellular
shading or protective pigmentation (13). The last conclusion
is supported by the observation that more photodamage is induced in
bacterioplankton than in larger eukaryotic plankton by the same amount
of solar UVR (20, 21). The detrimental effects of sunlight
on bacterioplankton are manifested by reduced DNA and protein synthesis
(1, 18), reduced exoenzymatic activity (36),
reduced amino acid uptake (4), reduced oxygen consumption (39), and a decrease in bacterial abundance (36,
39).
DNA photodamage by UVR is wavelength dependent. UV-A (wavelengths, 320 to 400 nm) causes only indirect damage to DNA, proteins, and lipids
through reactive oxygen intermediates. UV-B (wavelengths, 290 to 320 nm) causes both indirect and direct damage because of the strong
absorption of wavelengths below 320 nm by DNA. The most abundant
products formed during irradiation with UV-B are the cyclobutane
pyrimidine dimers (CPDs) (35). A CPD can be lethal if the
lesion blocks DNA synthesis and RNA transcription or can be mutagenic
if the lesion is bypassed by DNA polymerase. Induction of CPD formation
in marine bacterioplankton under laboratory and field UVR conditions
has been studied by Jeffrey and coworkers (20, 21).
In response to UV damage, bacteria have developed different repair
pathways, including photoenzymatic repair (PER), nucleotide excision
repair (NER) (also called dark repair), and recombinational repair
(postreplication repair). PER involves direct monomerization of CPDs by
a single enzyme (photolyase) with near-UV or visible light as a source
of energy (45). PER has been observed in marine bacterioplankton exposed to artificial UV-B after secondary irradiation with UV-A or photosynthetically active radiation (PAR) (22). The existence of PER in natural populations has been inferred from the
results of CPD kinetic analyses performed with bacterioplankton samples
obtained from the marine water column throughout the solar day
(21). Similar field studies have also shown that marine bacterioplankton repair much of the DNA damage at night after diurnal
exposure to sunlight, indicating that NER is a fundamental repair
mechanism as well (21, 33, 39). Species-specific differences
in UV-B sensitivity have been found in algae (23), cyanobacteria (51), ciliates (53), and
metazooplankton (54). Studies of photobiological responses
of marine bacterial species are very rare (17), and little
is known about the diverse responses of marine bacteria during UV
exposure and subsequent recovery. Understanding these responses is
crucial for predicting the extent to which UVR may affect bacterial
diversity by selecting phototolerant species or by inducing
photoadaptation. Therefore, we studied the molecular and biological
responses of five marine bacteria to UV-B radiation under standardized
experimental conditions. By measuring survival and induction and repair
of CPDs under different light conditions, we estimated the relative
contributions of dark- and light-dependent repair systems to recovery.
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MATERIALS AND METHODS |
Bacterial strains and growth procedures.
Sphingomonas
sp. strain RB2256, a facultatively oligotrophic ultramicrobacterium,
was isolated from natural seawater from Resurrection Bay, Seward,
Alaska, by an extinction seawater method; this organism was a
numerically important member of the total community at the time of
sampling (46). Strain RB2256 shares several features with
indigenous marine bacteria, including a small cell size and a low
apparent DNA content (46). Other marine bacteria, including
Vibrio natriegens ATCC 14048, Pseudoalteromonas haloplanktis CIP 103197T (Collection Institut Pasteur,
Paris, France), Deleya aquamarina CIP 74.8T, and
Pseudomonas stutzeri (49), were chosen because
they are frequently isolated from marine environments on high-nutrient media. The responses of marine bacteria were compared to the responses of Salmonella typhimurium CIP 60.62T, an enteric
bacterium that can experience UV stress outside its normal habitat
(intestinal tracts), including wastewater discharge events. S. typhimurium was maintained on tryptone soy broth (TSB) (Becton
Dickinson) solidified with 15 g of Bacto Agar (Difco) liter
1. The marine bacteria were maintained on a complex
marine medium containing artificial seawater (ASWJP) (41)
supplemented with 1 g of yeast extract liter1 and
5 g of Bacto Peptone (Difco) liter1 (ASWJP+PY); this
medium was solidified with 15 g of Bacto Agar liter1. Cultures were grown at 37°C (S. typhimurium) or 25°C (marine bacteria) in 200-ml flasks
containing 100 ml of TSB (S. typhimurium) or 100 ml of
ASWJP+PY (marine bacteria) with shaking (200 rpm). One-milliliter
portions of overnight cultures were used as inocula. The growth phases
of cultures were determined from growth curves obtained by measuring
the optical density at 600 nm (OD600). Stationary-phase cells were obtained after growth for 6 h (S. typhimurium), 18 h (V. natriegens, P. haloplanktis, D. aquamarina, and P. stutzeri), or 48 h (strain RB2256). Depending on the
experiment, cells were harvested in the mid-log phase (maximum
OD600/2) or at the beginning of the stationary phase
(maximum OD600) by centrifugation at 6,000 × g
for 10 min at 10°C. The pellets were washed twice in 0.9% (wt/vol)
NaCl (S. typhimurium) or ASWJP (marine bacteria). Bacterial suspensions were diluted in order to obtain a concentration of 108 cells per ml, and the resulting preparations were used
for irradiation experiments.
UV-B irradiation.
A portion of each cell suspension was
transferred to a sterile plastic tissue culture dish (100 by 20 or 150 by 25 mm; Corning) so that the depth of the liquid was not greater than
2 mm. The petri dish (without a lid) was then placed in an incubator
and incubated at 20°C with slow shaking (50 rpm). The petri dish was covered with a sheet of Kodacel (Kodak, Rochester, N.Y.) to block out
UV-C (50% transmittance at 295 nm) and then exposed to four 20-W UV-B
lamps (Philips). UV-B irradiance was quantified with a radiometer
(model IL-1400A; International Light, Newburyport, Mass.) coupled to a
UV-B detector (26) (with maximum sensitivity at 295 nm). The
average intensity was 2.3 W m2. The UV dose (in joules
per square meter) was calculated by multiplying the intensity by the
exposure time (in seconds). The amount of PAR was determined with a
model GUV511 radiometer (Biospherical Instruments, San Diego, Calif.).
Photoreactivation and liquid holding.
