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INTRODUCTION |
Bacillus subtilis is a
gram-positive bacterium that undergoes sporulation when deprived of an
essential nutrient or nutrients. The resulting elliptical spore is
metabolically dormant and resistant to a variety of harsh environmental
conditions. However, in the presence of specific germinant molecules,
such as L-alanine, the spore can resume metabolic activity
and eventually return to vegetative growth through the processes of
spore germination, which does not require macromolecular synthesis, and
spore outgrowth, which begins with the resumption of macromolecular
synthesis (35). An essential event during spore outgrowth is
the synthesis of new cell wall peptidoglycan (PG). Much of the spore's
PG is degraded during the first minutes of germination (8).
However, the spore PG which is not degraded comprises the germ cell
wall, which is thought to provide the template for PG synthesis during
spore outgrowth (3). The PG generated during the latter
period then generates the rod-shaped vegetative cell.
One group of enzymes essential for PG synthesis is the
penicillin-binding proteins (PBPs), which catalyze PG strand elongation (transglycosylase activity) and regulate PG side chain cross-linking through transpeptidase and D,D-peptidase
activities (reviewed in reference 9). A number of
PBPs are present in growing B. subtilis cells, and some are
in the dormant spore; the initiation of transcription of a number of
pbp genes also occurs very early in spore outgrowth
(17, 18, 20, 30). However, the specific function of most
individual PBPs in vegetative growth and spore outgrowth has been
difficult to discern, as mutations in one or more pbp genes
often result in few phenotypic effects, probably because many of these
proteins perform redundant functions (31).
There are, however, several PBPs whose loss results in a clear
phenotype (5, 18, 30, 37). One such PBP is the class B
high-molecular-weight (HMW) PBP2a encoded by pbpA, as the
loss of this putative monofunctional transpeptidase results in both delayed spore outgrowth and an increased cell diameter early in spore
outgrowth (18). In contrast, the loss of PBP2a does not cause phenotypic changes during vegetative growth (18). In
this report we further characterize the outgrowth of spores lacking PBP2a and examine PG metabolism during this period. The importance of
other HMW PBPs during spore outgrowth was also studied by constructing strains lacking PBP2a and additional HMW PBPs and examining the properties of these strains during spore outgrowth, as well as during
vegetative growth and sporulation.
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MATERIALS AND METHODS |
Bacterial strains, growth, sporulation, and spore germination and
outgrowth.
The B. subtilis strains used are shown in
Table 1; all strains are derived from
PS832, a derivative of strain 168. B. subtilis was
transformed with either chromosomal DNA or plasmid DNA as previously
described (1), and transformants were selected on 2× SG
plates (14) by their resistance to appropriate antibiotics: chloramphenicol (Cmr) (3.0 µg/ml); the macrolides
(MLSr) (lincomycin, 12.5 µg/ml; erythromycin, 0.5 µg/ml); or spectinomycin (Spr) (100 µg/ml). B. subtilis was routinely grown and sporulated at 37°C in
antibiotic-free 2× SG medium (14). The spores were then
purified by repeated washing in water as described previously (21). Purified spores were germinated at 37°C in a number
of different media, including 2× YT (31), Penassay broth
(PAB) (Difco), and Luria broth (LB) (containing [per liter] 10 g
of tryptone, 5 g of yeast extract, and 10 g of NaCl), after a
30-min heat shock in water at 70°C as described previously
(21); all germination media were supplemented with
L-alanine (4 mM) to stimulate the initiation of spore
germination. To remove molecules released during the initial stages of
germination, all spores outgrowing in PAB medium were harvested by
centrifugation at 3,000 × g for 5 min at 24°C 30 min
after the initiation of spore germination and the pellet was
resuspended in an equal volume of prewarmed (37°C) fresh medium. The
optical density at 600 nm (OD600) of all vegetative cells
and germinating and outgrowing spores was monitored with a Genesys 5 spectrophotometer.
Membrane isolation, penicillin-binding assays, spore heat
resistance, and analysis of PG structure.
Membranes were isolated
from B. subtilis cells in log-phase growth in 2× SG medium
at an OD600 of approximately 0.5 or from spores outgrowing
in 2× YT medium 2 h after the initiation of spore germination, as
previously described (31). The purified membranes were
incubated with fluorescein-hexanoic acid 6-aminopenicillanic acid
(FLU-C6-APA), the proteins were separated by sodium dodecyl sulfate-10% polyacrylamide gel electrophoresis, and labeled PBPs were
visualized with a FluorimagerSI (Vistra), as described previously (31).
Spore wet-heat resistance was measured by incubating purified spores in
water at 85°C for 15 min and plating serial dilutions of unheated and
heated spores on LB plates (27). Spore cortex PG was
isolated, hydrolyzed, reduced, and analyzed by reverse-phase high-performance liquid chromatography (HPLC) as previously described (2, 25). PG from outgrowing spores of various strains was isolated 30, 60, and 90 min after the initiation of spore germination and analyzed in a similar manner.
Light microscopy and viability assays.
