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Journal of Bacteriology, June 2000, p. 3017-3021, Vol. 182, No. 11
0021-9193/00/$04.00+0
Copyright © 2000, American Society for Microbiology. All rights reserved.
Color-Sensitive Motility and Methanol Release
Responses in Rhodobacter sphaeroides
Remco
Kort,1,
Wim
Crielaard,1
John L.
Spudich,2 and
Klaas J.
Hellingwerf1,*
Laboratory for Microbiology, E. C. Slater Institute, University of Amsterdam, 1018 WS Amsterdam, The
Netherlands,1 and Department of
Microbiology and Molecular Genetics, The University of Texas Medical
School, Houston, Texas 770302
Received 26 October 1999/Accepted 2 March 2000
 |
ABSTRACT |
Blue-light-induced repellent and demethylation responses,
characteristic of behavioral adaptation, were observed in
Rhodobacter sphaeroides. They were analyzed by
computer-assisted motion analysis and through the release of volatile
tritiated compounds from
[methyl-3H]methionine-labeled cells,
respectively. Increases in the stop frequency and the rate of methanol
release were induced by exposure of cells to repellent light signals,
such as an increase in blue- and a decrease in infrared-light
intensity. At a 
of >500 nm the amplitude of the methanol release
response followed the absorbance spectrum of the photosynthetic
pigments, suggesting that they function as photosensors for this
response. In contrast to the previously reported motility response to a
decrease in infrared light, the blue-light response reported here does
not depend on the number of photosynthetic pigments per cell,
suggesting that it is mediated by a separate sensor. Therefore, color
discrimination in taxis responses in R. sphaeroides
involves two photosensing systems: the photosynthetic pigments and an
additional photosensor, responding to blue light. The signal generated
by the former system could result in the migration of cells to a light
climate beneficial for photosynthesis, while the blue-light system
could allow cells to avoid too-high intensities of (harmful) blue light.
| |
INTRODUCTION |
Changes in the environmental light
climate induce behavioral responses in photosynthetic purple bacteria,
which allow them to migrate towards an environment beneficial for
photosynthesis. Reports on photosynthetic bacteria that accumulate in
infrared light are among the classics of microbiology (2,
3). In more recent years, more-complex behavioral responses of
these organisms towards light have been reported. Free-swimming
Ectothiorhodospira halophila cells, in addition, show a
repellent response, with a maximal relative increase in the flagellar
reversal frequency at 450 nm (20). A colony of
Rhodospirillum centenum is repelled by intense green light
(550 to 600 nm) and is attracted by red or infrared light that is
absorbed by the photosynthetic pigments (15). Light-induced
motility responses have also been analyzed in Rhodobacter
sphaeroides WS8-N (1). This bacterium responds to a
step-down in yellow-green light (530 to 600 nm), and infrared light in
a background of red monitoring light (650 ± 10 nm), by an
increase in its stop or reorientation frequency, with adaptation taking
approximately 2 min. The photosynthetic apparatus is the photosensor
for this response (6).
Stimulus-induced turnover of methylated carboxyl groups of
methyl-accepting transducer proteins is a common feature of prokaryotic taxis-signaling pathways. Nevertheless, previous methanol release assays with intact R. sphaeroides cells have not revealed
changes in methanol release upon the addition or removal of
chemoeffectors (19). The recent finding of chemotaxis
operons encoding MCP homologues, methyltransferases and -esterases,
however, supports the involvement of a methylation-dependent adaptation
pathway for tactic responses in this bacterium (7). This
study describes a blue-light-induced taxis response in R. sphaeroides RK1 that is distinct from the response to
photosynthetic light previously reported (6).
Color-sensitive, light-induced changes in the release of volatile
[3H]methanol from cells labeled with radioactive
methionine provide evidence for the involvement of methyl-accepting
transducers in signaling by both phototaxis systems.
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MATERIALS AND METHODS |
Strain and culture conditions.
R. sphaeroides strain
RK1 (11) was cultured at 30°C under anaerobic conditions
in white light (15 W · m
2), in 20-ml screw-cap
tubes containing Sistrom's minimal medium A supplemented with
succinate as the carbon source (18). Culturing under
semianaerobic conditions in the dark was carried out in 80%-filled
300-ml Erlenmeyer flasks at a low rotation speed on an orbital shaker
(25 rpm), and culturing under aerobic conditions in the dark was
carried out in 4%-filled 500-ml Erlenmeyer flasks at a high rotation
speed (250 rpm). Cells grown at a high light intensity were illuminated
with white light at 100 W · m2, while for low
light intensities 3 W · m2 was used.
