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Journal of Bacteriology, May 1999, p. 3201-3211, Vol. 181, No. 10
0021-9193/99/$04.00+0
Copyright © 1999, American Society for Microbiology. All rights reserved.
Septal Localization of Penicillin-Binding
Protein 1 in Bacillus subtilis
Lotte B.
Pedersen,1
Esther R.
Angert,2 and
Peter
Setlow1,*
Department of Biochemistry, University of
Connecticut Health Center, Farmington, Connecticut
06032,1 and Department of Molecular and
Cellular Biology, The Biological Laboratories, Harvard University,
Cambridge, Massachusetts 021382
Received 30 November 1998/Accepted 26 February 1999
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ABSTRACT |
Previous studies have shown that Bacillus subtilis
cells lacking penicillin-binding protein 1 (PBP1), encoded by
ponA, have a reduced growth rate in a variety of growth
media and are longer, thinner, and more bent than wild-type cells. It
was also recently shown that cells lacking PBP1 require increased
levels of divalent cations for growth and are either unable to grow or
grow as filaments in media low in Mg2+, suggesting a
possible involvement of PBP1 in septum formation under these
conditions. Using epitope-tagging and immunofluorescence microscopy, we
have now shown that PBP1 is localized at division sites in vegetative
cells of B. subtilis. In addition, we have used
fluorescence and electron microscopy to show that growing ponA mutant cells display a significant septation defect,
and finally by immunofluorescence microscopy we have found that while FtsZ localizes normally in most ponA mutant cells, a
significant proportion of ponA mutant cells display FtsZ
rings with aberrant structure or improper localization, suggesting that
lack of PBP1 affects FtsZ ring stability or assembly. These results
provide strong evidence that PBP1 is localized to and has an important function in the division septum in B. subtilis. This is the
first example of a high-molecular-weight class A PBP that is localized to the bacterial division septum.
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INTRODUCTION |
The bacterial peptidoglycan is
composed of glycan strands with peptide side chains which cross-link
the glycan strands to each other. During growth, new disaccharide
pentapeptide units become inserted into the existing peptidoglycan by
the penicillin-binding proteins (PBPs) (17, 23, 36, 37).
PBPs can be divided into three groups: low-molecular-weight PBPs and
class A and class B high-molecular-weight (HMW) PBPs (17).
Low-molecular-weight PBPs are usually monofunctional
DD-carboxypeptidases, which are involved in regulating the number of
peptide cross-links (17). Class A HMW PBPs possess both
transglycosylase and transpeptidase activities (24, 55) and
are somewhat functionally redundant (26, 46), while class B
HMW PBPs are monofunctional transpeptidases, some of which have
essential functions in cell septation and regulation of cell shape
(1, 17, 18, 38).
During cell growth rod-shaped bacteria display two modes of
peptidoglycan synthesis: elongation and septation (23, 36, 37). For Escherichia coli, it is widely assumed that
the class B HMW PBP2 is involved in elongation while the class B HMW
PBP FtsI (also known as PBP3) is needed for septation (38, 53, 61). Homologues of PBP2 and FtsI are also found in Bacillus subtilis (34, 62). Since class B HMW PBPs have no
transglycosylase activity (1, 17, 57), these proteins cannot
synthesize peptidoglycan. Therefore, it has been proposed that the
class A HMW PBPs synthesize and to some extent cross-link primers,
which are then used by class B HMW PBPs during septation and elongation (61). This model is supported by studies using specific
inhibitors of the major E. coli class A HMW PBPs
(61) and by affinity chromatography studies, which
indicate that the E. coli class A HMW PBPs PBP1a and PBP1b
interact either directly or indirectly with the class B HMW PBPs PBP2
and FtsI (23). In addition, genetic studies have
shown that E. coli PBP1b is a possible helper protein for FtsI (15), and antibiotic inhibition studies have indicated that triggering of cell lysis by inhibitors of PBP1a and PBP1b in
E. coli is linked to cell division (16),
suggesting a role for class A HMW PBPs in cell division. Recent
evidence from our laboratory suggests that in growth media low in
Mg2+ the class A HMW PBP1 may also play a role in cell
division in B. subtilis, because cells lacking PBP1
sometimes grow as filaments (35). PBP1, encoded by
ponA, is expressed predominantly during vegetative growth,
and mutants lacking PBP1 have a reduced growth rate in a variety of
growth media (45, 46). Although vegetative B. subtilis cells contain two other class A HMW PBPs (PBP2c and PBP4)
(43, 44), PBP1 appears to be functionally more important than PBP2c and PBP4 (35, 46).
In addition to PBPs, septum formation in E. coli requires
the concerted action of the division proteins FtsZ, FtsA, FtsK, FtsL,
FtsN, FtsQ, FtsW, and ZipA, which are believed to form a complex at the
division site, called the divisome (10, 37, 48), whose key
component is the FtsZ ring (9, 48). Except for FtsL, the
other Fts proteins as well as ZipA have now been localized to the
division site in E. coli (2-4, 11, 19, 31, 58-60,
64); in B. subtilis the division proteins DivIB (an
FtsQ homologue), DivIC, and FtsZ have also been localized to
division sites (21, 25, 31). However, no class A HMW PBPs
have been demonstrated to localize to division sites in either
organism, although immunoelectron microscopy showed that PBP1b from
E. coli is present in higher amounts around inner
membrane-outer membrane contact areas than in other parts of the cell
(7).
In this communication, we report studies of the subcellular
localization of PBP1 in exponentially growing B. subtilis
cells using immunofluorescence microscopy; these studies show that PBP1 localizes to division sites.
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MATERIALS AND METHODS |
B. subtilis strains, recombinant plasmids, and growth
conditions.
All B. subtilis strains used are
derivatives of PS832, a prototrophic revertant of strain 168. Transformation of B. subtilis was as described previously
(6), and transformants were selected on 2× SG
(30) agar plates with chloramphenicol (Cm; 5 µg/ml). Strain PS2062 carrying a spectinomycin resistance (100 µg/ml) cassette (Spr) in the ponA gene
(ponA::Spr) has been described
previously (45). Strain LP27 was generated by transformation
of PS832 with plasmid pLP3 (see below), resulting in integration of
pLP3 into the chromosome by a single crossover event at the 3' end of
the coding region of ponA. Therefore, strain LP27 contains a
24-bp DNA sequence encoding the FLAG epitope immediately before the
stop codon of the ponA gene, followed by a Cm resistance marker and the sequence corresponding to the 3' noncoding region of
ponA. Routinely, cells were grown at 30 or 37°C overnight
on 2× SG agar plates with or without appropriate antibiotics,
inoculated into 20, 50, or 100 ml of 2× YT medium (containing [per
liter] 16 g of tryptone, 10 g of yeast extract, 5 g of
NaCl) or Penassay broth with no antibiotics, and grown at 30 or 37°C.
Plasmids and epitope tagging.
Plasmid pDPC273 was a gift
from David L. Popham (Virginia Polytechnic Institute, Blacksburg, Va.).
It is a derivative of plasmid pJH101 (14), which contains an
~3.8-kb insert comprising the region from the HindIII
site in the gene upstream of ponA (prfA) to the
EcoRV site downstream of ponA (45).