Outgrowing spores
(0.5 ml) were fixed in 2.5% glutaraldehyde for 20 min at 22°C,
washed in 0.5 ml of phosphate-buffered saline (PBS; 137 mM NaCl, 3 mM
KCl, 5.4 mM Na2HPO4, 1.7 mM
KH2PO4 [pH 7.3]), placed on 0.01%
polylysine-coated coverslips, and visualized by differential
interference contrast microscopy with a Noran confocal laser scanning
microscope employing a 100× Plan-APO chromatic oil immersion lens
(Zeiss) as described previously (31). To visualize the cell
walls and DNA of outgrowing spores, the fixed cells on coverslips were
treated with lysozyme (1 mg/ml) at 24°C for 30 s, rinsed several
times in PBS, and incubated with 2% bovine serum albumin in PBS for
1 h, and wheat germ agglutinin conjugated to Oregon green
(Molecular Probes) was added along with 4',6-diamidino-2-phenylindole (DAPI) to final concentrations of 5 and 1.25 µg/ml, respectively (22). After a 2-h incubation, the cells were rinsed eight
times with PBS, placed on slides by using the SlowFade light antifade kit (Molecular Probes), and viewed at the appropriate wavelength (22).
Spore viability was assessed by plating purified spores on either LB
plates or LB plates supplemented with 10 mM MgCl2. The plates were incubated overnight at 37°C, and the colonies were counted. The LIVE/DEAD BacLight bacterial viability kit (Molecular Probes) was also used as directed by the manufacturer to assess viability during spore outgrowth in 2× YT medium. Samples (0.5 ml) of
outgrowing spores were harvested at various times after the initiation
of spore germination and incubated with 1.5 µl of LIVE/DEAD stain for
2 to 3 min. An aliquot of the suspension was placed on a 1%
agarose-coated slide, a coverslip was gently applied, and the cell
fields were examined with the fluorescein filter on an Olympus B-max
microscope employing a 40× Dplan-APO lens. The live cells were
counted, and then the rhodamine filter was used to assay the number of
dead cells in the same field. At each time point, at least 300 cells
were examined.
Measurement of macromolecular synthesis during spore
outgrowth.
To assess PG synthesis during spore outgrowth in a
medium where cells remained viable and yet morphological changes were
evident, 12.5 ml of prewarmed (37°C) 2× YT medium containing 4 mM
L-alanine, 100 µM
N-acetyl-D-glucosamine, and 5 µCi of
[3H]N-acetyl-D-glucosamine (6.6 Ci/mmol) were inoculated with heat-shocked dormant spores to an initial
OD600 of approximately 0.5. Duplicate 0.5-ml samples were
harvested throughout subsequent spore germination and outgrowth, added
to 0.5 ml of 10% trichloroacetic acid (TCA) at 4°C, and incubated on
ice for at least 1 h, and the TCA precipitate was harvested by
filtration through GF/C membranes (Whatman) (10, 24). The
membranes were washed five times with 5 ml of 5% TCA and four times
with 5 ml of 95% ethanol and dried under a heat lamp; 3.5 ml of
Biosafe II counting fluid was added, and the samples were counted for 5 min with a Delta 300 scintillation counter (Searle). To determine the
extent to which PG synthesis required new protein synthesis, spore
germination and outgrowth was carried out as described above but with a
final total N-acetyl-D-glucosamine concentration
of 10 µM to improve the sensitivity of the assay and with the
addition of chloramphenicol (100 µg/ml) (20). In all of
these experiments, the percentage of
[3H]N-acetyl-D-glucosamine
incorporated into the TCA-insoluble fraction was determined and divided
by the initial OD of the spore culture to correct for differences in
the sizes of the initial inocula.
To confirm that the incorporation of
[3H]N-acetyl-D-glucosamine into a
TCA-insoluble form did represent PG synthesis, we used the method
previously described (24). Samples were harvested as
described above but were boiled for 10 min in 5% TCA to denature proteases which might interfere with subsequent reactions. Then the TCA
was removed by centrifugation (10,000 × g; 24°C) and
the pellet was washed with 1.0 M Tris HCl (pH 8.0) and dissolved in 1 ml of 20 mM Tris-HCl (pH 8.0) for digestion with either lysozyme (0.5 mg/ml) or trypsin (0.1 mg/ml, with 10 mM CaCl2). After
incubation overnight at 37°C, the digests were centrifuged for
several minutes at 10,000 × g and aliquots of both the
pellet and supernatant fraction were counted as described above. More
than 90% of the counts were solubilized by lysozyme, while less than
10% of the counts were solubilized by trypsin treatment (data not
shown). Consequently, the great majority of
[3H]N-acetyl-D-glucosamine
incorporated into TCA-insoluble form was in cell wall material.
Previous work has demonstrated that more than 90% of radiolabeled
N-acetyl-D-glucosamine incorporated into
TCA-insoluble material in growing B. subtilis cells is
incorporated into the cell wall, with 70 to 75% incorporated into the
PG fraction and the remainder into glucosamine-containing teichoic
acids (7, 24).