Single-cell motion analysis.
R. sphaeroides cells were
cultured under anaerobic conditions in the light and harvested for
analyses at an optical density at 660 nm (OD660) of ~0.8.
The setup used in this study has been described elsewhere (8,
24). Briefly, cells were monitored under a cover glass by
dark-field microscopy with a 150-W tungsten-halogen lamp (Ushio Inc.)
and a 600-nm long-pass filter (light intensity, 8.3 W · m2, as determined with a Kettering radiant power meter
[Scientific Instruments]). Blue-light repellent stimuli (using pulses
of 3 s) were delivered via an HBO 103W/2 mercury short-arc lamp
(Osram), through 400-, 450-, or 500-nm broad-band interference filters (FWHM, ± 20 nm). The corrected light intensities (i.e., corrected for
the surface of the light spot relative to the surface of the light
intensity meter) were 11, 83, and 51 W · m2 for
the three wavelengths, respectively. The motion analysis software was
run on a SPARC IPC workstation. The average linear speed of cells in a
suspension was obtained by combining two data sets in which the paths
of single motile cells were tracked for a period of 4 s. The first
data set was obtained starting at 1 s before the blue-light pulse
and continuing for the 3 s of the pulse, and the second data set
was obtained for the last 1 s of the pulse and the first 3 s
after the pulse; for both, the frame rate was 15 frames/s (i.e., 67 ms/frame). The following settings were used for the calculation of the
centroids: neighbor width/height, 2/2; minimum number of pixels, 1;
maximum number of pixels, 4,096. The settings for the calculation of
the paths were as follows: search mask size, 15; minimum path duration,
40; average minimum movement, 1. All calculated paths were visually
inspected, and paths of nonmotile cells were removed with the path
editor. About 150 paths, obtained from 10 independent recordings, were
merged into a single file and used for calculation of the average
linear speed of the cells.
Methanol release assay.
The flow assay for measurements of
photostimulus-induced changes in carboxylmethylesterase activity, based
on a procedure developed for Escherichia coli chemotaxis
(9) and modified for Halobacterium salinarum
(14), was used with 2 ml of cell suspension
(OD660, ~0.8) for each experiment. Cells were cultured under anaerobic conditions in the light (unless specified otherwise) in
Sistrom medium plus 0.1 mM methionine in order to enhance the uptake of
methionine. They were washed three times in Sistrom medium without
methionine and were then incubated under semianaerobic or aerobic
conditions for 40 min in the presence of 200 µl of L-[methyl-3H]methionine (specific
activity, 75 Ci/mmol; concentration, 1 mCi/ml; [methionine] = 1.3 µM) (DuPont). Subsequently, cells were incubated on a
0.2-µm-pore-size syringe filter (Nalgene), and label was chased from
the cells with Sistrom medium containing 0.1 mM methionine during a
10-min period (flow rate, 1 ml · min1) at 28°C.
Next, 0.5-ml fractions were collected in 1.5-ml Eppendorf tubes for 10 min, and a light stimulus obtained from a tungsten or xenon lamp, in
combination with heat filters and broad-band interference filters, was
applied. All light intensities were adjusted to 20 W · m2, except for light of 400 nm, which was used at 5.0 W · m2. The amplitudes for the sustained responses
(interference filters of 500 nm and longer) in Fig. 5 were calculated
as the difference between the average number of counts per minute over
the three fractions collected before the light pulse (n = 3) and the average over the three fractions collected 2 min after
the start of the light pulse (n = 3). The amplitudes
for the transient responses (400- and 450-nm interference filters) in
Fig. 5 were calculated as the difference between the average number of
counts per minute over the three fractions collected before the light
pulse and the three fractions collected 2 min after the start of the
light pulse (n = 6) and the number of counts per minute
of the first fraction collected after the light pulse. The duration of
the light pulses varied from 8 to 10 min. The photostimuli had a
negligible effect on the temperature at the position of the immobilized
cells in the filter. Eppendorf tubes containing the collected samples were transferred to vials with scintillation fluid, incubated overnight
at room temperature in the dark to allow the transfer of volatile,
labeled methanol, and analyzed by liquid scintillation spectrometry.
| |
RESULTS |
Characterization of the motility response.
A step-up in blue
light in a background of infrared light caused a motility response in
swimming R. sphaeroides cells cultured under anaerobic
conditions in the light (a single-cell track is shown in Fig.