For tagging of PBP1 with the FLAG epitope, a megaprimer PCR-based
method (49) was used. All oligonucleotides were purchased
from GIBCO and are listed in Table 1. All
PCRs were carried out with Thermus aquaticus DNA polymerase
(GIBCO). The first round of PCR was carried out by using the primers
ponAP2 and ponAP3 with chromosomal DNA from strain PS832 as a template
and gave a 215-bp product corresponding to the region from nucleotide
(nt) 4050 to nt 4234 of the ponA operon (45) plus
24 bp encoding the FLAG epitope. The 215-bp PCR product was purified by
using a PCR purification kit (QIAGEN) and used as a megaprimer in a
second round of PCR with ponAP1 used as the upstream primer and
linearized pDPC273 plasmid DNA used as the template. The product of the
second round of PCR (720 bp), corresponding to the region from nt 3553 to nt 4234 of the ponA operon (45), was ligated
into pCR2.1 (Invitrogen), and after transformation into E. coli INV
F' competent cells (Invitrogen), plasmids were prepared
and the inserts were sequenced by using an automated DNA sequencer. A
plasmid with an insert whose sequence was as expected was digested with
EcoRI and BamHI, and the 712-bp insert was
ligated into plasmid pJH101 (14) digested with the same
enzymes to generate plasmid pLP3, which was used to transform B. subtilis to Cm resistance. Transformants were screened by PCR using primers ponAP4 and p1219, and pFLAG and p1219, respectively.
Membrane preparation, penicillin-binding assay, SDS-PAGE, and
Western blot analysis.
Cells from 50- or 100-ml log-phase cultures
(optical density at 600 nm [OD600] of ~0.5) grown in
2× YT medium were harvested by centrifugation (7,000 × g, 10 min), washed with 5 ml of wash buffer (50 mM Tris [pH 8],
1 mM MgCl2, 1 M KCl, 1 mM phenylmethylsulfonyl fluoride
[PMSF]), resuspended in 0.7 ml of buffer A (100 mM Tris-HCl [pH 8],
1 mM MgCl2, 1 mM 
-mercaptoethanol [-ME], 1 mM PMSF) containing 285 µg of lysozyme/ml, and incubated on ice for 20 min.
Cells were broken by sonication in the presence of glass beads, the
lysate was cleared by centrifugation (20,000 × g for 15 min at 4°C), and membranes were pelleted by centrifugation at
100,000 × g for 1 h at 4°C. The supernatant
fluid from the latter step was stored at 
20°C and used for DNA
quantitation. The membranes were washed with 1 ml of buffer B (50 mM
Tris-HCl [pH 8], 1 mM -ME, 1 mM PMSF), resuspended in 90 µl of
buffer B, and stored at 80°C. Membranes were incubated with
fluorescein-hexanoic acid-6-aminopenicillanic acid (FLU-C6-APA) and
assayed for penicillin-binding activity as previously described
(46). For Western blotting, different amounts (10 to 40 µg) of crude membrane protein in 10 µl of buffer B were mixed with
10 µl of 2× sodium dodecyl sulfate (SDS) sample buffer
(29) and boiled for 4 min, and proteins were separated by
SDS-10% polyacrylamide gel electrophoresis (PAGE). Various amounts
(9.4 to 600 ng) of C-terminal FLAG-bacterial alkaline phosphatase (BAP)
(Sigma) were loaded on the same gels to serve as a standard for
quantitative Western blotting. After electrophoresis the proteins on
SDS gels were transferred to Immobilon-Psq membranes
(Millipore) in transfer buffer (3 g of Tris base/liter, 14.4 g of
glycine/liter, 20% methanol [vol/vol], 0.08% [wt/vol] SDS), and
the membranes were subjected to immunoblotting as described below.
Immunoblotting and protein and DNA quantitation.
Immunoblotting was carried out essentially as in the protocol for the
BM Chromogenic Western blotting kit (Boehringer). The primary antibody
was anti-FLAG M2 monoclonal antibody (4 µg/ml) (Sigma). The secondary
antibodies were a commercially available mixture of goat anti-mouse
immunoglobulin G (IgG) and goat anti-rabbit IgG, both conjugated to
alkaline phosphatase (1:2,000 dilution) (Boehringer). The blots were
scanned, and the relative FLAG content of each band was determined by
using an IS-1000 digital imaging system and software from Alpha
Innotech Corporation. Protein concentrations were determined by the
Lowry procedure (32) by using bovine serum albumin (BSA) as
the standard. DNA quantitation was performed with the Hoechst
33258-based fluorescent DNA quantitation kit (Bio-Rad) by using calf
thymus DNA (Bio-Rad) as a standard. Calf thymus DNA has about the same
A+T content (58%) as B. subtilis DNA (56.5%
[28]), and hence is appropriate to use as a standard in this assay.
Preparation of cells for immunofluorescence microscopy.
For
FLAG localization, cells were grown at 37°C in 2× YT medium to an
OD600 of ~0.5 and prepared for immunofluorescence
microscopy as described previously (20, 41) with
modifications. Briefly, cells were fixed for 20 min at room temperature
in 4.4% (wt/vol) paraformaldehyde-28 mM NaPO4 (pH 7),
washed three times with phosphate-buffered saline (PBS; containing
[per liter] 8 g of NaCl, 0.2 g of KCl, 1.44 g of
Na2PO4, 0.24 g of
KH2PO4 [pH 7.4]), resuspended in 100 µl of
GTE (50 mM glucose, 20 mM Tris-HCl [pH 7.5], 10 mM EDTA), treated
with lysozyme (2 mg/ml) for 30 to 60 s at room temperature, washed
twice with PBS, resuspended in 70 µl of PBS, and applied to
poly-L-lysine-coated microscope slides. The excess cell
suspension was removed, and the slides were washed two times with PBS
and allowed to dry completely. After rehydration with PBS, the slides were blocked for 20 min at room temperature with 2% (wt/vol) BSA in
PBS and incubated for 2.5 h at room temperature in PBS containing 10 µg of anti-FLAG M2 monoclonal antibody/ml and 0.1% (wt/vol) BSA.
The slides were washed six times with PBS and incubated overnight at
4°C in PBS containing a 1:100 dilution of biotinylated goat anti-mouse IgG (Molecular Probes) and 0.1% (wt/vol) BSA. Slides were
subsequently washed six times with PBS and incubated for 2 h at
room temperature with PBS containing 0.1% (wt/vol) BSA and 5 µg of
indocarbocyanine (Cy3)-conjugated streptavidin (Jackson ImmunoResearch
Laboratories) per ml. After being washed six times with PBS, the slides
were mounted by using the SlowFade antifade kit from Molecular Probes.
For FtsZ localization, cells were grown as described above and fixed by
adding 1 ml of cell culture to 10 ml of cold (
20°C)
80% methanol
as described (
22) with modifications. After fixing
for
1 h at room temperature, 200 µl of 16% (wt/vol)
paraformaldehyde
was added to the cell suspension, and after a 5-min
incubation
at room temperature, cells were pelleted by centrifugation
at
3,000 ×
g for 15 s at room temperature, the
supernatant was discarded,
and cells were gently resuspended in the
residual fluid by tapping
the tube. Then 1 ml of 80% methanol was
added, and the cells were
spun gently as described above, the
supernatant was removed, and
the pellet was resuspended in GTE.
Lysozyme was added to 2 mg/ml,
and the cells were placed on
poly-
L-lysine-coated microscope slides
and incubated at
room temperature for 30 to 60 s. Excess cell
suspension was
removed, and the slides were washed, dried, rehydrated
and blocked with
BSA as described for FLAG localization, and incubated
for 2 h at
room temperature with a 1:500 dilution of affinity-purified
rabbit
antiserum against
B. subtilis FtsZ (a gift from Petra A.
Levin, Massachusetts Institute of Technology, Cambridge, Mass.)
in PBS
containing 2% (wt/vol) BSA. After being washed 10 times
with PBS, the
slides were incubated for 2 h with 3.75 µg of Cy3-conjugated
donkey anti-rabbit serum (Jackson ImmunoResearch Laboratories)
per ml
in PBS containing 2% (wt/vol) BSA, washed 10 times with
PBS, and
mounted for microscopy as for cells stained for the FLAG
epitope.
Staining of cell walls and nucleoids.
For staining of cell
walls and/or nucleoids of immunostained cells, 2 µg of
4',6'-diamidino-2-phenylindole (DAPI) (Sigma) per ml and/or 2 µg of
Oregon green- or fluorescein isothiocyanate (FITC)-conjugated wheat
germ agglutinin (WGA) (Molecular Probes) was included in the Cy3 mix.