In order to measure PG turnover during outgrowth of wild-type and
pbpA spores, 12.5 ml of prewarmed (37°C) 2× YT medium
(containing 4 mM L-alanine) and 5 µCi of
[3H]N-acetyl-D-glucosamine (6.6 Ci/mmol) was inoculated with heat-shocked spores to an
OD600 of 0.5. Thirty minutes after the initiation of spore
germination, the outgrowing spores were harvested by centrifugation and
resuspended in 12.5 ml of prewarmed (37°C) 2× YT medium containing 1 mM unlabeled N-acetyl-D-glucosamine, and
TCA-insoluble radioactivity was measured during subsequent incubation
as described above (10). PG turnover during vegetative growth was assayed by adding 5 µCi of
[3H]N-acetyl-D-glucosamine (6.6 Ci/mmol) to log-phase wild-type B. subtilis
(OD600, 0.5) in 12.5 ml of 2× YT medium, incubating the
cells for 10 min, harvesting the cells by centrifugation, resuspending
the cell pellet in fresh 2× YT medium containing 1 mM unlabeled
N-acetyl-D-glucosamine, and analyzing aliquots for TCA-precipitable radioactivity as described above.
DNA synthesis during spore germination and outgrowth was measured in
12.5-ml cultures of 2× YT medium containing 6.25 µCi of
[3H-methyl]thymidine (6.7 Ci/mmol), 10 µM unlabeled
thymidine, and 4 mM 2-deoxyadenosine (32). Protein synthesis
was measured similarly but by the addition of 12.5 µCi of
L-[3,4,5-3H(N)]leucine (179 Ci/mmol) to 12.5 ml of 2× YT medium without additional unlabeled leucine
(32). In both cases TCA-precipitable radioactivity was
measured as described above.
Electron microscopy and autoradiography.
The location of
newly synthesized cell wall components was examined during spore
outgrowth by inoculating 5 ml of prewarmed (37°C) 2× YT medium with
heat-shocked wild-type or pbpA spores to an
OD600 of 1.5. After approximately 60 min, 500 µCi of
[3H]N-acetyl-D-glucosamine (6.9 Ci/mmol) was added to each culture, and 2 min later 250 µg of cold
N-acetyl-D-glucosamine was added. After a 10-min
chase (72 min after the initiation of spore germination), the samples
were harvested by centrifugation for 5 min (7,500 × g), fixed for 20 min in 1 ml of 2.5% glutaraldehyde in 0.1 M sodium cacodylate buffer (pH 7.4), washed in 0.1 M sodium cacodylate buffer, and embedded in low-melting-temperature agarose. The agarose pellets were postfixed in 1% osmium tetroxide-0.8% potassium
ferricyanide in 0.1 M cacodylate buffer, stained with 0.5% aqueous
uranyl acetate, dehydrated in graded ethanol solutions, and embedded in
epoxy resin (34).
Thin sections were cut with a diamond knife, placed on celloidin-coated
glass slides, and stained with uranyl acetate and lead citrate, and
then the slides were carbon coated in a vacuum evaporator, dipped in
Ilford L4 emulsion diluted 1:3.5 with water, dried, and stored in
lighttight boxes at 4°C for 20 weeks (11). The
autoradiographs were then developed in Agfa-Gevaert solution physical
developer (12) and fixed in 24% sodium thiosulfate, and
sections collected on copper specimen grids were viewed with a Philips
CM10 transmission electron microscope.
To determine if the cell wall was the source of most of the developed
silver grains in the electron microscopic autoradiographs described
above, the shortest distance from the center of each grain to the
center of the nearest cell wall was measured with a 7× magnifying
eyepiece. Where several grains were clustered in a circle equivalent to
the average diameter of an Ilford L4 silver bromide crystal (140 nm
[12]), the center of the cluster was considered the
center of the grain. To confirm that developed silver grains were
derived from the cell wall, the distance on either side of the center
of the cell wall that included 50% of the counted grains (the
half-distance) was determined (6). This analysis gave
half-distances for wild-type and pbpA spores of 80 and 89 nm, respectively, compared to a value of 83 nm obtained in previous
studies of cell wall synthesis in Bacillus megaterium (6). These values are also consistent with the published
value for the half-distance for tritium when samples are prepared by similar methods (13). Consequently, silver grains which were within the half-distance were considered to be derived from the wall
and only those grains were further analyzed with regard to distribution
along the outgrowing spore (see below). At least 200 grains within the
half-distance were analyzed in both the outgrowing wild-type and
pbpA spores, with 15 elongating wild-type spores and 23 elongating pbpA spores analyzed.
To determine the location of the newly synthesized PG for each strain,
the position of a silver grain (within the half-distance from the wall)
was used to determine the position of labeled PG in the wall by taking
the shortest distance between the grain and the wall; the distance
between this position and the center of the cell or of the forming
septum was then measured and expressed as a percentage of total cell
length (see Fig. 3). Due to the increased diameter of the
pbpA spores, a higher percentage of wall material is present
at the poles of pbpA spores than in wild-type spores. To
correct for this difference, an average cell length and width were
determined for each strain (see Fig. 3); for this analysis an
outgrowing spore was considered a rectangle with semicircles at each
pole. The average outgrowing spore for each strain was then divided
into 10 bins with identical amounts of wall material, and the grains
were placed into each bin based upon their distance to the center of
the cell as a percentage of cell length. As the cell is symmetrical,
bins at equal distances from the cell center were combined, so the
grains were grouped into five compartments. Grains associated with
forming septa were excluded from the analysis; outgrowing spores, which
are round, do not provide a reference point to analyze grain
distribution and were also excluded. All statistical analyses employed
Excel software (Microsoft).
| |
RESULTS |
Macromolecular synthesis, PG turnover, and location of PG synthesis
during spore outgrowth.