UV-B-irradiated cell
suspensions were exposed to different types of photoreactivating light,
including UV-A light (20-W bulbs [Philips]) and visible light (20-W
cool white bulbs [General Electric]). The petri dishes containing
UV-B-irradiated cells were covered with a sheet of Mylar-D to block out
UV-B (cutoff, 320 nm) (1) and placed under two
photoreactivating lamps. The UV-A fluence rate was measured with a
model IL-1400A radiometer coupled to a UV-A detector (with maximal
sensitivity at 355 nm). The UV-A intensities used for the
photoreactivating treatments were 2.6 W m2 (UV-A
treatment) and 0.026 W m2 (0.0027 microeinsteins of
PAR/cm2/s) (visible light treatment). The cell suspensions
were continuously shaken at 50 rpm during photoreactivation.
The numbers of CFU per milliliter were corrected for possible
evaporation during exposure by using the following formula:
CFUcorr ml1 = (CFU × final
volume)/initial volume. Tests to determine possible liquid
holding recovery in the dark were performed by placing UV-B-irradiated
cell suspensions in plastic tubes in the dark and incubating them
without agitation at room temperature.
Starvation conditions.
The effects of starvation on both
UV-B resistance and PER capacity were assessed by recording the numbers
of CFU of S. typhimurium and V. natriegens
exposed to radiation for different times in nutrient-depleted
solutions. Erlenmeyer flasks (capacity, 500 ml) containing 200 ml of
0.9% NaCl (S. typhimurium) or 200 ml of ASWJP (V. natriegens) were autoclaved (120°C, 15 min), cooled, and
inoculated with cells from stationary-phase cultures washed as
described above. The flasks were incubated in the dark at 20°C. Subsamples were removed each day for 4 days. The cells were exposed to
UV-B radiation (950 and 1,650 J m2 for V. natriegens and S. typhimurium, respectively, resulting in about 0.1% survival for each strain) and then to UV-A for 2 h
(photoreactivation). The numbers of CFU were determined before and
after each radiation treatment.
Culturable counts.
A 100-µl portion of each cell
suspension was removed in order to prepare serial dilutions in 0.9%
NaCl (S. typhimurium) or ASWJP (marine bacteria). Portions
(100 µl) of the appropriate serial dilutions were spread in
triplicate onto TSB plates (S. typhimurium) or ASWJP+PY
plates (marine bacteria). The numbers of CFU were determined after
24 h of incubation in the dark at 37°C for S. typhimurium and after 2 to 7 days of incubation in the dark at
25°C for the marine bacteria. The dilution and plating procedures
were carried out under low-luminosity conditions to avoid any stray
photoreactivating light. The fraction of surviving cells was calculated
by dividing the number of CFU in the treated sample by the number of
CFU in the unirradiated sample at time zero.
DNA photoproducts.
Culture subsamples (5 to 15 ml) were
filtered through 0.22-µm-pore-size polysulfone membranes (Gelman),
and the membranes were then stored at 
20°C in microcentrifuge
tubes. Five hundred microliters of STE buffer (10 mM Tris [pH 8], 1 mM EDTA, 100 mM NaCl) containing 0.5% sodium dodecyl sulfate was added
to each tube, and then the tubes were capped, vortexed for 30 s,
and placed in a boiling water bath for 2 min. After they were chilled
on ice for 5 min, the filters were removed, and the DNA was extracted with 0.8 ml of chloroform-isoamyl alcohol (49:1). The aqueous phase was
collected after centrifugation at 3,000 × g at 4°C
for 30 min. To precipitate the nucleic acids, 1 µl of a 25-mg
ml1 glycogen solution, 0.1 volume of 3 M sodium acetate
(pH 5), and an equal volume of isopropanol were added to each
preparation, and the preparations were incubated overnight at 20°C.
Each precipitate was collected by centrifugation at 13,000 × g at 4°C for 30 min. The pellet was washed with 0.5 ml of
cold 70% ethanol and resuspended in 0.5 ml of STE buffer. DNA was
quantified by fluorometry with Hoechst 33258 dye (43).
CPDs were quantified by using a highly sensitive radioimmunoassay
described by Mitchell (34). This assay measures the binding of radiolabeled UV-irradiated DNA to antibody raised in rabbits against
triplet (acetone)-sensitized UV-B-irradiated DNA, conditions which
induce cyclobutane dimers exclusively. The quantity of CPDs was
determined by comparison to standard DNA (pUC19) for which the
rates of photoproduct formation were known. Samples were analyzed in duplicate, and the amount of CPD that accumulated was compared to
the amount in a solution of calf thymus DNA previously exposed to
the same UV-B dose.
Flow cytometric detection of DNA content.
Bacterial samples
were fixed with 2% (vol/vol) (final concentration) formaldehyde and
stored in the dark at 4°C until they were analyzed. The DNA content
was determined as described by Lebaron et al. (27) by using
SYBR-I (Molecular Probes Inc., Eugene, Oreg.). SYBR-I was added at a
final concentration of 104 M to the stock solution, and
the preparation was incubated for 15 min in the dark with 30 mM (final
concentration) potassium citrate (pH 7.4). All experiments were
performed with a FACS-Calibur flow cytometer (Becton Dickinson)
equipped with an air-cooled laser providing 15 mW at 488 nm and the
standard filter setup. All parameters were measured as logarithmic
signals. Green fluorescence was determined in the FL1 channel (530 ± 15 nm). Relative cellular sizes were estimated by using the side
scatter signal. Yellow-green fluorescent microspheres (2-µm-diameter
fluorescent size standard beads; Polysciences Inc., Warrington, Pa.)
were systematically added to each sample as an internal reference to
normalize cell fluorescence and side scatter values.
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RESULTS |
Diversity of UV-B resistance.
Although the UV-B intensity used
in this study (2.3 W m2) was higher than the natural
intensity, the UV-B doses applied (up to 3,000 J m2) were
similar to daily UV-B doses recorded in the field at the surface of
water (8). To ensure that the measured biological responses
were reliable, we previously tested the principle of reciprocity, which
requires that a response be a function of the dose and not a function
of the dose rate. For reciprocity to be satisfied, different
combinations of dose rate and exposure time, which result in the same
dose, should yield the same response. Stationary-phase cultures of
S. typhimurium were exposed to three UV-B intensities (0.5, 1.1, and 2.3 W m2) for different times, which resulted in
eight doses (0 to 2,000 J m2) for each intensity. The
inactivation rates (loss of CFU) obtained for the three UV-B
intensities did not differ (data not shown). The high cell densities
used in this study did not introduce bias through a protective effect;
the inactivation rates obtained with different S. typhimurium cell densities (106 to 108
cells per ml) were comparable, indicating that shielding did not occur
(data not shown).