1). The average linear speed of 150 cell
paths was determined (Fig. 2). The cells
stopped and reoriented after a delay of 0.27 s; detailed
inspection of the recorded response (15 frames per s) revealed the
start of the blue-light pulse at 1.07 s (frame 16) and a decrease
in the average speed after 1.33 s (frame 20). During this delay
the speed of individual cells increased slightly, which may be due to
the increased light intensity, causing a higher proton motive force.
The latter will affect the cellular swimming speed (a phenomenon called
photokinesis). After 1.27 s, the cells reached their lowest
average speed. This is mainly caused by the cells stopping rather than
by a decrease in swimming speed. After the pulse (duration, 3 s),
the cells started to swim again with a delay of 0.07 s. The cells
recovered to their prestimulus swimming speed in about 1 s.
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FIG. 1.
Representative single-cell track of R. sphaeroides RK1. The track starts at the upper right corner. The
arrow indicates the start of the step-up in blue-light intensity. The
time interval between two recorded frames (dots) is 67 ms, and infrared
light was used as the monitoring light. For further details, see
Materials and Methods.
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FIG. 2.
Time dependence of the effect of blue light on the
average speed of free-swimming R. sphaeroides RK1 cells.
After a delay of 1 s, cells received a step-up in blue light for
3 s. The tracks of 150 individual cells were averaged for this
figure. For further details, see Materials and Methods.
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Full adaptation to the prestimulus stopping frequency was not observed,
even when the duration of the continuous blue-light
illumination was
extended up to several minutes. Cells either
stopped completely or
continued swimming with an increased stop
frequency during the entire
blue-light exposure. Replacement of
the 450-nm interference filter by a
500-nm filter abolished the
stop response at the light intensity used.
On the other hand,
a step-up in light of 400 nm or in white light
(i.e., without
the use of any interference filter) did result in a stop
response,
which was similar to the response observed when the 450-nm
filter
was used (data not shown). Decreasing the 450-nm pulse duration
from 3 to 1 s did not significantly affect the amplitude of the
response (for the calculation of this amplitude, see Materials
and
Methods). However, a decrease in the length of the pulse to
100 ms
resulted in a reduction of the response to approximately
20%.
Exposure of anaerobically cultured
R. sphaeroides RK1 cells
to oxygen also resulted in an increased stop frequency, as observed
previously for
R. sphaeroides WS8-N (
5). No
effect of blue
light on the motility of cells was observed during their
response
to oxygen; cells need to be cultured and observed under
anaerobic
conditions in order for this response to be observed. Cells
grown
anaerobically at high (100 W · m
2) and low
(3 W · m
2) light intensities did not show
significant differences in the
amplitude of the blue-light motility
response. In addition to
the response to a step-up in blue light, the
R. sphaeroides RK1
cells exhibited a similar motility
response (increased reorientation
frequency) to a step-down in infrared
light (
> 700 nm), which
was earlier described for
R. sphaeroides WS8-N (
6). After periods
of 5 to 15 min of
frequent exposure to blue-light pulses of 3
s, only part of the
cell population continued to respond to such
pulses. Eventually the
cells completely lost this motility response,
while their nonstimulated
motility, and their response to a decrease
in infrared light, was not
affected.
Methanol release.
Light-induced release of volatile tritiated
compounds was analyzed using a flow assay, with intact R. sphaeroides cells placed on a filter. With anaerobically grown
cells, a transient increase in methanol production was observed upon a
step-up in blue-light intensity (Fig.
3A); this is similar to that observed
from a methylation-dependent adaptation reaction of a repellent
response in E. coli. When grown under semianaerobic
conditions, R. sphaeroides cells also showed an increased
release of methanol (data not shown), while with aerobically grown
cells, this increase was not detectable (Fig. 3B). The results in Fig.
3 match the occurrence of the reported blue-light motility response,
which also was observed in cells grown anaerobically in the light and
was less pronounced in cells grown under semianaerobic conditions in
the dark.
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FIG. 3.
Effect of growth conditions on the release of
3H-labeled methanol by R. sphaeroides RK1 cells
upon a step-up in blue light: The arrow indicates the time of provision
of the step-up in blue light (450 nm). Cells were grown anaerobically
in the light (A) or aerobically in the dark (B). Each point indicates
the amount of radioactive methanol collected during a 30-s period. For
further details, see Materials and Methods.
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The effect of infrared light on methanol release was also investigated.
R. sphaeroides cells do not display a pronounced phototactic
response to an increase in infrared light, but a decrease in this
light
causes a transient stop, followed by adaptation (
6).