The alteration in cell morphology and the
slowed outgrowth of pbpA spores seen previously
(18) suggested that PG synthesis might be altered during
outgrowth of spores lacking PBP2a. Indeed, consistent with the slower
outgrowth of pbpA spores compared to that of wild-type
spores (Fig. 1A), PG synthesis during
spore outgrowth in 2× YT medium was also slightly slowed by the loss of PBP2a (Fig. 1B). In this medium [3H]thymidine
incorporation into DNA began to increase significantly between 60 and
90 min after the initiation of germination of wild-type spores,
pbpA spores, and spores of all other pbp mutants
examined and then paralleled the changes in the OD600
during subsequent outgrowth (data not shown). The relative rates of
protein synthesis were also similar to those found for PG synthesis
(data not shown). Chloramphenicol addition blocked (>98%) PG
synthesis during the first 120 min of germination and outgrowth of both
wild-type and pbpA spores (data not shown), indicating that
PG synthesis during outgrowth requires protein synthesis.
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FIG. 1.
Spore outgrowth (A) and PG synthesis (B) in strains
lacking HMW PBPs. The spores were heat shocked and inoculated into 2×
YT medium with 4 mM L-alanine containing
[3H]N-acetyl-D-glucosamine, as
described in Materials and Methods. The cells were harvested, and
TCA-insoluble radioactivity was measured as described in Materials and
Methods. The percentage of total radioactivity in the sample that was
TCA insoluble was divided by the initial OD600 to give the
units in Fig. 1B. The kinetics of spore outgrowth shown are from the
same experiment in which PG synthesis was monitored.  , PS832 (wild
type);  , PS2465 (pbpA);  , PS2466 (pbpA
ponA);  , PS2468 (pbpA pbpD).
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PG turnover was also measured during outgrowth of wild-type and
pbpA spores; for wild-type spores, the newly synthesized PG was stable for approximately 30 min, which is consistent with our
measurements of PG turnover in vegetative cells as well as with
previous work (Fig. 2) and data not shown
[16, 23]). However, newly synthesized PG in
pbpA spores was stable for longer than 30 min, and
subsequent PG turnover was faster in the outgrowing wild-type spores,
consistent with their higher rate of outgrowth (Fig. 2). Interestingly,
measurements of cell wall width from electron micrographs of spores
harvested 75 min after the initiation of spore germination revealed
that during outgrowth pbpA spores possess a thicker wall
than wild-type spores (59 ± 10 nm for pbpA spores
versus 45 ± 7.4 nm for wild-type spores; n > 100
spores for each strain; P < 0.05 by the two-tailed
Student's t test). It is possible that the increased
thickness of the outgrowing pbpA spore wall is due at least
in part to the slower PG turnover in pbpA spores (see
Discussion).
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FIG. 2.
PG turnover during outgrowth of wild-type and
pbpA spores. The spores were heat shocked and germinated in
2× YT medium with 4 mM L-alanine for 30 min in the
presence of
[3H]N-acetyl-D-glucosamine. Then
the cells were harvested and resuspended in fresh 2× YT medium with
unlabeled N-acetyl-D-glucosamine, and
TCA-insoluble radioactivity was measured, as described in Materials and
Methods. The amount of TCA-insoluble radioactivity immediately after
resuspension was set at 100%. , OD600 of PS832 (wild
type); , OD600 of PS2465 (pbpA);  , percent
incorporated
[3H]N-acetyl-D-glucosamine
remaining in PS832 (wild type);  , percent incorporated
[3H]N-acetyl-D-glucosamine
remaining in PS2465 (pbpA).
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An alternative explanation is that the wall may be thicker in only some
areas of pbpA spores during outgrowth due to a change in the
location of PG incorporation compared to that in wild-type spores. To
examine this possibility, wild-type and pbpA spores were
pulse labeled with
[3H]N-acetyl-D-glucosamine 60 min
after the initiation of spore germination and subjected to electron
microscopic autoradiography to determine the sites of PG synthesis
during spore elongation (Fig. 3A and
C). Initial analyses
confirmed that in these autoradiographs the silver grains were derived
from label in the cell wall (see Methods). Further statistical analysis
of grain distribution along the wall of the outgrowing spores revealed
that when cell shape was corrected for by dividing each cell into
compartments containing equal amounts of wall, the grain distribution
was similar for both wild-type and pbpA spores and appeared
uniform along the cell (Fig. 3B and D). This suggests that after the
initiation of spore elongation, sites of PG synthesis are similar for
the wild-type and pbpA spores, although this analysis does
not address sites of earlier PG synthesis (i.e., when outgrowing spores
are still round).
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FIG. 3.
Electron microscopic autoradiography of wild-type and
pbpA spores. The spores were heat shocked and inoculated
into 2× YT medium with 4 mM L-alanine, as described in
Materials and Methods. Sixty minutes after the initiation of spore
germination, 500 µCi of
[3H]N-acetyl-D-glucosamine was
added and the culture was incubated for 2 min, followed by the addition
of unlabeled N-acetyl-D-glucosamine to 1 mM.