With stationary-phase cells of most bacteria exposed to artificial UV-B
radiation there was an exponential decrease in the
number of CFU. The
only exception was strain RB2256; a shoulder
was observed at doses up
to 500 J m
2 with this strain (Fig.
1). Large variations in the UV-B survival
rates were observed with the different organisms. The marine bacteria
could be divided into the following three resistance categories:
sensitive (
V. natriegens and
P. haloplanktis),
intermediate (
P. stutzeri and
D. aquamarina), and resistant (strain RB2256). Ninety-nine
percent inactivation was observed after 800-, 1,600-, and
3,000-J
m
2 UV-B doses with members of the sensitive,
intermediate, and resistant
groups, respectively. The response of
S. typhimurium was similar
to the response of the
sensitive strains at doses up to 1,000
J m
2 but resembled
the response of the intermediate strains at higher
UV-B doses.
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FIG. 1.
Effects of UV-B radiation on the survival of different
bacteria collected during the stationary phase. The data are expressed
as percentages of the initial counts (~108 CFU
ml1) and are means (± standard deviations) based on the
data from two different experiments. Symbols:  , V. natriegens;  , P. haloplanktis;  , S. typhimurium;  , P. stutzeri;  , D. aquamarina;  , strain RB2256.
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Accumulation of DNA photoproducts.
Because DNA is the primary
target for UV-B radiation, it was important to estimate variations in
DNA content and distribution in the different bacteria. Flow cytometric
DNA histograms showed that bacterial populations harvested in the
stationary phase were mainly composed of cells containing a single
genome (Table 1). On the basis of mean
fluorescence values for cells containing a single genome, we determined
that the chromosome of strain RB2256 was 1.5 to 2 times smaller than
the chromosomes of the other strains (Table 1). The cells of strain
RB2256 were also smaller, as estimated by the side scatter signal
(Table 1).
To determine if the diversity of UV-B resistance resulted mainly from
effective protection or DNA repair, CPDs were quantified
after two
doses of irradiation (1,000 and 2,000 J m
2) (Fig.
2). DNA damage frequencies (expressed as
the number of
CPDs per megabase of DNA) were determined for
stationary-phase
cells immediately following UV-B irradiation and prior
to initiation
of any DNA repair processes. The number of CPDs per
megabase of
DNA in a solution of calf thymus DNA was comparable to the
values
recorded for purified DNA exposed to 1 solar day at the surface
of marine water (between 500 and 1,750 CPDs/Mb of DNA) (
1,
20,
29,
44), indicating that the UV-B doses used in this
study were
comparable to natural daily doses.
V. natriegens,
P. haloplanktis, and
D. aquamarina accumulated the same
amounts of
CPDs after different UV-B doses despite their different
levels
of resistance. Surprisingly, the levels of CPDs obtained for
these
strains were equivalent to the level obtained for the DNA
solution
exposed to the same UV-B dose, suggesting that no cellular
photoprotective
mechanisms were present. In contrast, strain RB2256
accumulated
very small amounts of CPDs after different UV-B doses. The
amount
of CPDs in strain RB2256 was fourfold less than the amounts in
the other marine bacteria after a 2,000-J m
2 dose. After
a 1,000-J m
2 UV-B dose, the level of DNA damage in
S. typhimurium was intermediate
between the levels of DNA
damage in strain RB2256 and the other
marine bacteria. With higher UV-B
doses, the
S. typhimurium damage
frequencies were comparable
to the damage frequencies observed
in the DNA solution.
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FIG. 2.
Accumulation of CPDs in marine bacteria and S. typhimurium during UV-B irradiation. The data are means (± standard deviations) based on the data from duplicate analyses.
Symbols: , V. natriegens; , P. haloplanktis; , S. typhimurium; , D. aquamarina; , strain RB2256; , DNA solution.
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Photoreactivation and liquid holding recovery.
PER of bacteria
after UV-B irradiation was investigated by using different light
sources (UV-A and cool white light). With all strains,
photoreactivation was greatest after UV-A exposure; maximum survival
was observed after 2 h of exposure, which corresponded to a dose
of 19 kJ m2 (Fig. 3). Under
photoreactivating light conditions, V. natriegens, P. stutzeri, and strain RB2256 exhibited similar levels of resistance to UV-B, and a shoulder was observed at doses up to 500 J
m2 (Fig. 4). Under these
conditions 99% inactivation was obtained after a 3,000-J
m2 UV-B dose. The very sensitive organism V. natriegens had a very effective photorepair mechanism. For this
strain, the number of cells that survived after a 1,500-J
m2 UV-B dose increased by more than four orders of
magnitude when the cells were allowed to photorepair via exposure to
UV-A. In contrast to the marine bacteria, S. typhimurium
exhibited effective photoreactivation only under UV-A irradiation
conditions, and the level of photorepair was less than the levels
observed in marine bacteria and exhibited no shoulder (i.e., repair
capacity). With S. typhimurium, 99% inactivation was
observed after a 2,250-J m2 UV-B dose.
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FIG. 3.
Effects of different light sources on photoreactivation
kinetics after UV-B exposure. V. natriegens, S. typhimurium, P. stutzeri, and strain RB2256 were
exposed to 1,500-, 1,650-, 2,000-, and 3,500-J m2 UV-B
doses, respectively, before they were exposed to photoreactivating
light. The data are expressed as percentages of the initial counts
(~108 CFU ml1) and are means (± standard
deviations) based on data obtained from triplicate plates.
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FIG. 4.
Effects of UV-B radiation without () and with ()
exposure to UV-A radiation for 2 h after UV-B treatment for
V. natriegens, S. typhimurium, P. stutzeri, and RB2256. The data are expressed as percentages of the
initial counts (~108 CFU ml1) and are means
(± standard deviations) based on the data from two different
experiments.
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The principle of reciprocity was also verified under photoreactivation
conditions. No differences in photorecovery were observed
when
stationary-phase cultures (
S. typhimurium was used as the
test organism) were exposed to UV-A light after equivalent UV-B
doses
were delivered by two UV-B intensities (0.5 and 2.3 W m
2)
(data not shown). To determine whether de novo protein synthesis
was
necessary for photoreactivation, we conducted experiments
with
V. natriegens (the most photoreactive organism in this study)
in the
presence of 50 µg of chloramphenicol ml
1 (the MIC of
chloramphenicol for this strain is approximately
2.5 µg
ml
1). After UV-B exposure, chloramphenicol was added, and
the cells
were exposed to UV-A for 2 h. Under these conditions,
photoreactivation
was not altered (data not shown). Of the different
strains tested
and within the time frame used (2 days), only
V. natriegens benefited
from a liquid holding period after UV-B
inactivation (Fig.