(WGA is a lectin that binds to oligomers of
N-acetylglucosamine and N-acetylmuramic acid and
has been found to bind well to lysozyme-treated cell walls of B. subtilis [5, 42, 52].) For cell wall and nucleoid
staining only, cells grown in 2× YT medium at 30 or 37°C were
treated like cells stained for the FLAG epitope except that incubations
with antibodies and streptavidin were omitted and the slides were
incubated for 2 h at room temperature with PBS containing 0.1%
(wt/vol) BSA, 0.2 µg of DAPI/ml, and 2 µg of Oregon green-WGA/ml
before the final PBS wash. In some cases, propidium iodide (10 µg/ml)
was used instead of DAPI to visualize RNA, DNA, and septa
(31).
Fluorescence microscopy and image analysis.
Cells (see Fig.
3) were visualized with a Zeiss Axiovert 100 fluorescence microscope
equipped with a 63× Plan-NEOFLUAR immersion lens (Zeiss) and a
standard filter block for visualizing Cy3 and Oregon green; images were
captured with a cooled charge-coupled device camera by using exposure
times of 700 (Cy3) or 500 (Oregon green) ms. Fluorescence microscopy of
cells (see Fig. 4 and 7) was performed with an Olympus BX60 microscope
equipped with a 100× UPlanFluor objective and standard filter sets for
visualizing DAPI, Cy3, and Oregon green or FITC; images were captured
with a MicroMax (Princeton Instruments)-cooled charge-coupled device camera driven by the MetaMorph software package (version 3.0; Universal
Imaging). The exposure times were 1 s for both Oregon green and
DAPI (see Fig. 4) and 500 ms for all fluorophores (see Fig. 7).
All images were transferred to a Power Macintosh computer and processed
by using Adobe Photoshop version 4.0. Images of control
cells were
captured and processed identically to those of cells
carrying
ponA-FLAG or
ponA::Sp
r. Cell lengths
were determined on images of Oregon green-WGA-stained
cells by using
the public domain NIH Image program (
38a) and
by drawing a
straight (straight cells) or segmented (bent cells)
central line
between the two poles of each cell and converting
the obtained pixel
value to micrometers. A pole was either a free
hemispherical cap at the
end of a cell or a very bright green
band between two cells in a chain.
By double staining of cells
with propidium iodide and Oregon green-WGA,
completed septa were
found to stain much brighter with Oregon green-WGA
than nascent
septa (data not shown). Although a small fraction (<5%)
of the
completed septa scored may be incomplete septa, the average cell
length obtained by this method (3.9 µm;
n = 137) for
wild-type
cells grown with a doubling time of 32 min is within the
range
of those reported for
B. subtilis SG38 grown with
doubling times
of 40 min (3.6 µm) and 30 min (4.5 µm), respectively
(
51).
Electron microscopy.
Vegetative B. subtilis cells
grown in 2× YT medium at 37°C were fixed, processed, and analyzed by
transmission electron microscopy as described previously
(50).
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RESULTS |
Epitope tagging of PBP1.
To localize PBP1 in B. subtilis by immunofluorescence microscopy, we added the FLAG
epitope (DYKDDDDK) at the C terminus of PBP1. The FLAG epitope is very
hydrophilic (pI of 3.85) and hence has a high probability of being
surface localized. By using PCR and a plasmid for integration into the
B. subtilis chromosome by a single crossover event (see
Materials and Methods), we generated a strain (LP27) in which a 24-bp
sequence encoding the FLAG epitope was inserted into the chromosomal
ponA gene (45) immediately before the stop codon.
Note that this tagged ponA (ponA-FLAG) is the
only ponA gene in this strain. To verify the presence of FLAG-tagged PBP1, membranes from log-phase cells of strain LP27 (ponA-FLAG) grown at 37°C in 2× YT medium were prepared
and membrane proteins were analyzed by SDS-10% PAGE and
immunoblotting with a monoclonal anti-FLAG antibody. As shown in Fig.
1A, membranes from ponA-FLAG
cells (lane 1) contained a protein of about 107 kDa that was recognized
by the FLAG antibody while no protein bands were detected in the lanes
containing membranes from wild-type cells (strain PS832; lane 3) or
cytoplasmic proteins from ponA-FLAG cells (lane 2). The
calculated molecular mass of PBP1-FLAG is 100.6 kDa, which corresponds
roughly to the size of the band detected by immunoblotting. Thus,
PBP1-FLAG can be specifically detected in membranes from
ponA-FLAG cells by using the monoclonal anti-FLAG antibody.
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FIG. 1.
Immunodetection and penicillin-binding activity of
PBP1-FLAG. (A) Western blot of PBP1-FLAG. Membranes from cells grown at
37°C in 100 ml of 2× YT medium were prepared and proteins were
solubilized in SDS sample buffer and subjected to SDS-10% PAGE as
described in Materials and Methods; about 40 µg of protein was loaded
into each well of the gel. The gel was transferred to a polyvinylidene
difluoride membrane, which was probed with 4 µg of monoclonal
anti-FLAG antibody (Sigma) per ml, and bound antibodies were visualized
by using alkaline phosphatase-conjugated secondary antibodies and a
colorimetric alkaline phosphatase substrate as described in Materials
and Methods. Lanes: 1, membrane proteins from strain LP27 expressing
PBP1-FLAG; 2, cytoplasmic proteins from strain LP27; and 3, membrane
proteins from wild-type cells (PS832). Molecular mass markers are shown
in kilodaltons on the left. The arrow indicates the position of
PBP1-FLAG. (B) Penicillin-binding activity of PBP1-FLAG. Membranes were
prepared as described for panel A and incubated with FLU-C6-APA for 30 min at 30°C as described (46), proteins (60 µg per well)
were separated by SDS-10% PAGE, and PBPs were visualized with a
fluorimager. Lanes: 1, membrane proteins from strain LP27 expressing
PBP1-FLAG; and 2, membrane proteins from wild-type cells (PS832). The
positions of the major vegetative B. subtilis PBPs are shown
on the left. The arrow indicates the position of PBP1 and PBP1-FLAG
(due to the large size of PBP1, we could not observe any size
difference between PBP1 and PBP1-FLAG).
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PBP1-FLAG is functionally indistinguishable from PBP1.
To
determine whether PBP1 function is affected by the addition of the FLAG
epitope, we first tested whether PBP1-FLAG is able to bind penicillin.
Membranes from log-phase cultures of wild-type and ponA-FLAG
cells grown at 37°C in 2× YT medium were incubated with FLU-C6-APA,
proteins separated by SDS-10% PAGE and PBPs visualized on a
fluorimager. The results indicated that PBP1-FLAG is capable of binding
penicillin like wild-type PBP1 (Fig. 1B), suggesting that tagging with
FLAG at the C terminus of PBP1 does not significantly affect the
penicillin-binding activity of the protein. Recent experiments
conducted in our laboratory have shown that ponA mutant cells require increased levels of divalent cations in the growth medium
and are therefore unable to grow in Penassay broth, which is relatively
low in Mg2+ (35). We therefore compared the
growth rate of ponA-FLAG cells grown in Penassay broth at
37°C to those of wild-type cells and cells of a ponA
mutant strain (PS2062) lacking PBP1 (45). Consistent with
previous findings (35), the ponA mutant was
unable to grow in Penassay broth, while both the ponA-FLAG
and wild-type cells grew with a doubling time of about 20 min. In
addition, fluorescence microscopy of cells grown at 37°C in 2× YT
medium revealed that ponA-FLAG cells are morphologically
identical to wild-type cells (data not shown, and see below), while
ponA mutant cells are much longer and thinner
(46). These findings indicate that PBP1-FLAG is functionally
equivalent to wild-type PBP1 in vivo, and it is therefore very unlikely
that the FLAG epitope interferes with PBP1 localization.
Number of PBP1 molecules per cell.