After 10 min of further incubation, the cells were harvested and
processed for electron microscopic autoradiography, as described in
Materials and Methods. Examples of wild-type (A) and pbpA
(C) outgrowing spores are shown, with arrows indicating the labeled
wall. The statistical analysis of silver grain distribution was done as
described in Materials and Methods and is shown for the average
wild-type (B) and pbpA (D) outgrowing spore after half of
each cell was divided into 5 bins, with equal amounts of wall material
in each bin. Bars, 1 µM.
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The loss of PBP1 or PBP4 further delays outgrowth of spores lacking
PBP2a.
Previous work has suggested that HMW PBPs perform partially
redundant functions in B. subtilis (31). Given
the unique phenotype associated with the loss of PBP2a, we were
interested in determining the effect of the loss of other HMW PBPs in
addition to PBP2a. Consequently, mutations inactivating the genes
encoding the predominant class A HMW putative bifunctional
transglycosylases and transpeptidases, PBP1, -2c, and -4, and the class
B putative monofunctional transpeptidase PBP3 were combined with a
mutation in pbpA (Table 1). That these mutant strains lacked
the appropriate PBPs was confirmed by analysis of PBP profiles in
membranes isolated from the appropriate strains (Fig.
4 and data not shown). In 2× SG or 2×
YT medium, the vegetative growth rates of strains lacking PBP2a and
either PBP2c, PBP3, or PBP4 were essentially identical to those of the
wild-type strain while the strain lacking PBP2a and PBP1 (encoded by
ponA) grew at a rate identical to that of a strain lacking
only PBP1 (data not shown) (30). All of these multiple
pbp mutant strains produced dormant spores with heat
resistance properties and PG cortex structures essentially identical to
those of wild-type spores (data not shown).
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FIG. 4.
PBP profiles from membranes of various strains.
Membranes were isolated from vegetative cells and labeled with
FLU-C6-APA, as described in Materials and Methods.
Approximately 10 µg of total membrane protein was run on sodium
dodecyl sulfate-10% polyacrylamide gel electrophoresis for 4 h
at 100 V, and labeled PBPs were visualized with a FluorimagerSI. The
PBP pattern of wild-type cell membranes is like that previously found
(lane 1) (18). Lane 1, PS832 (wild type); lane 2, PS2466
(pbpA ponA); lane 3, PS2467 (pbpA pbpC); lane 4, PS2468 (pbpA pbpD). The PBPs were designated as previously
described (31).
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Spores of all multiple pbp mutants initiated germination
similarly to wild-type spores (Fig. 1A and data not shown). The loss of
either PBP2c or PBP3 in addition to PBP2a did not further alter either
the rate of spore outgrowth, the previously observed morphological changes or the rate of PG synthesis during spore outgrowth, or spore
viability compared to the effects of the loss of PBP2a alone (data not
shown). However, the loss of either PBP4 or PBP1 in a pbpA
background further slowed spore outgrowth in 2× YT medium (Fig. 1A)
and decreased PG synthesis comparably to the decrease in the rate of
spore outgrowth (Fig. 1B); these effects were much greater than the
effects of loss of either PBP1 or -4 alone (data not shown) (29,
30). Examination of spores 120 min after the initiation of spore
germination in 2× YT medium revealed that 45% of outgrowing spores
lacking PBP4 and PBP2a had the increased diameter observed in spores
lacking only PBP2a while the remaining spores showed no elongation and
no increase in diameter (n > 200 spores; data not
shown). In contrast, 85% of spores lacking only PBP2a exhibited the
increased diameter 120 min after the initiation of spore outgrowth. No
spores lacking PBP1 and PBP2a outgrowing in 2× YT medium showed the
increased cell diameter exhibited by outgrowing pbpA spores,
and outgrowth of pbpA ponA spores was accompanied by severe
bending (Fig. 5C and Table
2).
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FIG. 5.
Morphologies of cells and outgrowing spores of the
wild-type strain and strains lacking PBP2a or PBP2a and PBP1. The
spores were heat shocked and inoculated into either PAB medium, PAB
medium with 500 mM NaCl, or 2× YT medium, all of which contained 4 mM
L-alanine. Outgrowing spores were harvested 120 min after
the initiation of spore germination unless otherwise noted. The
following strains (genotypes) and growth or outgrowth conditions are
shown: (A) PS2062 (ponA), vegetative cells in PAB medium
with 50 µM MgCl2; (B) PS2466 (pbpA ponA),
vegetative cells in PAB medium with 50 µM MgCl2; (C)
PS2466 (pbpA ponA), spores outgrowing in 2× YT medium with
no additions, harvested 150 min after the initiation of spore
germination; (D) PS832 (wild type), spores outgrowing in PAB medium
with no additions; (E) PS2465 (pbpA), spores outgrowing in
PAB medium with no additions; (F) PS2465 (pbpA ponA), spores
outgrowing in PAB medium with no additions; and (G) PS2465
(pbpA), spores outgrowing in PAB medium with 500 mM NaCl.
Bars, 10 µM.