5).
Maximum recovery
of
V. natriegens occurred after 12 h of incubation
in
the dark. When liquid holding recovery was taken into account,
the resistance of
V. natriegens was improved and more
closely
resembled the resistance of the intermediate organisms
(i.e.,
D. aquamarina and
P. stutzeri) (data not
shown). Under these conditions
99% inactivation was observed
after a 1,500-J m
2 UV-B dose (data not shown).
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FIG. 5.
Kinetics of liquid holding recovery after UV-B
irradiation. V. natriegens, S. typhimurium,
P. stutzeri, and strain RB2256 were exposed to 950-, 1,650-, 2,000-, and 4,000-J m2 UV-B doses, respectively, before
liquid holding. The data are expressed as percentages of the initial
counts (~108 CFU ml1) and are means (± standard deviations) based on the data from two different experiments.
Symbols: , V. natriegens; , S. typhimurium;
, P. stutzeri; , strain RB2256.
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Effects of growth rate on UV-B resistance and photoreactivation
capacity.
We compared the UV-B resistance of V. natriegens, P. stutzeri, S. typhimurium,
and strain RB2256 under exponential- and stationary-phase growth
conditions. Figure 6 shows the ratios of
stationary-phase cells to exponential-phase cells that survived after
different doses of UV-B light. In both cases, the percentages of
survival were expressed as a function of the initial CFU counts. The
resistance of stationary-phase cells was slightly greater than (ratio,
less than 5) or comparable to the resistance of exponential-phase cells for the marine bacteria. A greater difference between stationary- and exponential-phase cells was observed with S. typhimurium
exposed to UV-B doses higher than 1,000 J m2.
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FIG. 6.
Influence of growth phase on the survival of V. natriegens, S. typhimurium, P. stutzeri, and
strain RB2256 after exposure to UV-B radiation. The data are ratios of
stationary-phase cells to exponential-phase cells that survived after
different UV-B doses. The percentages of survival at both growth phases
used to calculate ratios were determined relative to the initial counts
(~108 CFU ml1) and are means based on the
data from two different experiments. Symbols: , V. natriegens; , S. typhimurium; , P. stutzeri; , strain RB2256.
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In addition, the effects of starvation on UV-B resistance and
photoreactivation of
V. natriegens and
S. typhimurium were examined.
During 4 days of starvation, the
percentage of viable
S. typhimurium cells did not change,
and the percentage of viable
V. natriegens cells decreased
slightly (data not shown). For both strains, the
number of CFU after
UV-B exposure and UV-A exposure did not change
significantly during
starvation, indicating that these conditions
did not alter UV-B
resistance or photoreactivation capacity (data
not
shown).
Potential for photoadaptation.
In order to identify UV-B
photoadaptive responses, we treated V. natriegens and
S. typhimurium with 1,000 J of UV-B light m2; this was followed by 2 h of exposure to
photoreactivating UV-A light, 10 h of incubation in the dark, and
a second exposure to 1,000 J of UV-B light m2. At
each point during this protocol, the numbers of CFU and CPDs were
determined (Fig. 7). Under these
conditions, all of the CPDs that were accumulated by V. natriegens cells were removed after photoreactivation, and CFU
were recovered up to 46% of the initial count. During the dark period,
no change in the number of culturable cells was observed. The DNA
damage and sensitivity after the second UV-B exposure were considerably
less than the DNA damage and sensitivity after the first exposure. When
this protocol was used, the S. typhimurium cells removed
about one-half of the accumulated CPDs during the photoreactivation
phase, but the level of recovery of CFU was only 5%. Liquid holding
slightly decreased the number of CFU. The second exposure to UV-B
resulted in a less significant loss of CFU than the first exposure but
greater cumulative DNA damage (the number of CPDs was twice the number
after the initial exposure).
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FIG. 7.
Changes in CFU and CPDs under the following conditions:
bars A, control (stationary-phase cells); bars B, treatment A plus UV-B
radiation (1,000 J m2); bars C, treatment B plus
photoreactivation (UV-A radiation for 2 h); bars D, treatment C
plus liquid holding (10 h); bars E, treatment D plus UV-B radiation
(1,000 J m2). The CFU data are expressed as percentages
of the initial counts (~108 CFU ml1) and
are means (± standard deviations) based on data obtained from
triplicate plates. The numbers above the bars are the numbers of CPDs
per megabase of DNA and are means (± standard deviations) based on
data obtained from duplicate analyses. ND, not determined.
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DISCUSSION |
UV-B resistance and DNA damage.
Significant species-specific
differences in susceptibility to UV-B irradiation were observed among
bacterial strains, particularly when photodependent repair mechanisms
were excluded. When culture media were used to measure bacterial
resistance, sufficient nutrients were supplied for the energy
requirements of NER. In the case of V. natriegens, we found
that maintenance of cells in a nutrient-depleted solution after
irradiation and prior to plating resulted in increased survival. Such
liquid holding recovery has been demonstrated previously for
Escherichia coli strains after UV-C irradiation and, more recently, for the fish pathogen Vibrio anguillarum
(31). The mechanisms of this response are not yet
understood, but Ganesan and Smith (12) observed that liquid
holding recovery occurs in E. coli K-12 only when excision
repair, but not recombination repair, is operative. From these results
it is evident that in order to understand how low concentrations of
dissolved organic carbon in seawater constitute a limiting factor for
optimal repair, the precise energy requirements of NER must be determined.
In contrast to previous observations of Antarctic diatoms exposed to
artificial UV-B radiation (
23), equivalent DNA damage
frequencies did not correlate with cell killing in the sensitive
and
intermediate organisms. The bacteria investigated have different
G+C
contents, as follows: 41 to 45% for
P. haloplanktis
(
6),
46 to 47% for
V. natriegens (
5),
50 to 53% for
S. typhimurium (
28), 56% for
D. aquamarina (
2), 61 to 66% for
P. stutzeri (
40), and 61.6 to 67.8% for
Sphingomonas sp. strain RB2256 (
50).