Interestingly, this symmetry (the removal of an attractant leads
to a
response very similar to that resulting from the addition
of a
repellent) is partially reflected in light-induced methanol
release, as
indicated in Fig.
4. An attractant
stimulus leads
to a sustained decrease in the release of methanol,
while a repellent
stimulus leads to a transient response (Fig.
4).
After the initiation
of the light stimulus, it typically takes about 4 data points,
i.e., 2 min, before the rate of methanol release has
reached a
new level (attractant response) or has returned to the
prestimulus
level (repellent response).
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FIG. 4.
Analysis of 3H-labeled methanol release in
R. sphaeroides RK1 cells upon step-up stimuli with infrared
(IR) and blue light. Infrared light was selected with a long-pass
filter (50% transmission at 758 nm). For blue light, a broad-band
450-nm filter was used.
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The observed opposite effects of blue and infrared photostimuli on
methanol release in intact
R. sphaeroides cells impelled
us
to investigate the wavelength dependence of this response.
Accordingly,
we used a set of broad-band interference filters
and a 600-nm long-pass
filter (see Materials and Methods). The
data in Fig.
5 show that the increase in
[
3H]methanol release has a wavelength dependence that
follows the
absorbance spectrum of the photosynthetic pigments for
wavelengths
of 500 nm and above. Thus, the photosynthetic pigments
presumably
are the photoreceptors for the methanol release response to
a
decrease in the intensity of light sustaining photosynthesis.
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FIG. 5.
Wavelength dependence of the light-induced release of
[3H]methanol in R. sphaeroides RK1 cells in
response to a step-up in light stimuli. The light intensity used at 400 nm was 4 times lower than the intensity used for all the other
wavelengths. For comparison, an absorption spectrum of anaerobically
grown R. sphaeroides RK1 cells is shown. Error bars,
calculated standard deviations. For further details, see Materials and
Methods.
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DISCUSSION |
The blue-light response.
When swimming R. sphaeroides cells are exposed to blue light, they stop and then
partially adapt to their prestimulus motility. Does this response lead
to migration away from blue light? For comparison, consider the very
similar motility response to a decrease in photosynthetic light in
R. sphaeroides WS8-N (which we also observe in cells of
strain RK1). Under anaerobic conditions both strains respond to a
step-down of photosynthetic light by a transient stop. It has been
reported, however, that WS8-N cells, exposed to a white-light beam for
a few minutes, accumulate outside that beam in the dark
(17). For a phototrophic bacterium, this is not easily
interpreted as a physiological response. Possibly, in nature R. sphaeroides does not often face such steep light intensity
gradients, and the accumulation in the dark can be explained as an
"overreaction" of the cells. Overly steep gradients may cause
nonphysiological complete stops, leading to the unexpected accumulation
pattern. However, the lack of photomotility responses in a
cheAII mutant, identified via selection of cells which do not sense a light-dark boundary, does show the importance of these responses for R. sphaeroides for the avoidance of a dark
environment (7).
The relatively slow adaptation of
R. sphaeroides to bright
photostimuli (adaptation takes approximately 2 min for the response
to
a step-down in photosynthetic light [
6], although with
subsaturating
light intensities, cells can show adaptation within
seconds [
16])
complicates the observation of
adaptation. With the procedure
used here, free-swimming cells are not
easily tracked for periods
longer than several seconds, because
gradually cells leave the
light spot. When the blue light was turned
off, even after minutes
of exposure, cells exhibited a decrease in stop
frequency, indicating
that they were not fully adapted. This may be due
to the large
change in light intensity; more-subtle changes may result
in full
adaptation within shorter
times.
As blue light is also used for photosynthesis,
R. sphaeroides RK1 also shows an increase in stop frequency in
response to
a decrease in blue light (and other wavelengths of
photosynthetic
light), while, as reported here, it also shows an
increase in
stop frequency in response high blue-light intensities in a
background
of infrared light. These two responses may bias the swimming
pattern
of
R. sphaeroides RK1 toward its most favorable
light climate
for photosynthesis, while simultaneously allowing it to
avoid
radiation damage. This pattern of opposite responses, as a
function
of either the color or the intensity of the actinic
illumination,
is often seen in phototrophic prokaryotes and algae
(
4,
8,
13,
15,
21).
The response to a decrease in infrared light is strongly dependent on
the capacity of the photosynthetic light-harvesting
apparatus. Cells
grown at high light intensities respond to a
much wider range of
step-down intensities than cells grown at
low light intensities,
evidently because photosynthesis saturates
at much lower intensities in
low-light-grown cells (
6). Similar
saturation is not
observed for the behavioral response to a step-up
in blue light, which
argues against a role for the photosynthetic
pigments as the underlying
photoreceptors in this latter response.