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In both this and previous work we observed that when we plated equal
numbers of wild-type and pbpA spores, the pbpA
spores gave 50% fewer colonies than the wild type (18)
(Table 3). However, spores lacking PBP2a
and PBP1 had greatly decreased viability, while those lacking either
PBP1 or PBP1 and PBP4 had viabilities essentially identical to that of
wild-type spores (Table 3 and data not shown). The addition of 10 mM
Mg2+ to LB plates significantly increased the viability of
spores lacking PBP2a and PBP1 (Table 3), suggesting that these spores require increased levels of divalent cations for outgrowth. This is
consistent with recent work, which demonstrated that decoated spores
lacking PBP1 require increased concentrations of Mg2+ for
outgrowth (19). However, colony formation from spores also requires cell growth, and ponA cells require a high
concentration of Mg2+ for growth (19).
To determine when during outgrowth spores were no longer viable,
LIVE/DEAD assays were performed during spore outgrowth in 2× YT
medium. After 90 min of spore outgrowth, and even more so after 120 min, a significant number of dead cells were observed with the strain
lacking PBP2a and PBP1 (Table 3), this cell death could explain the
extremely slow outgrowth of spores of this strain. The percentage of
dead cells given at 120 min is actually a significant underestimate of
the true number, since chains of cells were present at this time,
suggesting that significant division of the cells that remained viable
had already occurred (data not shown). In contrast to the results with
spores lacking PBP1 and -2a, outgrowth of spores lacking PBP2a or PBP2a
and -4 was not accompanied by significant cell death (Table 3).
PG structure during outgrowth of spores lacking HMW PBPs.
While the rate of PG synthesis was decreased similarly in outgrowing
spores lacking PBP1 and -2a or PBP4 and -2a, the viabilities of the two
strains during spore outgrowth were drastically different. One
explanation for this difference is that the structures of the PGs being
synthesized by the two strains might differ significantly. To explore
this possibility, PG was purified from spores of a variety of HMW PBP
mutants after 30, 60, and 90 min of outgrowth in 2× YT medium and the
PG was digested and analyzed by HPLC. The amounts of PG synthesized by
30 and 60 min are not dramatically different in various mutant strains
(Fig. 1B), so PG from comparable amounts of outgrowing spores was
loaded onto the HPLC column for each mutant strain. However, for
samples analyzed at 90 min of outgrowth, PG from twice as many spores
lacking PBP2a and either PBP1 or -4 was loaded than from spores of
other strains. The pattern of muropeptide peaks varied significantly
among the 30-, 60-, and 90-min samples (data not shown), but at each
time point similar patterns were found in the wild-type strain and most
PBP mutants examined, even though distinct morphological differences
were observed between some strains at 90 min. However, 60 min after the
initiation of spore germination, spores lacking PBP1 and PBP2a exhibited significant changes in PG structure, as determined by reverse-phase HPLC (Fig. 6).
Cochromatography with known standards, amino acid and amino sugar
analysis, and mass spectrometry of peaks 4, 5, and 6 revealed that in
the strain lacking PBP1 and PBP2a (Fig. 6A), there was an increase in
levels of disaccharide tetrapeptide (peak 4) and a corresponding
decrease in a disaccharide tetrapeptide cross-linked to either a
disaccharide tetrapeptide with a glycine present in the cross-link or a
disaccharide pentapeptide with glycine in either the fourth or fifth
position of the side chain (peak 5/6) compared to a strain lacking only
PBP1 (Fig. 6B) or to the wild-type strain (data not shown). Peaks 5 and
6 had identical amino acid and amino sugar compositions, so we suspect the split peak is due to a partial modification of the muropeptide, possibly partial amidation of the peptide side chain. The ratio of
cross-linked species to monomer (Fig. 6A, ratio of peak 5/6 to peak 4)
was 1.10 in ponA spores compared to 0.72 in pbpA
ponA spores. Similar (within 7%) results were seen in duplicate
analyses of two independent spore preparations of both strains. This
suggests that the percentage of total PG cross-linking is decreased
during outgrowth of spores lacking PBP1 and PBP2a compared to that in wild-type spores. Since these changes in PG structure were not detected
in the strain lacking PBP1 and PBP2a either 30 or 90 min after
initiation of spore germination, this suggests that the change in PG
cross-linking is transient.
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|
FIG. 6.
Differences in structures of PG from outgrowing
wild-type spores and spores lacking PBP1 and PBP2a. PG from outgrowing
spores was harvested 60 min after the initiation of spore germination
and purified, digested, reduced, and separated by HPLC with a 0.1%
TFA-20% acetonitrile gradient as described previously
(25). The muropeptide profile of a strain lacking both PBP1
and PBP2a (A) is compared to the profile of a strain lacking only PBP1
(B); the latter is identical to that from the wild-type strain (data
not shown). The arrows denote peaks whose magnitude is significantly
different in the strains lacking PBP1 and PBP1 and -2a. Differences in
retention times between the two gradients are the result of slight
differences in buffer composition among the different HPLC runs.
|
|
Divalent cation sensitivity of outgrowing spores lacking HMW
PBPs.