Although a
high G+C content is known to protect DNA against damage
by thymine
dimerization (
48), the differences in G+C content
among the
bacteria studied here did not appear to affect the quantity
of CPDs
induced. Likewise, differences in the amount of DNA per
cell could not
explain the differences in resistance observed
between intermediate and
sensitive organisms since most cells
of all of the organisms tested in
the stationary phase had one
genome and the genomes were similar
sizes.
The fact that comparable levels of CPDs per megabase of DNA were
accumulated by sensitive and intermediate organisms and calf
thymus DNA
in solution indicated that there was a complete lack
of photoprotection
in these bacteria. In contrast, the high level
of resistance of strain
RB2256 cells to UV-B light was associated
with low CPD frequencies and
indicated that a constitutive photoprotective
mechanism was present in
these cells. The yellow pigmentation
in RB2256 cells is thought to be
insufficient to explain the resistance
of the cells to UV-B irradiation
since the small size of the cells
probably precludes effective
protection by pigmentation (
13).
Consistent with this
hypothesis, Gascón et al. (
14) observed
that strains
of
Rhodobacter sphaeroides with high levels of pigment
(associated with phototrophic growth) were more sensitive to UV-C
irradiation than strains with less pigment (associated with
heterotrophic
growth) were. In addition to pigments, DNA binding
proteins have
been implicated in photoprotection (
47).
Small, acid-soluble
proteins bind to the DNA of
Bacillus and
Clostridium spores and
provide strong protection against DNA
damage due to UV irradiation
and other stresses (e.g., heat and
hydrogen peroxide). It has
also been shown that certain
E. coli strains which lack specific
DNA binding proteins (protein HU)
are sensitive to UV-C and
irradiation (
7,
29). Since
these proteins mitigate DNA cleavage
by
rays (
7) but do
not reduce CPD frequencies after UV-C
irradiation (
29), they
are probably not involved in the photoprotection
observed in RB2256
cells. The mechanism behind the multiple resistance
of strain RB2256 to
a variety of exogenous stress-inducing agents,
such as heat shock,
H
2O
2, and ethanol (
9), is intriguing
and
worthy of future research
efforts.
Photoreactivation in marine bacteria.
Photoreactivation occurs
in each of the three marine organisms studied and appears to be an
important mechanism of DNA repair, especially for the sensitive
bacterium V. natriegens. When PER is taken into account, the
diversity of resistance is largely reduced. The different levels of
photoreactivation observed with marine bacteria may result from
differences in the specific activities of the individual DNA
photolyases or from differences in the intracellular concentrations of
the enzymes (37). Different photoreactivation capacities may
also occur in members of the same genus, as described previously for
six Bacillus species (37). Observations made with
V. natriegens treated with chloramphenicol indicated that the initial photolyase content is sufficient to ensure total
photoreactivation. Moreover, the photoreactivation capacity was not
attenuated by prior starvation with both V. natriegens and
S. typhimurium cells. Unlike NER and postreplication repair,
PER does not require energy mobilization and may therefore be very
important in aquatic microbial populations that are nutrient limited.
Indeed, photoreactivation may be the main repair mechanism that
operates immediately after exposure to solar UVR.
The rates of photoreactivation were greatest during UV-A exposure,
although diverse responses were observed. The marine organisms
responded to a broader spectrum for photoreactivation than
S. typhimurium, and efficient PER was observed under UV-A and cool
white light conditions. Photolyases are categorized on the basis
of the
chromophore involved in energy transfer, and
S. typhimurium photolyase belongs to the folate class (
30) and
exhibits maximum
activity at 384 nm (i.e., UV-A), which is
consistent with our
results. Other known photolyases fall into
either a deazaflavin
class (maximum activity at 440 nm) (
19)
or a "no-second-chromophore"
class (maximum activity at both 370 and 450 nm) (
24). The patterns
of photoreactivation observed
in marine strains suggest that the
efficiency of photoreactivation in
marine water should correlate
with UV-A penetration. Our results agree
with those of Kaiser
and Herndl (
22), who found that UV-A
radiation produced more
efficient photoreactivation than PAR produced
in both aged and
freshly collected
seawater.
Physiological conditioning and photoadaptation.
Our results
show that growth phase has little impact on the resistance of marine
bacteria to UV-B irradiation. Significant interstrain variability in
growth phase responses has been observed in different bacteria exposed
to UV-C radiation (10, 15, 16, 25), although the types of
damage induced by UV-C radiation were essentially the same as the types
of damage induced by UV-B light. Some workers observed increased
resistance (10, 16) but other studies revealed increased
sensitivity when cells entered the stationary phase (25).
Our results agree with the results of other studies and show that the
sensitivities of cells harvested from exponential- and stationary-phase
cultures did not differ (15). Our results show that
starvation of stationary-phase cells in ASWJP or an NaCl solution did
not change the UV-B resistance of the cells. In contrast, Nyström
et al. (38) showed that marine Vibrio sp. strain
S14 kept in seawater depleted of C, N, and P exhibited increased
resistance to UV-C and UV-B (302-nm) radiation. The lengths of
starvation time required by this strain to reach the maximal level of
stress resistance were 30 h for UV-C radiation and about 70 h
for UV-B radiation. In the same way, Enterococcus faecalis
cells in the exponential growth phase and the early stationary phase
became progressively more resistant to UV-C radiation when they were
incubated in tap water (an oligotrophic environment); maximum survival
occurred after 35 days (16). Many studies have shown that
stationary-phase or starved bacteria are far more resistant to
stresses, such as starvation, abnormal temperatures, hydrogen peroxide,
and UV-A radiation (photooxidation), than their exponentially growing
counterparts are. Expression of the genes responsible for this
increased resistance is controlled by the stationary-phase-specific
sigma factor 
S, which is encoded by rpoS
(32). To our knowledge, genes controlled by rpoS
responsible for UV-C or UV-B resistance have never been described. It
has been shown that mutations in the E. coli rpoS gene
(initially named nur) may reduce the efficiency of repair after exposure to 313-nm UV-B radiation but not after exposure to
290-nm UV-B radiation (52). This result may be explained by
the increase in the proportion of photooxidations to dimerizations in
the upper part of the UV-B spectrum (i.e., near the UV-A spectrum).