This response is observable
only at relatively high light intensities,
i.e., when the length of the
light pulse is longer than 100 ms
and the intensity does not decrease
below 30% of the default value
(see Materials and Methods). The
blue-light response cannot be
observed in cells exposed to oxygen,
which itself increases the
reorientation frequency. This is in line
with the idea that multiple
signals feed into the taxis transduction
pathway in these cells.
This is further supported by the phenotype of
an
R. sphaeroides RK1 mutant in which the chemotaxis operon
II was interrupted.
This mutant has lost all light-induced behavioral
responses, while
its motility is unimpaired (unpublished
data).
The methanol release assay.
The nature of the labeled volatile
compound (i.e., methanol) has not been independently confirmed in this
study, but methanol is the only compound detected so far in this type
of experiment carried out with eubacteria. In a previous study,
methanol release in R. sphaeroides WS8 was not observed in
response to the addition or removal of a combination of the attractants
serine and succinate (19). This may have been due to the
lower specific activity and amount of label added in the latter study.
In
E. coli the transient increase in the rate of methanol
release occurs during the process of adaptation of the taxis response,
as a result of increased methylesterase activity of phosphorylated
CheB. A similar mechanism presumably is operative in
R. sphaeroides,
in which one
cheB and two
cheR
genes have been identified recently
(
7). The methanol
release responses in
R. sphaeroides are reminiscent
of those
observed in
E. coli. Attractant stimuli decrease
methylesterase
activity, which then recovers in a period of minutes to
the prestimulus
level. A repellent stimulus produces a transient
increase in methylesterase
activity. The sustained decrease, and
transient increase, in methanol
release shown in Fig.
4 can be
rationalized by the assumption
that phosphorylated CheB has a
relatively short lifetime, which
causes a transient response
(
12). Inhibition of CheA kinase
activity, which increases
the number of nonphosphorylated CheB
molecules, has a more sustained
effect.
The photosensors mediating the light responses.
The
photosynthetic apparatus presumably mediates the motility and methanol
release responses to a step-down in photosynthetic light. Furthermore,
adaptation to a decrease in infrared light involves a
methylation-dependent system. A methyl-accepting protein capable of
sensing the membrane potential could form part of the machinery for
this signal transfer route.
The molecular identity of the photoreceptor mediating the blue-light
response in
R. sphaeroides remains to be resolved. Sensory
rhodopsins (
8) have not been detected in purple bacteria. In
E. coli, intermediates of the heme biosynthesis pathway act
as
sensors for a blue-light tumbling response (
22,
23).
However,
both the wavelength dependence and the oxygen requirement of
the
latter response make it incompatible with the blue-light-induced
behavioral response in
R. sphaeroides reported
here.
Recently, the
pyp gene, encoding a photoactive yellow
protein (PYP), has been identified in
R. sphaeroides RK1
(
11), while
it was not detectable in the WS8-N strain
(
10). Although the
blue-light motility response is present
in the RK1 strain and
not in WS8-N,
pyp deletion mutants of
R. sphaeroides RK1 displayed
unimpaired blue-light
responses. This excludes the possibility
that the PYP from
Rhodobacter is the photosensor for the blue-light
repellent
response in this organism (
10).
| |
ACKNOWLEDGMENTS |
R.K. thanks Xue-Nong Zhang, Kwang-Hwan Jung, and Bastianella
Perazzona for expert assistance with the motion analyses and flow
assays and Elena Spudich for fruitful discussions. Judy Armitage is
acknowledged for making plasmid pDS1 available for mutagenesis experiments in Rhodobacter.
We acknowledge the support of collaborative research grant 960237 from
NATO to J.L.S. and SIR travel grant 14-1779 to Houston, Tex., from the
Dutch Organization of Scientific Research (NWO) to R.K.
| |
FOOTNOTES |
*
Corresponding author. Mailing address: Laboratory for
Microbiology, E. C. Slater Institute, University of Amsterdam,
Nieuwe Achtergracht 127, 1018 WS Amsterdam, The Netherlands. Phone:
31-20-5257055. Fax: 31-20-5257056. E-mail:
K.Hellingwerf{at}chem.uva.nl.
Present address: European Synchrotron Radiation Facility, 38043 Grenoble, France.
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Journal of Bacteriology, June 2000, p. 3017-3021, Vol. 182, No. 11
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Abstract
Introduction