Given that the addition of 10 mM MgCl2 to LB
plates restored viability to spores lacking PBP1 and PBP2a, we further
explored the spore outgrowth and vegetative growth of multiple HMW PBP mutants in PAB medium, which contains low levels of divalent cations. All strains grew in PAB medium except the strain lacking PBP1 and
PBP2a, which, like a strain lacking only PBP1, did not grow (reference
19 and data not shown); however, the strain lacking PBP1 and 
2a appeared to require more Mg2+ for growth than
the strain lacking only PBP1 (data not shown). The addition of 50 µM
Mg2+ to PAB medium allowed some growth of a strain lacking
both PBP1 and PBP2a, and microscopic examination of these cells
revealed dramatic bending and filamentation, more so than with a strain lacking only PBP1 (Fig. 5A and B), suggesting a role for PBP2a in
vegetative growth.
The rate of outgrowth of pbpA spores in PAB medium was
significantly lower than that of wild-type spores (Table 2). Ninety minutes after the initiation of spore germination in PAB medium, pbpA cultures had many small round cells while wild-type
spores had initiated elongation (data not shown). After 120 min of
spore outgrowth an increasing number of pbpA cells did
exhibit an increased diameter, but little cell elongation had taken
place compared to that with wild-type spores (Fig. 5D and E). It is
possible that the release of cations during germination is responsible for pbpA spore outgrowth in PAB medium, as this appears to
be the case for spores lacking PBP1 (19). Spores lacking
PBP1 and PBP2a outgrowing in PAB medium did not show the increased cell diameter exhibited by outgrowing pbpA spores, and very
little elongation occurred compared to that of wild-type spores (Fig. 5D and F).
The addition of 10 mM Mg2+ to PAB medium greatly improved
the rate of outgrowth of spores of all pbp mutant strains
whose outgrowth was previously found to be severely compromised (Table
2). The bent cells previously observed in the strain lacking PBP2a and PBP1 were no longer present, while straight rods with decreased diameter were (data not shown).
Sensitivity of outgrowing spores lacking HMW PBPs to monovalent
cations.
Previously we had found that the inclusion of 500 mM NaCl
in PAB medium prevented outgrowth of spores lacking PBP1 and resulted in lysis of wild-type spores 120 to 150 min after the initiation of
spore outgrowth (19). While wild-type spores were able to initially elongate in PAB medium with 500 mM NaCl (19),
pbpA spores remained round and swollen and no elongation was
observed (<1% of spores elongated) (Fig. 5G); however, lysis occurred
as with wild-type spores outgrowing in this medium. The inclusion of
1.0 M sorbitol in PAB medium did not give these effects (data not
shown). The structure of PG purified from these outgrowing spherical
pbpA spores 120 min after the initiation of spore
germination was identical to that of PG harvested from wild-type spores
which had initiated elongation. The addition of 10 mM MgCl2
improved both the rate of outgrowth and elongation efficiency of
pbpA spores in PAB medium with 500 mM NaCl and prevented
cell lysis in the presence of high salt but did not affect the
formation of cells with increased diameters (Table 2 and data not
shown). Spores lacking PBP2a and PBP4 exhibited outgrowth similar to
that of pbpA spores in PAB medium with 500 mM NaCl with
regard to rate and morphology, while spores lacking PBP1 and -2a did
not initiate outgrowth in this medium (Table 2), as was found with
spores lacking only PBP1 (19).
| |
DISCUSSION |
One difficulty in assessing the function of individual HMW PBPs is
that these proteins tend to possess at least partially redundant enzyme
activities (31). However, during outgrowth of B. subtilis spores, PBP2a, a class B HMW PBP with putative transpeptidase activity, appears to be uniquely important, at least
initially, for cell elongation and the determination of proper cell
diameter (18). Studies of the PG metabolism of outgrowing pbpA spores revealed slight decreases in PG synthesis and a
significant delay in PG turnover compared to that in wild-type spores,
suggesting that the coupling of PG synthesis and degradation may be
altered in pbpA spores. However, in both strains no PG
turnover was detected for at least 30 min after PG synthesis,
consistent with models for PG metabolism during vegetative growth,
where it has been established that new PG is inserted along the inner
wall and spreads to the outer surface, where it is degraded by
autolysins (24).
The slower PG turnover early in pbpA spore outgrowth may
also explain, at least in part, the increased thickness of the cell wall in those spores at that time, as the magnitude of the decrease in
PG turnover early in outgrowth appears greater than the slight slowing
of PG synthesis. However, pbpA spores eventually give rise
to vegetative cells which are identical to those of the wild-type strain (18). Consequently, any imbalance between PG
synthesis and degradation early in the outgrowth of pbpA
spores must be corrected as outgrowth proceeds. An additional
explanation for the increased cell wall thickness of pbpA
spores early in outgrowth is that, since outgrowing wild-type and
pbpA spores have similar volumes (18), newly
synthesized PG is inserted into a smaller surface area in the
outgrowing spherical pbpA spore than in the more cylindrical
outgrowing wild-type spore, as a sphere has a smaller surface area than
a cylinder.
pbpA spores which had begun elongation showed no dramatic
differences from wild type spores in the distribution of newly
synthesized PG, after the data had been normalized for cell shape. This
analysis excluded pbpA spores which remained round, where
incorporation of PG precursors truly may have been different.