As demonstrated with
V. natriegens, photoadaptation could
occur after UV-B irradiation and subsequent recovery (photoreactivation
and dark holding). This photoadaptation resulted in increased
resistance and decreased CPD accumulation during a second UV-B
irradiation period. The reduced CPD frequency observed after the
second
irradiation suggests that
V. natriegens could mitigate
the
DNA damage by some inducible photoprotective mechanism. Whether
bacterial photoadaptation occurs in aquatic environments has not
been
clearly established. Pakulski et al. (
39) observed recovery
of bacterial production and respiration in subtropical coral reef
bacteria incubated at a fixed depth during a second day of exposure
to
natural sunlight. These authors explained these findings by
either
photoinduced selection for light-tolerant cells or physiological
adaptation to ambient light regimens that occurred during exposure.
Herndl et al. (
18) showed that no photoadaptation occurred
in
bacterioplankton obtained from surface water (depth, 0.5 m)
that
were as sensitive (as determined by thymidine incorporation) to
surface UV-B radiation as subpycnocline bacteria were (depth,
20 m). Based on the diversity of the photobiological responses
of
different marine bacterioplankton to solar UVR observed in
this study,
it is apparent that an integrated approach involving
molecular,
physiological, and taxonomic end points will be required
to better
understand the impact of UV-B radiation on bacterioplankton
populations. Our results demonstrate the variability of the responses
of marine bacteria that are exposed to UV-B radiation and emphasize
the
difficulties in deriving models that predict UV effects in
the
environment from laboratory studies. The clear differences
between the
photobiology of culturable marine bacteria isolated
on high-nutrient
media and the photobiology of the facultatively
oligotrophic
ultramicrobacterium
Sphingomonas sp. strain RB2256
should
encourage isolation of the latter type of organisms, which
may
constitute a significant part of the open ocean bacterial
community.
| |
ACKNOWLEDGMENTS |
We thank Staffan Kjelleberg (School of Microbiology and
Immunology, The University of New South Wales, Sydney, Australia) for
providing Sphingomonas sp. strain RB2256 and Jason Kase for critically reviewing the manuscript.
This work was supported by a postdoctoral fellowship to F.J. from the
Conseil Régional Languedoc-Roussillon (France) and by National
Science Foundation Office of Polar Programs grant OPP-9419037 to W.H.J.
| |
FOOTNOTES |
*
Corresponding author. Mailing address: Observatoire
Océanologique de Banyuls, CNRS-UMR 7621, INSU et Université
Paris VI, BP 44, F-66651 Banyuls-sur-Mer Cedex, France. Phone:
33-(0)4-68-88-73-42. Fax: 33-(0)4-68-88-73-95. E-mail:
joux{at}obs-banyuls.fr.
| |
REFERENCES |
| 1.
|
Aas, P.,
M. M. Lyons,
R. Pledger,
D. L. Mitchell, and W. H. Jeffrey.
1996.
Inhibition of bacterial activities by solar radiation in nearshore waters and the Gulf of Mexico.
Aquat. Microb. Ecol.
11:229-238.
|
| 2.
|
Akagawa, M., and K. Yamasato.
1989.
Synonymy of Alcaligenes aquamarina, Alcaligenes faecalis subsp. homari, and Deleya aesta: Deleya aquamarina comb. nov. as the type species of the genus Deleya.
Int. J. Syst. Bacteriol.
39:462-466[Abstract/Free Full Text].
|
| 3.
|
Azam, F.,
T. Fenchel,
J. G. Field,
J. S. Gray,
L. A. Meyer-Reil, and F. Thingstad.
1983.
The ecological role of water-column microbes in the sea.
Mar. Ecol. Prog. Ser.
10:257-263.
|
| 4.
|
Bailey, C. A.,
R. A. Neihof, and P. S. Tabor.
1983.
Inhibitory effect of solar radiation on amino acid uptake in Chesapeake Bay bacteria.
Appl. Environ. Microbiol.
46:44-49[Abstract/Free Full Text].
|
| 5.
|
Baumann, P.,
A. L. Furniss, and J. V. Lee.
1984.
Genus Vibrio, p. 518-538.
In
N. R. Krieg, and J. G. Holt (ed.), Bergey's manual of systematic bacteriology, vol. 1. The Williams & Wilkins Co., Baltimore, Md.
|
| 6.
|
Baumann, P.,
M. J. Gauthier, and L. Baumann.
1984.
Genus Alteromonas, p. 342-352.
In
N. R. Krieg, and J. G. Holt (ed.), Bergey's manual of systematic bacteriology, vol. 1. The Williams & Wilkins Co., Baltimore, Md.
|
| 7.
|
Boubrik, F., and J. Rouvière-Yaniv.
1995.
Increased sensitivity to irradiation in bacteria lacking protein HU.
Proc. Natl. Acad. Sci. USA
92:3958-3962[Abstract/Free Full Text].
|
| 8.
|
Demers, S.,
C. Belzile,
D. R. S. Lean,
B. Mostajir,
S. Roy,
S. De Mora,
D. Bird,
M. Gosselin,
J.-P. Chanut, and M. Levasseur.
1998.
An experimental tool to study the effects of ultraviolet radiations on planktonic communities: a mesocosm approach.
Environ. Technol.
19:667-682.
|
| 9.
|
Eguchi, M.,
T. Nishikawa,
K. MacDonald,
R. Cavicchioli,
J. C. Gottschal, and S. Kjelleberg.
1996.
Responses to stress and nutrient availability by the marine ultramicrobacterium Sphingomonas sp. strain RB2256.
Appl. Environ. Microbiol.
62:1287-1294[Abstract].
|
| 10.
|
Freedman, M. L., and A. K. Bruce.
1971.
The relationship of radioresistance to balanced growth-rate in Micrococcus radiodurans.
Int. J. Radiat. Biol.
19:111-121.
|
| 11.
|
Fuhrman, J. A.,
T. D. Sleeter,
C. A. Carlson, and L. M. Proctor.
1989.
Dominance of bacterial biomass in the Sargasso Sea and its ecological implications.
Mar. Ecol. Prog. Ser.
57:207-217.
|
| 12.
|
Ganesan, A. K., and K. C. Smith.
1968.
Dark recovery processes in Escherichia coli irradiated with ultraviolet light.
J. Bacteriol.
96:365-371[Abstract/Free Full Text].
|
| 13.
|
Garcia-Pichel, F.
1994.
A model for internal self-shading in planktonic organisms and its implications for the usefulness of ultraviolet sunscreen.
Limnol. Oceanogr.
39:1704-1717.
|
| 14.
|
Gascón, J.,
A. Oubiña,
A. Pérez-Lezaun, and J. Urmeneta.
1995.