To date one of the
least-abundant bacterial proteins that has been localized by
immunofluorescence microscopy is FtsI from E. coli
(60), which is present at about 100 molecules per cell (13, 60). To determine whether it would be feasible to
localize PBP1-FLAG in B. subtilis by immunofluorescence
microscopy, the cellular abundance of this protein was analyzed.
Membranes from ponA-FLAG cells grown at 30 or 37°C in 2×
YT medium were prepared and subjected to SDS-10% PAGE followed by
immunoblotting with the anti-FLAG antibody. The blots were scanned, and
the total amount of FLAG epitope in each sample was determined by using C-terminal FLAG-BAP as a standard. An example of this analysis is shown
in Fig. 2. The DNA content of the samples
used for the membrane preparations was determined and used to calculate
the amount of PBP1-FLAG molecules per DNA equivalent in the samples. Assuming that FLAG-BAP and PBP1-FLAG react equally well with the anti-FLAG antibody, and by using a molecular weight for the B. subtilis chromosome of 2,781.77 × 106
(28) and a molecular weight of FLAG-BAP of 49,100, the
numbers of PBP1 molecules per chromosome equivalent were estimated to be 154 ± 33 for cells grown at 30°C with a doubling time of 32 min and 231 ± 86 for cells grown at 37°C with a doubling time of 20 min (three independent experiments). B. subtilis cells
grown in media with doubling times of 40 or 30 min contain about 2.9 or
3.3 chromosomes per cell, respectively (51). Hence, we
estimate that there are between 450 and 1,000 molecules of PBP1 per
cell in our experiments. Using a method based on in vivo labeling of PBPs with [3H]benzylpenicillin, the numbers of molecules
of PBP1a and PBP1b per E. coli cell have been estimated to
be 221 and 127, respectively, for cells grown at 35°C in
Luria-Bertani medium (13).
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FIG. 2.
Quantitative Western blotting of PBP1-FLAG. Cells of
strain LP27 were grown in 50 ml of 2× YT medium at 37°C, harvested,
and broken by sonication and the membrane and cytoplasmic fractions
were separated by high-speed centrifugation as described in Materials
and Methods. Membranes were resuspended in 90 µl of buffer B (50 mM
Tris-HCl [pH 8], 1 mM -ME, 1 mM PMSF), and 5, 3, or 2 µl was
brought to a 10-µl volume with buffer B, mixed with 10 µl of 2×
SDS sample buffer, boiled for 4 min, and subjected to SDS-10% PAGE
and immunoblotting with anti-FLAG antibody as described in the legend
for Fig. 1A. The blot was scanned, and the relative amounts of FLAG
molecules in the samples were determined by comparison with known
amounts (range, 9.4 to 150 ng) of purified FLAG-BAP protein (Sigma),
which were run on the same gel. The number of PBP1-FLAG molecules per
cell was calculated as described in the text. The inset in the graph
shows the blot with the different amounts of FLAG-BAP (lower band) and
PBP1-FLAG (upper band). Open squares in the graph are values for
FLAG-BAP standard protein; black circles are values for the
different amounts of membranes from ponA-FLAG mutant cells
loaded on the gel.
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PBP1 localizes to division sites.
We used log-phase
ponA-FLAG cells to determine the subcellular localization of
PBP1 by immunofluorescence microscopy. Briefly, cells were fixed,
permeabilized, applied to poly-L-lysine-coated microscope
slides, and incubated with anti-FLAG antibody followed by incubation
with biotinylated secondary antibodies and streptavidin conjugated to
the red fluorophore Cy3 (Jackson ImmunoResearch). The cells were also
stained with WGA conjugated to the fluorophore Oregon green (Molecular
Probes) to allow visualization of cell walls and septa. Cells from a
log-phase culture of the wild-type strain (PS832) were subjected to the
same treatments to serve as a negative control. Fluorescence microscopy
of immunostained ponA-FLAG cells revealed the presence of
bright red bands of fluorescence at division sites (Fig.
3A, B, and C), which were absent in
wild-type cells analyzed in parallel (Fig. 3D). We also found no such
fluorescent bands in vegetative cells of a strain expressing C-terminal
FLAG-tagged GerBA (a putative membrane protein that is normally
expressed in the forespore during sporulation and hence has no role in
cell division [12]) from a xylose-inducible promoter
(39), ruling out the possibility that the targeting of
PBP1-FLAG to the division site is due only to the FLAG epitope. About
69% of the ponA-FLAG cells examined (n = 182) showed ponA-FLAG localization at midcell (Fig. 3A,
B, and C), while the remaining 31% had no midcell bands. In addition,
some fluorescent bands (17% of 151 fluorescent bands observed) were
found to colocalize with septa separating two cells in a chain (Fig.
3A, B, and C). Presumably these bands represent PBP1 that was left over
from a previous division event. In some cells, the red fluorescence
observed was a bright dot rather than a band. Such dots could reflect
PBP1 localization or may be a fixation artifact. In numerous
experiments we found that if cells are not fixed sufficiently, bright
dots could often be observed at old poles, at new poles, and at what
appear to be future cell division sites (i.e., at midcell and at
one-fourth and three-fourths of the cell's length). Initially we
thought that these dots reflected PBP1 localization, and it has become
apparent to us that extreme caution must be taken when interpreting
data from such immunofluorescence experiments. In general, we found
that cells that are not properly fixed have a high background
fluorescence and look very smudgy compared with cells that are properly
fixed. Other authors have previously reported that the presence of dots
at the cell pole is a common artifact associated with
immunofluorescence microscopy of bacteria (60). Taken
together, the results suggest that PBP1 localizes to midcell around the
time that the cell is about to divide and disappears from this site
shortly after completion of the septum.
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FIG. 3.
Immunofluorescence of ponA-FLAG (A, B, and C)
or wild-type (D) cells grown in 2× YT medium at 37°C. The figure
shows immunodetection of the FLAG epitope only (red), staining of cell
walls and septa with Oregon green-conjugated WGA (green), and an
overlay of those images showing simultaneously visualized PBP1-FLAG and
cell walls and septa. Arrows point to PBP1-FLAG localization at
midcell; arrowheads show PBP1-FLAG bands that colocalize with completed
septa between two cells in a chain (completed septa stain very brightly
with Oregon green-WGA, in contrast to nascent septa at midcell, which
appear as faint green bands; see also Materials and Methods). Bar, 10 µm.
|
|
Impaired septum formation in a mutant lacking PBP1.
The
evidence that PBP1 is localized predominantly to the site of cell
division suggests that this protein might have some important role in
septum formation, and recent results acquired in our laboratory suggest
that PBP1 is required for septum formation in cells grown in media low
in Mg2+ (35). It has previously been shown that
in 2× SG medium, ponA mutant cells are longer and grow more
slowly than wild-type cells; in 2× YT and minimal medium, growth of
ponA mutant cells is also significantly slower than that of
wild-type cells (46). Given that PBP1 localizes to division
sites in cells grown in 2× YT medium, we speculated that PBP1 may also
be needed for septum formation under these normal growth conditions.
Indeed, fluorescence microscopy of ponA mutant cells grown
in 2× YT medium at 30°C and double stained with Oregon green-WGA and
DAPI to visualize cell walls, septa, and nucleoids showed that
ponA mutant cells not only are longer than wild-type cells
(mean cell length was 6.4 µm for ponA mutant cells
[n = 98] and 3.9 µm for wild-type cells
[n = 137]), consistent with a previous report
(46), but also contain a larger average number of nucleoids
per cell (2.5; n = 98) than wild-type cells (1.8;
n = 137), suggesting a reduced efficiency of septation
of the ponA mutant (Fig. 4 and
5). Similar results were obtained when
cells were grown at 37°C (results not shown). Interestingly, bright
green dots could often be observed at the periphery of Oregon
green-WGA-stained ponA mutant cells grown in 2× YT medium
at 30°C (39% of the cells; n = 163) (Fig. 4A) or at
37°C (39%; n = 222), suggesting that at these sites septation was initiated but failed to go to completion. The proportion of ponA mutant cells with bright green dots was lower, but
still significant, when the cells were grown in 2× YT medium to which 1 mM or 10 mM MgCl2 had been added (23% of the cells
[n = 279] grown at 37°C in 2× YT with 1 mM
MgCl2 had bright green dots, while with 10 mM
MgCl2, the number was 25% [n = 282]),
suggesting that the lack of PBP1 results in a significant septation
defect even under these conditions. Based on electron microscopy data (Fig. 6, and see below) we estimate that
about 50% of the cell wall dots were located between well-segregated
nucleoids while the remaining dots were located at positions
overlapping the DNA. No cell wall dots were observed in wild-type cells
(n = 339) grown at 37°C in 2× YT medium, indicating
that the observed septation defect was indeed due to the
ponA mutation.