Structural analysis of purified PG from outgrowing pbpA
spores also revealed no differences from that in outgrowing wild-type
spores, even when distinct morphological differences were observed. As
we currently have no way of observing and quantitating changes in the
three-dimensional structure of PG, we are unable to determine the exact
nature of the defect during elongation of pbpA spores.
Both the kinetics and the morphology of outgrowing pbpA
spores are sensitive to levels of monovalent and divalent cations in
the growth medium. Interestingly, in PAB medium with 500 mM NaCl,
pbpA spores remain spheres, with no detectable elongation before lysis, confirming the importance of PBP2a in spore elongation. The inclusion of high levels of divalent cations in this medium promotes spore elongation, even in the absence of PBP2a, but the mechanism of this effect of divalent cations is unclear. Previously we
found that cells lacking PBP1 require increased levels of divalent cations for growth (19). The requirement of pbpA
spores for elevated Mg2+ levels might be related, as a
strain lacking both PBP1 and PBP2a requires higher levels of
Mg2+ for spore outgrowth and vegetative growth than either
single mutant alone (Table 3 and data not shown). This finding suggests that PBP2a also contributes to PG synthesis during vegetative growth,
as has been previously reported (33).
To further define a role for PBP2a in spore outgrowth, we examined
strains lacking PBP2a and one of the predominant class A HMW PBPs
(PBP1, -2c, or -4) and a strain lacking PBP2a and the class B PBP3. The
additional loss of either PBP2c or PBP3 had no effects different from
those of the loss of PBP2a alone. However, the loss of PBP4 further
slowed spore outgrowth, while the loss of PBP1 had an even more
dramatic effect on spore outgrowth and viability. This is consistent
with previous work demonstrating that PBP1 and PBP4 perform partially
redundant functions, with PBP4 being the subordinate class A HMW PBP
and PBP2c having an even more minor role (31).
Previous work has demonstrated that PBP1, a class A HMW PBP with
putative transglycosylase and transpeptidase activities, is also
involved in establishing the correct diameter of vegetative cells
(30). Since PBP1 is expressed early in spore outgrowth (20, 30), it is not surprising that this protein is also
important for reestablishing the rod-shaped cell morphology, starting
from the elliptical dormant spore. The increased diameter observed in
pbpA spores during outgrowth is absent in spores lacking
PBP1 and PBP2a, suggesting that the activity of PBP1 is required for the increased diameter. Additionally, analysis of PG from spores lacking PBP1 and PBP2a suggests that the degree of PG cross-linking is
transiently decreased 60 min after the initiation of spore germination.
One possibility is that wild-type spores also undergo this transient
decrease in cross-linking during spore outgrowth, but due to the time
points we analyzed, this decrease was not detected. Another possibility
is that pbpA ponA spores are transiently subjected to
increased PG hydrolytic activity during outgrowth. However, a third and
more intriguing possibility is that PBP1 and PBP2a possess a redundant
transpeptidase activity which is crucial to proper spore outgrowth, the
loss of which results in decreased cross-linking, which is corrected
later in outgrowth as levels of other PBPs increase. PBP1 and PBP2a
have both been found on the inner forespore membrane of dormant spores,
consistent with the idea that these enzymes act in proximity to one
another during spore outgrowth (4).
One possible model to explain the different morphology and PG structure
of outgrowing spores lacking either PBP1, PBP2a, or both PBPs is that
during wild-type spore outgrowth PBP1 synthesizes PG strands of a
specific length, whose peptide side chains are then cross-linked by
PBP2a and PBP1 to facilitate elongation at the appropriate diameter.
Spores lacking PBP1 thus produce shorter glycan strands, which are
initial substrates for PBP2a cross-linking, in such a way that a
narrower cell diameter is established but elongation still occurs,
perhaps facilitated by PBP2a, which may generate cross-links
perpendicular to the glycan strands. However, whether the cross-linked
glycan strands run parallel to each other or are in a more disorganized
nonparallel configuration is not clear. At this time we also cannot
determine whether the absence of specific PBPs alters the orientation
of cross-linked glycan strands with respect to each other. In the
absence of PBP2a, PBP1 polymerizes PG strands of the appropriate
length, which are then cross-linked inappropriately, perhaps by other
HMW PBPs or PBP1. Thus, instead of elongation, a large sphere forms,
whose diameter is slowly reduced as PG remodeling occurs and other HMW
PBPs are synthesized (17, 28, 29). In the absence of both
PBP1 and -2a, short glycan strands which are cross-linked less
efficiently, are made, resulting in poor cell elongation and cell death.
An interesting finding in this work is the presence of glycine in the
PG purified from outgrowing spores, as previous studies of PG from
vegetative B. subtilis cells did not report the presence of
glycine (36, 38). More recently, glycine was detected in the
PG of dormant spores lacking CwlD, a putative
muramoyl-L-alanine amidase (26), and glycine is
an important component in the PG of other gram-positive organisms, such
as Staphylococcus aureus (15). It is unclear
whether glycine is incorporated into the wall transiently during spore
outgrowth or whether there is truly a glycine component of B. subtilis vegetative PG; this may be an important area for further study.
We are grateful to Kit Pogliano for protocols and to Steven Harris for
the use of his microscope.
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Abstract
Introduction