Sensitivity of selected bacterial species to UV radiation.
Curr. Microbiol.
30:177-182[Medline].
|
| 15.
|
Hartke, A.,
S. Bouche,
X. Gansel,
P. Boutibonnes, and Y. Auffray.
1994.
Starvation-induced stress resistance in Lactococcus lactis subsp. lactis IL1403.
Appl. Environ. Microbiol.
60:3474-3478[Abstract/Free Full Text].
|
| 16.
|
Hartke, A.,
J.-C. Giard,
J. M. Laplace, and Y. Auffray.
1998.
Survival of Enterococcus faecalis in an oligotrophic microcosm: changes in morphology, development of general stress resistance, and analysis of protein synthesis.
Appl. Environ. Microbiol.
64:4238-4245[Abstract/Free Full Text].
|
| 17.
|
Helbling, E. W.,
E. R. Marguet,
V. E. Villafañe, and O. Holm-Hansen.
1995.
Bacterioplankton viability in Antarctic waters as affected by solar ultraviolet radiation.
Mar. Ecol. Prog. Ser.
126:293-298.
|
| 18.
|
Herndl, G. J.,
G. Müller-Niklas, and J. Frick.
1993.
Major role of ultraviolet-B in controlling bacterioplankton growth in the surface layer of the ocean.
Nature
361:717-719.
|
| 19.
|
Jagger, J.,
H. Tabeke, and J. Snow.
1970.
Photoreactivation and killing in Streptomyces: action spectra and kinetic studies.
Photochem. Photobiol.
12:185-196[Medline].
|
| 20.
|
Jeffrey, W. H.,
P. Aas,
M. M. Lyons,
R. B. Coffin,
R. J. Pledger, and D. L. Mitchell.
1996.
Ambiant solar radiation-induced photodamage in marine bacterioplankton.
Photochem. Photobiol.
64:419-427.
|
| 21.
|
Jeffrey, W. H.,
R. J. Pledger,
P. Aas,
S. Hager,
R. B. Coffin,
R. Von Haven, and D. L. Mitchell.
1996.
Diel and depth profiles of DNA photodamage in bacterioplankton exposed to ambiant solar ultraviolet radiation.
Mar. Ecol. Prog. Ser.
137:283-291.
|
| 22.
|
Kaiser, E., and G. J. Herndl.
1997.
Rapid recovery of marine bacterioplankton activity after inhibition by UV radiation in coastal waters.
Appl. Environ. Microbiol.
63:4026-4031[Abstract].
|
| 23.
|
Karentz, D.,
J. E. Cleaver, and D. L. Mitchell.
1991.
Cell survival characteristics and molecular responses of Antarctic phytoplankton to ultraviolet-B radiation.
J. Phycol.
27:326-341.
|
| 24.
|
Kato, R.,
K. Hasegawa,
Y. Hidaka,
S. Kuramitsu, and T. Hoshino.
1997.
Characterization of a thermostable DNA photolyase from an extremely thermophilic bacterium, Thermus thermophilus HB27.
J. Bacteriol.
179:6499-6503[Abstract/Free Full Text].
|
| 25.
|
Keller, L. C., and R. B. Maxcy.
1984.
Effect of physiological age on radiation resistance of some bacteria that are highly radiation resistant.
Appl. Environ. Microbiol.
47:915-918[Abstract/Free Full Text].
|
| 26.
|
Kirk, J. T. O.,
B. R. Hargreaves,
D. P. Morris,
R. B. Coffin,
B. David,
D. Frederickson,
D. Karentz,
D. R. S. Lean,
M. P. Lesser,
S. Madronich,
J. H. Morrow,
N. B. Nelson, and N. M. Scully.
1994.
Measurement of UV-B in two freshwater lakes: an instrument intercomparison.
Arch. Hydrobiol. Beih.
43:71-99.
|
| 27.
|
Lebaron, P.,
N. Parthuisot, and P. Catala.
1998.
Comparison of blue nucleic acid dyes for flow cytometric enumeration of bacteria in aquatic systems.
Appl. Environ. Microbiol.
64:1725-1730[Abstract/Free Full Text].
|
| 28.
|
Le Minor, L.
1984.
Genus Salmonella, p. 427-458.
In
N. R. Krieg, and J. G. Holt (ed.), Bergey's manual of systematic bacteriology, vol. 1. The Williams & Wilkins Co., Baltimore, Md.
|
| 29.
|
Li, S., and R. Waters.
1998.
Escherichia coli strains lacking protein HU are UV sensitive due to a role for HU in homologous recombination.
J. Bacteriol.
180:3750-3756[Abstract/Free Full Text].
|
| 30.
|
Li, Y. F., and A. Sancar.
1991.
Cloning, sequencing, expression and characterization of DNA photolyase from Salmonella typhimurium.
Nucleic Acids Res.
19:4885-4890[Abstract/Free Full Text].
|
| 31.
|
Liltved, H., and B. Landfald.
1996.
Influence of liquid holding recovery and photoreactivation on survival of ultraviolet-irradiated fish pathogenic bacteria.
Water Res.
30:1109-1114.
|
| 32.
|
Loewen, P. C.,
B. Hu,
J. Strutinsky, and R. Sparling.
1998.
Regulation in the rpoS regulon of Escherichia coli.
Can. J. Microbiol.
44:707-717[Medline].
|
| 33.
|
Lyons, M. M.,
P. Aas,
J. D. Pakulski,
L. Van Waasbergen,
D. L. Mitchell,
R. V. Miller, and W. H. Jeffrey.
1998.
Ultraviolet radiation induced DNA damage in coral reef microbial communities.
Mar. Biol.
130:537-543.
|
| 34.
|
Mitchell, D. L.
1996.
Radioimmunoassay of DNA damaged by ultraviolet light, p. 73-83.
In
G. Pfeiffer (ed.), Technologies for DNA damage and mutations. Plenum Publishing Corp., New York, N.Y.
|
| 35.
|
Mitchell, D. L., and D. Karentz.
1993.
The induction and repair of DNA photodamage in the environment, p. 345-377.
In
A. R. Young, L. O. Bjorn, J. Moan, and W. Nultsch (ed.), Environmental UV photobiology. Plenum, New York, N.Y.
|
| 36.
|
Müller-Niklas, G.,
A. Heissenberger,
S. Puskeric, and G. J. Herndl.
1995.
Ultraviolet-B radiation and bacterial metabolism in coastal waters.