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FIG. 4.
Fluorescence micrographs showing filamentation in a
ponA mutant (strain PS2062) grown at 30°C in 2× YT
medium. (A and B) ponA mutant cells; (C) wild-type cells.
(Ai, Bi, and Ci) DAPI staining of nucleoids (blue); (Aii, Bii, and Cii)
staining of cell walls and septa with Oregon green-conjugated WGA
(green); (Aiii, Biii, and Ciii) overlay of DAPI staining and cell wall
staining. Bar, 10 µm. Arrows in panels Aii and Aiii show a bright
green dot in the periphery of ponA mutant cells between two
nucleoids (arrow in Ai). Such dots were seen in 39% of the
ponA mutant cells examined (n = 163) but
were not seen in wild-type cells (0%; n = 137).
Similar results were obtained when cells were grown in 2× YT medium at
37°C (39% of the ponA mutant cells had dots
[n = 222]; 0% of wild-type cells had dots
[n = 339]).
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|
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FIG. 5.
Cell length and number of nucleoids for ponA
mutant (A) and wild-type (B) cells grown at 30°C in 2× YT medium.
Images similar to those shown in Fig. 4 were used for the analysis.
Cell lengths were measured as described in Materials and Methods, and
the number of visible, fully segregated nucleoids present in each cell
was counted. A total of 98 and 137 cells are depicted in panels A and
B, respectively. The mean cell length and nucleoid content obtained
were, respectively, 6.4 µm and 2.5 for ponA mutant cells
and 3.9 µm and 1.8 for wild-type cells.
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|
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FIG. 6.
Electron micrograph showing cross-sections of
ponA mutant and wild-type cells grown at 37°C in 2× YT
medium. (A and B) ponA mutant cells; (C) wild-type cell. The
arrowheads indicate aberrant septa or cell wall clusters in
ponA mutant cells. Note that the cell wall clusters shown in
panel A are at positions overlapping the DNA. Bar, 0.5 µm.
|
|
To investigate the septation defect of the
ponA mutant in
more detail, sections of
ponA mutant and wild-type cells
were prepared
and examined by electron microscopy (Fig.
6). This
analysis showed
that while most dividing
ponA mutant cells
(73%;
n = 103) had
septa that looked like those of
wild-type cells (Fig.
6C), a significant
proportion (27%;
n = 103) had abnormal clusters of cell wall material
assembled at
distinct sites at the cell periphery (Fig.
6A) or
displayed incomplete
septa with aberrant morphology (Fig.
6B).
Presumably these abnormal
septa or wall clusters correspond to
the fluorescent dots seen by
fluorescence microscopy of
ponA mutant
cells (Fig.
4A, and
see above), and this observation further supports
a role for PBP1 in
septal peptidoglycan synthesis. However, it
cannot be ruled out that
these abnormal septa or cell wall clusters
are a secondary effect of
the reduced diameter and/or general
changes in the cell wall structure
of the
ponA mutant (
46).
FtsZ localization in cells lacking PBP1.
An essential and very
early event in cell division is the assembly and polymerization of FtsZ
into a cytokinetic ring at midcell (9, 48). Given the effect
of the ponA mutation on septum formation, we were interested
in analyzing the effect of this mutation on FtsZ ring formation.
Log-phase cultures of ponA mutant and wild-type cells were
fixed and subjected to immunofluorescence microscopy by using a rabbit
antiserum against B. subtilis FtsZ followed by staining with
Cy3-conjugated secondary antibodies as well as DAPI and FITC-conjugated
WGA to visualize nucleoids, septa, and cell walls. Examination of the
immunostained cells showed that the ponA mutation had a
significant but very pleiotropic effect on FtsZ localization. Some
ponA mutant cells had normal FtsZ localization, which was
seen as a bright ring-like structure at the midpoint of the cell with
one or two well-segregated nucleoids on each side (Fig.
7B, leftmost cell). Others had two
closely positioned FtsZ rings at anticipated division sites, which were not always perpendicular to the long axis of the cell (Fig. 7A). Cells
with normal-appearing FtsZ rings at inappropriate sites, such as an
acentric ring in a cell with three or more nucleoids (Fig. 7B), as well
as cells with no visible FtsZ rings, were also observed. In some of the
latter cells, a diffuse or fuzzy signal between two separated nucleoids
was sometimes detected, suggestive of aggregated or disassembled FtsZ
protein. To compare FtsZ localization in ponA mutant and
wild-type cells, immunostained cells were scored for FtsZ localization
by using the following criteria: (i) normal FtsZ rings, which were
defined as a nice ring-like localization at the midpoint of the cell
with one or two separated nucleoids on each side; (ii) abnormal FtsZ
rings, which were either double rings or FtsZ rings at inappropriate
sites; and (iii) no FtsZ rings, a category used when cells had one or
two nucleoids and no FtsZ rings. This analysis showed that most
ponA mutant cells (52%; n = 507) had normal
FtsZ rings and a significant proportion (22%; n = 507)
had abnormal FtsZ rings, while the remaining 26% (n = 507) had no FtsZ rings. In contrast, 82% (n = 425) of the wild-type cells had normal FtsZ rings and 3%
(n = 425) had abnormal rings, while 15% (n = 425) showed no FtsZ rings (Table
2). These results therefore demonstrate
that the ponA mutation has a significant, but very
pleiotropic, effect on FtsZ localization, presumably because the
effects of the ponA mutation itself are very pleiotropic (35, 46).
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FIG. 7.
Immunolocalization of FtsZ in ponA mutant (A
and B) and wild-type (C) cells grown in 2× YT medium at 37°C. (Ai,
Bi, and Ci) Overlay of FtsZ localization (red and yellow), DAPI
staining of nucleoids (blue), and staining of cell walls and septa with
FITC-conjugated WGA (green); (Aii, Bii, and Cii) overlay of FtsZ
localization (red) and DAPI staining (blue). The arrows in panels Ai
and Aii indicate an aberrant FtsZ ring, and those in panels Bi and Bii
indicate an FtsZ ring at an inappropriate location. Bar, 10 µm.
|
|
| |
DISCUSSION |
The main contributions of the present work can be summarized as
follows: (i) PBP1 localizes to the division site in B. subtilis cells, (ii) a significant proportion of ponA
mutant cells grown under normal growth conditions are filamentous or
have a septation defect, and (iii) while FtsZ localizes correctly in
most ponA mutant cells grown under normal conditions, a
significant proportion of ponA mutant cells contain FtsZ
rings that have aberrant morphology or are localized at inappropriate sites.
Role of PBP1 in septum formation.
The localization of PBP1 to
the division site strongly suggests that PBP1 is directly involved in
septal peptidoglycan synthesis. The only other PBP that has been
directly implicated in septum synthesis is the class B HMW PBP2b, which
is homologous to E. coli FtsI (62). Given that
class B HMW PBPs have no transglycosylase activity (1, 17,
57), it is likely that PBP2b and PBP1 cooperate during septal
peptidoglycan synthesis in B. subtilis, as has been proposed
for E. coli FtsI and PBP1a and PBP1b (61). However, in contrast to PBP2b, which is an essential protein
(62), inactivation of ponA is not lethal under
normal growth conditions (35, 45, 46). This suggests the
presence in B. subtilis of another protein with
transglycosylase activity involved in septum synthesis. Possible
candidates for such a protein are the two other known B. subtilis HMW class A PBPs, PBP2c and PBP4, because previous
studies have shown that the growth and morphology defects associated
with a ponA insertional mutation are exacerbated greatly by
the additional loss of PBP4 and only slightly by the loss of PBP2c
(46). However, cells lacking PBP4, PBP2c, or both do not
display any significant growth defects under normal growth conditions
(43, 44, 46), suggesting that any contribution of either of
these proteins to septal wall synthesis is clearly dispensable when
PBP1 is present. Furthermore, a mutant lacking the PBPs PBP1, PBP2c,
and PBP4 is viable, although its growth is seriously impaired
(46), suggesting the involvement of yet another (unknown)
protein. Recently the B. subtilis genome sequencing project
identified ywhE as a gene encoding a putative class A HMW
PBP (28). We are now planning to generate a mutant that lacks all four class A HMW PBP-encoding genes to determine whether such
a mutant is viable. In this respect it is worth noting that inactivation of PBP1a and PBP1b in E. coli is lethal
(63), despite the presence of a third class A HMW PBP in
this organism (23) as well as a putative monofunctional
peptidoglycan transglycosylase (54). Inactivation of a
single ponA homolog in Neisseria gonorrhoeae is
also lethal (47), again despite the presence of a putative monofunctional peptidoglycan transglycosylase in this organism (54).
Septal peptidoglycan synthesis is not the only function of
PBP1.
Previous studies have shown that inactivation of
ponA produces cells that are not only longer than wild-type
cells but also thinner and more bent (46), suggesting a role
for PBP1 in lateral wall synthesis. In contrast, B. subtilis
cells containing a C-terminal truncated version of PBP2b, a protein
presumed to function only in cell division, grow as filaments, but
their cell diameter was not significantly different from that of
wild-type cells (62). In addition, our results do not
exclude the possibility that PBP1 is present in the membrane lining the
cylindrical parts of the cell wall, because the local concentration of
PBP1 at such sites may be too low to be detected. Taken together, the
results make it seem likely that during cell elongation, PBP1 is
involved in cylindrical cell wall synthesis and is present at fairly
low local concentrations at a number of sites in the membrane lining
the cylinder. Subsequently, upon initiation of cell division, PBP1 localizes to the midpoint of the cell by some unknown mechanism, becomes part of the divisome complex (37), and promotes the formation of septal peptidoglycan, possibly by synthesizing primers which are subsequently cross-linked by PBP2b.
Timing of PBP1 localization.
One of the earliest events in
septation in E. coli is the polymerization of the
tubulin-like FtsZ protein into a cytokinetic ring at midcell, which
contracts during the septation process and maintains a position at the
leading edge of the invaginating septum (9, 48). Formation
of the FtsZ ring occurs in the absence of functional FtsA, FtsQ, FtsI,
FtsK, or FtsW (2, 4, 27, 64). Conversely, septal
localization of FtsA and FtsK is FtsZ-dependent (4, 64) and
FtsI localization depends on FtsZ, FtsA, FtsL, and FtsQ (58,
59), while FtsN localization depends on prior FtsZ and FtsA
localization and requires the function of FtsI and FtsQ (3).
In most cases, these localization studies are indicative of the stage
in septation during which a given protein acts. For example,
ftsZ mutant cell filaments have a smooth morphology,
indicating a role for FtsZ very early in the division process (8,
56), while ftsI mutant cell filaments have
indentations at septal positions suggesting a role for FtsI in the
continuation rather than initiation of septum formation (36,
38). Inactivation of PBP1a and PBP1b with the antibiotics
cefsulodin and moenomycin, which inhibit the transpeptidase activity
and transglycosylase activity, respectively, does not prevent
initiation of constriction during cell division in E. coli
(36, 61), suggesting that PBP1a and PBP1b, like FtsI, are
not required for initiation of septum formation in E. coli.
In the present study we found that 17% of 151 fluorescent PBP1 bands
observed colocalized with completed septa between two
cells in a chain
(Fig.
3A, B, and C). In addition, in the majority
of cells with midcell
PBP1 fluorescence (70% of 126 cells), septal
peptidoglycan was also
visible at these positions as judged by
the presence of faint green
bands upon staining with Oregon green-WGA.
This is in contrast to FtsZ,
which was not observed to colocalize
with septa visualized by FITC-WGA
staining (Fig.
7 and data not
shown), suggesting that PBP1 localizes to
the division site after
assembly of the FtsZ ring and that PBP1
probably acts at a relatively
late stage in septation. In support of
this idea, immunofluorescence
and electron microscopy of
B. subtilis ponA mutant cells showed
that a significant proportion of
the cells (27 to 39%) had abnormal
septa or clusters of cell wall
material at the cell periphery,
indicating that in such cells, septum
formation had initiated
but failed to complete. Furthermore, by
immunofluorescence microscopy
of
ponA mutant cells we found
that FtsZ rings formed normally
in the majority of the cells (56%;
n = 507), indicating that localization
of FtsZ to the
division site does not require prior localization
of PBP1. However, we
also found that a significant proportion
of the
ponA mutant
cells (22%;
n = 507) contained FtsZ rings that
had
aberrant morphology or were present at inappropriate locations,
indicating that the lack of PBP1 somehow interferes with the stability
or assembly of the FtsZ ring. That the effect of the
ponA
mutation
on FtsZ localization is so pleiotropic is not surprising,
given
that PBP1 is involved not just in cell division but also in
lateral
cell wall synthesis and maintenance of the correct cell
diameter,
as discussed above. Consequently, we find it difficult to
explain
precisely how the
ponA mutation affects FtsZ
localization, but
there are several possibilities: (i) the reduction of
the diameter
of the cell could affect FtsZ ring assembly or stability;
(ii)
some general change in the cell wall structure may alter
interactions
between the cell wall and the membrane, which in turn may
interfere
with the binding of FtsZ to its membrane nucleation site;
(iii)
changes in septal peptidoglycan structure or interactions between
proteins at the division site could interfere with the stability
of the
FtsZ ring; and (iv) effects on cell growth might interfere
with general
cues for cell division. However, we think it is important
to stress
that the majority of the
ponA mutant cells had normal
FtsZ
localization, indicating that PBP1 is not completely essential
for the
localization and binding of FtsZ at the division
site.
Interestingly, some of the FtsZ localization patterns observed in the
ponA mutant are reminiscent of those reported in a study
by
Addinall and colleagues (
2) for
E. coli filaments
with mutations
in the
ftsA,
ftsQ, or
ftsI gene. These authors found that a large
proportion of
such filaments had more than two FtsZ rings, some
had indentations or
FtsZ rings with variable spacing, and some
filaments had no FtsZ rings,
and they suggested that blocking
of septal peptidoglycan synthesis
blocks progression of the FtsZ
ring and may thus lead to its
destabilization (
2). In a similar
study it was found that
inactivation of FtsI by temperature or
cephalexin also had very
pleiotropic effects on FtsZ localization,
and it was shown that
inactivation of FtsI inhibits constriction
of the FtsZ ring and delays
the assembly of FtsZ rings at future
division sites, resulting in a
lower overall average number of
FtsZ rings per cell under these
conditions (
40). It is possible
that the lack of PBP1 at the
septum in
B. subtilis affects FtsZ
ring formation by a
similar
mechanism.