Aquat. Microb. Ecol.
9:111-116.
|
| 37.
|
Nicholson, W. L.
1995.
Photoreactivation in the genus Bacillus.
Curr. Microbiol.
31:361-364[Medline].
|
| 38.
|
Nyström, T.,
R. M. Olsson, and S. Kjelleberg.
1992.
Survival, stress resistance, and alterations in protein expression in the marine Vibrio sp. strain S14 during starvation for different individual nutrients.
Appl. Environ. Microbiol.
58:55-65[Abstract/Free Full Text].
|
| 39.
|
Pakulski, J. D.,
P. Aas,
W. Jeffrey,
M. Lyons,
L. Von Waasenbergen,
D. Mitchell, and R. Coffin.
1998.
Influence of light on bacterioplankton production and respiration in a subtropical coral reef.
Aquat. Microb. Ecol.
14:137-148.
|
| 40.
|
Palleroni, N. J.
1984.
Genus Pseudomonas, p. 141-199.
In
N. R. Krieg, and J. G. Holt (ed.), Bergey's manual of systematic bacteriology, vol. 1. The Williams & Wilkins Co., Baltimore, Md.
|
| 41.
|
Paul, J. H.
1982.
The use of Hoechst dyes 33258 and 33342 for enumeration of attached and planktonic bacteria.
Appl. Environ. Microbiol.
43:939-944[Abstract/Free Full Text].
|
| 42.
|
Paul, J. H., and D. Carlson.
1984.
Genetic material in the marine environment: implication for bacterial DNA.
Limnol. Oceanogr.
29:1091-1097.
|
| 43.
|
Paul, J. H., and B. Myers.
1982.
Fluorimetric determination of DNA in aquatic microorganisms by use of Hoechst 33258.
Appl. Environ. Microbiol.
43:1393-1399[Abstract/Free Full Text].
|
| 44.
|
Regan, J. D.,
W. L. Carrier,
H. Gucinski,
B. L. Olla,
H. Yoshida,
R. K. Fujimura, and R. I. Wicklund.
1992.
DNA as a solar dosimeter in the ocean.
Photochem. Photobiol.
56:35-42[Medline].
|
| 45.
|
Sancar, A.
1994.
Structure and function of DNA photolyase.
Biochemistry
33:2-9[Medline].
|
| 46.
|
Schut, F.,
E. J. De Vries,
J. C. Gottschal,
B. R. Robertson,
W. Harder,
R. A. Prins, and D. K. Button.
1993.
Isolation of typical marine bacteria by dilution culture: growth, maintenance, and characteristics of isolates under laboratory conditions.
Appl. Environ. Microbiol.
59:2150-2160[Abstract/Free Full Text].
|
| 47.
|
Setlow, P.
1992.
I will survive: protecting and repairing spore DNA.
J. Bacteriol.
174:2737-2741[Free Full Text].
|
| 48.
|
Setlow, R. B.
1966.
Cyclobutane-type pyrimidin dimers in polynucleotides.
Science
153:379-386[Abstract/Free Full Text].
|
| 49.
|
Stewart, G. J.,
C. A. Carlson, and J. L. Ingraham.
1983.
Evidence for an active role of donor cells in natural transformation of Pseudomonas stutzeri.
J. Bacteriol.
156:30-35[Abstract/Free Full Text].
|
| 50.
|
Takeuchi, M.,
H. Sawada,
H. Oyaizu, and A. Yokota.
1994.
Phylogenetic evidence for Sphingomonas and Rhizomonas as nonphotosynthetic members of the alpha-4 subclass of the proteobacteria.
Int. J. Syst. Bacteriol.
44:308-314[Abstract/Free Full Text].
|
| 51.
|
Vincent, W. F., and A. Quesada.
1994.
Ultraviolet radiation effects on cyanobacteria: implications for Antarctic microbial ecosystems.
Antarct. Res. Ser.
62:111-124.
|
| 52.
|
Webb, R. B., and R. W. Tuveson.
1982.
Differential sensitivity to inactivation of nur and nur+ strains of Escherichia coli at six selected wavelengths in the UVA, UVB, and UVC ranges.
Photochem. Photobiol.
36:525-530[Medline].
|
| 53.
|
Wickman, S., and M. Carsten.
1998.
Effects of ultraviolet-B radiation on two Arctic microbial food webs.
Aquat. Microb. Ecol.
16:163-171.
|
| 54.
|
Williamson, C. E.,
H. E. Zagarese,
P. C. Schule,
B. R. Hargreaves, and J. Seva.
1994.
The impact of short-term exposure to UV-B radiation on zooplankton communities.
Limnol. Oceanogr.
41:1024-1034.
|
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-
Eguchi, M., Ostrowski, M., Fegatella, F., Bowman, J., Nichols, D., Nishino, T., Cavicchioli, R.
(2001). Sphingomonas alaskensis Strain AFO1, an Abundant Oligotrophic Ultramicrobacterium from the North Pacific. Appl. Environ. Microbiol.
67: 4945-4954
[Abstract]
[Full Text]
-
Kim, J. J., Sundin, G. W.
(2001). Construction and Analysis of Photolyase Mutants of Pseudomonas aeruginosa and Pseudomonas syringae: Contribution of Photoreactivation, Nucleotide Excision Repair, and Mutagenic DNA Repair to Cell Survival and Mutability following Exposure to UV-B Radiation. Appl. Environ. Microbiol.
67: 1405-1411
[Abstract]
[Full Text]
-
Ostrowski, M., Cavicchioli, R., Blaauw, M., Gottschal, J. C.
(2001). Specific Growth Rate Plays a Critical Role in Hydrogen Peroxide Resistance of the Marine Oligotrophic Ultramicrobacterium Sphingomonas alaskensis Strain RB2256. Appl. Environ. Microbiol.
67: 1292-1299
[Abstract]
[Full Text]
-
Winter, C., Moeseneder, M. M., Herndl, G. J.
(2001). Impact of UV Radiation on Bacterioplankton Community Composition. Appl. Environ. Microbiol.
67: 665-672
[Abstract]
[Full Text]
-
Arrieta, J. M., Weinbauer, M. G., Herndl, G. J.
(2000). Interspecific Variability in Sensitivity to UV Radiation and Subsequent Recovery in Selected Isolates of Marine Bacteria. Appl. Environ. Microbiol.
66: 1468-1473
[Abstract]
[Full Text]
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Abstract
Introduction