Recently it was shown by fluorescence microscopy that
E. coli cells which lack a protein required for the synthesis of the
major membrane lipid component phosphatidylethanolamine (PE) have
a
defect in cell division and contain FtsZ rings with aberrant
structure
at potential division sites (reference
33 and
references
therein). For example, some PE-deficient filaments were
found
to contain FtsZ bands that were not perpendicular to the long
axis of the cell, a number of very long filaments only had a few
FtsZ
rings, and others had regularly spaced FtsZ rings with normal
morphology (
33). One suggested explanation for these
findings
is that changes in the phospholipid composition of the cell
membrane
may affect FtsZ ring formation indirectly by altering the
interaction
between FtsZ and its membrane nucleation site
(
33). Given the
effect of the
ponA mutation on
overall cell wall structure and
cell dimensions, the cell membrane
structure may also be affected
and consequently the inactivation of
ponA may affect FtsZ ring
formation by a mechanism similar
to that of PE
deficiency.
| |
ACKNOWLEDGMENTS |
We thank David L. Popham for the gift of plasmid pDPC273 and for
advice regarding epitope tagging of PBP1, Petra A. Levin for the gift
of anti-FtsZ antibodies and for helpful discussions, Kit Pogliano and
Ann Cowan for helpful advice regarding immunofluorescence microscopy,
Arthur L. Hand for performing electron microscopy, Frank Morgan and
Susan Krueger for assistance with computer work and image analysis,
Richard Losick for generous support and use of his equipment, Madan
Paidhungat for the Pxyl-GerBA-FLAG strain, and Lawrence I. Rothfield for helpful discussions and comments on the manuscript.
This work was supported by a grant from the National Institutes of
Health to P.S. (GM19698), a postdoctoral fellowship from the Danish
Natural Science Research Council to L.B.P. (9601026), and a
postdoctoral fellowship to E.R.A. from the Jane Coffin Childs Memorial
Fund for Medical Research.
| |
FOOTNOTES |
*
Corresponding author. Mailing address: Department of
Biochemistry, University of Connecticut Health Center, 263 Farmington Ave., Farmington, CT 06032. Phone: (860) 679-2607. Fax: (860) 679-3408. E-mail: setlow{at}sun.uchc.edu.
| |
REFERENCES |
| 1.
|
Adam, M.,
C. Fraipont,
N. Rhazi,
M. Nguyen-Distèche,
B. Lakaye,
J.-M. Frère,
B. Devreese,
J. van Beeumen,
Y. van Heijenoort,
J. van Heijenoort, and J.-M. Ghuysen.
1997.
The bimodular G57-V577 polypeptide chain of the class B penicillin-binding protein 3 of Escherichia coli catalyzes peptide bond formation from thiolesters and does not catalyze glycan chain polymerization from the lipid II intermediate.
J. Bacteriol.
179:6005-6009[Abstract/Free Full Text].
|
| 2.
|
Addinall, S. G.,
E. Bi, and J. Lutkenhaus.
1996.
FtsZ ring formation in fts mutants.
J. Bacteriol.
178:3877-3884[Abstract/Free Full Text].
|
| 3.
|
Addinall, S. G.,
C. Cao, and J. Lutkenhaus.
1997.
FtsN, a late recruitment to the septum in Escherichia coli.
Mol. Microbiol.
25:303-309[Medline].
|
| 4.
|
Addinall, S. G., and J. Lutkenhaus.
1996.
FtsA is localized to the septum in an FtsZ-dependent manner.
J. Bacteriol.
178:7167-7172[Abstract/Free Full Text].
|
| 5.
|
Allen, K. A.,
A. Neuberger, and N. Sharon.
1973.
The purification, composition and specificity of wheat-germ agglutinin.
Biochem. J.
131:155-162[Medline].
|
| 6.
|
Anagnostopoulus, C., and J. Spizizen.
1961.
Requirements for transformation in Bacillus subtilis.
J. Bacteriol.
81:74-76.
|
| 7.
|
Bayer, M. H.,
W. Keck, and M. E. Bayer.
1990.
Localization of penicillin-binding protein 1b in Escherichia coli: immunoelectron microscopy and immunotransfer studies.
J. Bacteriol.
172:125-135[Abstract/Free Full Text].
|
| 8.
|
Begg, K. J., and W. D. Donachie.
1985.
Cell shape and division in Escherichia coli: experiments with shape and division mutants.
J. Bacteriol.
163:615-622[Abstract/Free Full Text].
|
| 9.
|
Bi, E., and J. Lutkenhaus.
1991.
FtsZ ring structure associated with division in Escherichia coli.
Nature
354:161-164[Medline].
|
| 10.
|
Bramhill, D.
1997.
Bacterial cell division.
Annu. Rev. Cell Dev. Biol.
13:395-424[Medline].
|
| 11.
|
Chen, J. C.,
D. S. Weiss,
J.-M. Ghigo, and J. Beckwith.
1999.
Septal localization of FtsQ, an essential cell division protein in Escherichia coli.
J. Bacteriol.
181:521-530[Abstract/Free Full Text].
|
| 12.
|
Corfe, B. M.,
A. Moir,
D. Popham, and P. Setlow.
1994.
Analysis of the expression and regulation of the gerB spore germination operon of Bacillus subtilis 168.
Microbiology
140:3079-3083[Abstract/Free Full Text].
|
| 13.
|
Dougherty, T. J.,
K. Kennedy,
R. E. Kessler, and M. J. Pucci.
1996.
Direct quantitation of the number of individual penicillin-binding proteins per cell in Escherichia coli.
J. Bacteriol.
178:6110-6115[Abstract/Free Full Text].
|
| 14.
|
Ferrari, F. A.,
A. Nguyen,
D. Lang, and J. A. Hoch.
1983.
Construction and properties of an integrable plasmid for Bacillus subtilis.
J. Bacteriol.
154:1513-1515[Abstract/Free Full Text].
|
| 15.
|
García del Portillo, F.,
M. A. de Pedro, and J. A. Ayala.
1991.
Identification of a new mutation in Escherichia coli that suppresses a pbpB (Ts) phenotype in the presence of penicillin-binding protein 1B.
FEMS Microbiol. Lett.
84:7-14.
|
| 16.
|
García del Portillo, F.,
M. A. de Pedro,
D. Joseleau-Petit, and R. d'Dari.
1989.
Lytic response of Escherichia coli cells to inhibitors of penicillin-binding proteins 1a and 1b as a timed event related to cell division.
J. Bacteriol.
171:4217-4221[Abstract/Free Full Text].
|
| 17.
|
Ghuysen, J.-M.
1994.
Molecular structures of penicillin-binding proteins and -lactamases.
Trends Microbiol.
2:372-380[Medline].
|
| 18.
|
Goffin, C.,
C. Fraipont,
J. Ayala,
M. Terrak,
M. Nguyen-Distèche, and J.-M. Ghuysen.
1996.
The non-penicillin-binding module of the tripartite penicillin-binding protein 3 of Escherichia coli is required for the folding and/or stability of the penicillin-binding module and the membrane-anchoring module confers cell septation activity on the folded structure.
J. Bacteriol.
178:5402-5409[Abstract/Free Full Text].
|
| 19.
|
Hale, C. A., and P. A. J. de Boer.
1997.
Direct binding of FtsZ to ZipA, an essential component of the septal ring structure that mediates cell division in E. coli.
Cell
88:175-185[Medline].
|
| 20.
|
Harry, E. J.,
K. Pogliano, and R. Losick.
1995.
Use of immunofluorescence to visualize cell-specific gene expression during sporulation in Bacillus subtilis.
J. Bacteriol.
177:3386-3393[Abstract/Free Full Text].
|
| 21.
|
Harry, E. J., and R. G. Wake.
1997.
The membrane-bound cell division protein DivIB is localized to the division site in Bacillus subtilis.
Mol. Microbiol.
25:275-283[Medline].
|
| 22.
|
Hiraga, S.,
C. Inchinose,
H. Niki, and M. Yamazoe.
1998.
Cell cycle-dependent duplication and bidirectional migration of SeqA-associated DNA-protein complexes in E. coli.
Mol. Cell
1:381-387[Medline].
|
| 23.
|
Höltje, J.-V.
1998.
Growth of the stress-bearing and shape-maintaining murein sacculus of Escherichia coli.
Microbiol. Mol. Biol. Rev.
62:181-203[Abstract/Free Full Text].
|
| 24.
|
Ishino, F.,
K. Mitsui,
S. Tamaki, and M. Matsuhashi.
1980.
Dual enzyme activities of cell wall peptidoglycan synthesis, peptidoglycan transglycosylase and penicillin-sensitive transpeptidase, in purified preparations of Escherichia coli penicillin-binding protein 1A.
Biochem. Biophys. Res. Commun.
97:287-293[Medline].
|
| 25.
|
Katis, V. L.,
E. J. Harry, and R. G. Wake.
1997.
The Bacillus subtilis division protein DivIC is a highly abundant membrane-bound protein that localizes to the division site.
Mol. Microbiol.
26:1047-1055[Medline].
|
| 26.
|
Kato, J.,
H. Suzuki, and Y. Hirota.
1985.
Dispensability of either penicillin-binding protein-1a or -1b involved in the essential process for cell elongation in Escherichia coli.
Mol. Gen. Genet.
200:272-277[Medline].
|
| 27.
|
Khattar, M. M.,
S. G. Addinall,
K. H. Stedul,
D. S. Boyle,
J. Lutkenhaus, and W. D. Donachie.
1997.
Two polypeptide products of the Escherichia coli cell division gene ftsW and a possible role for FtsW in FtsZ function.
J. Bacteriol.
179:784-793[Abstract/Free Full Text].
|
| 28.
|
Kunst, F.,
N. Ogasawara,
I. Moszer,
A. M. Albertini,
G. Alloni, et al.
1997.
The complete genome sequence of the gram-positive bacterium Bacillus subtilis.
Nature
390:249-256[Medline].
|
| 29.
|
Laemmli, U. K.
1970.
Cleavage of structural proteins during the assembly of the head of bacteriophage T4.
Nature
227:680-685[Medline].
|
| 30.
|
Leighton, T. J., and R. H. Doi.
1971.
The stability of messenger ribonucleic acid during sporulation in Bacillus subtilis.
J. Biol. Chem.
254:3189-3195.
|
| 31.
|
Levin, P. A., and R. Losick.
1996.
Transcription factor Spo0A switches the localization of the cell division protein FtsZ from a medial to a bipolar pattern in Bacillus subtilis.
Genes Dev.
10:478-488[Abstract/Free Full Text].
|
| 32.
|
Lowry, O. H.,
N. J. Rosebrough,
A. L. Farr, and R. J. Randall.
1951.
Protein measurement with the Folin phenol reagent.
J. Biol. Chem.
193:265-275[Free Full Text].
|
| 33.
|
Mileykovskaya, E.,
Q. Sun,
W. Margolin, and W. Dowhan.
1998.
Localization and function of early cell division proteins in filamentous Escherichia coli cells lacking phosphatidylethanolamine.
J. Bacteriol.
180:4252-4257[Abstract/Free Full Text].
|
| 34.
|
Murray, T.,
D. L. Popham, and P. Setlow.
1997.
Identification and characterization of pbpA encoding Bacillus subtilis penicillin-binding protein 2A.
J. Bacteriol.
179:3021-3029[Abstract/Free Full Text].
|
| 35.
|
Murray, T.,
D. L. Popham, and P. Setlow.
1998.
Bacillus subtilis cells lacking penicillin-binding protein 1 require increased levels of divalent cations for growth.
J. Bacteriol.
180:4555-4563[Abstract/Free Full Text].
|
| 36.
|
Nanninga, N.
1991.
Cell division and peptidoglycan assembly in Escherichia coli.
Mol. Microbiol.
5:791-795[Medline].
|
| 37.
|
Nanninga, N.
1998.
Morphogenesis of Escherichia coli.
Microbiol. Mol. Biol. Rev.
62:110-129[Abstract/Free Full Text].
|
| 38.
|
Nguyen-Distèche, M.,
C. Fraipont,
N. Buddelmeijer, and N. Nanninga.
1998.
The structure and function of Escherichia coli penicillin-binding protein 3.
Cell. Mol. Life Sci.
54:309-316[Medline].
|
| 38a.
| NIH Image program. National Institutes of Health,
Bethesda, Md. http://rsb.info.gov/nih-image. [9 September 1998, last
date accessed.]
|
| 39.
| Paidhungat, M., L. B. Pedersen, and P. Setlow.
1999. Unpublished results.
|
| 40.
|
Pogliano, J.,
K. Pogliano,
D. S. Weiss,
R. Losick, and J. Beckwith.
1997.
Inactivation of FtsI inhibits constriction of the FtsZ cytokinetic ring and delays the assembly of FtsZ rings at potential division sites.
Proc. Natl. Acad. Sci. USA
94:559-564[Abstract/Free Full Text].
|
| 41.
|
Pogliano, K.,
E. Harry, and R. Losick.
1995.
Visualization of the subcellular location of sporulation proteins in Bacillus subtilis using immunofluorescence microscopy.
Mol. Microbiol.
18:459-470[Medline].
|
| 42.
|
Pogliano, K.,
A. E. M. Hofmeister, and R. Losick.
1997.
Disappearance of the  E transcription factor from the forespore and the SpoIIE phosphatase from the mother cell contributes to establishment of cell-specific gene expression during sporulation in Bacillus subtilis.
J. Bacteriol.
179:3331-3341[Abstract/Free Full Text].
|
| 43.
|
Popham, D. L., and P. Setlow.
1993.
Cloning, nucleotide sequence, and regulation of the Bacillus subtilis pbpF gene, which codes for a putative class A high-molecular-weight penicillin-binding protein.
J. Bacteriol.
175:4870-4876[Abstract/Free Full Text].
|
| 44.
|
Popham, D. L., and P. Setlow.
1994.
Cloning, nucleotide sequence, mutagenesis, and mapping of the Bacillus subtilis pbpD gene, which codes for penicillin-binding protein 4.
J. Bacteriol.
176:7197-7205[Abstract/Free Full Text].
|
| 45.
|
Popham, D. L., and P. Setlow.
1995.
Cloning, nucleotide sequence, and mutagenesis of the Bacillus subtilis ponA operon, which codes for penicillin-binding protein (PBP) 1 and a PBP-related factor.
J. Bacteriol.
177:326-335[Abstract/Free Full Text].
|
| 46.
|
Popham, D. L., and P. Setlow.
1996.
Phenotypes of Bacillus subtilis mutants lacking multiple class A high-molecular-weight penicillin-binding proteins.
J. Bacteriol.
178:2079-2085[Abstract/Free Full Text].
|
| 47.
|
Ropp, P. A., and R. A. Nicholas.
1997.
Cloning and characterization of the ponA gene encoding penicillin-binding protein 1 from Neisseria gonorrhoeae and Neisseria meningitidis.
J. Bacteriol.
179:2783-2787[Abstract/Free Full Text].
|
| 48.
|
Rothfield, L. I., and S. S. Justice.
1997.
Bacterial cell division: the cycle of the ring.
Cell
88:581-584[Medline].
|
| 49.
|
Sarkar, G., and S. S. Sommer.
1990.
The "megaprimer" method of site-directed mutagenesis.
BioTechniques
8:404-407[Medline].
|
| 50.
|
Setlow, B.,
A. R. Hand, and P. Setlow.
1991.
Synthesis of a Bacillus subtilis small, acid-soluble spore protein in Escherichia coli causes cell DNA to assume some characteristics of spore DNA.
J. Bacteriol.
173:1642-1653[Abstract/Free Full Text].
|
| 51.
|
Sharpe, M. E.,
P. M. Hauser,
R. G. Sharpe, and J. Errington.
1998.
Bacillus subtilis cell cycle as studied by fluorescence microscopy: constancy of cell length at initiation of DNA replication and evidence for active nucleoid partitioning.
J. Bacteriol.
180:547-555[Abstract/Free Full Text].
|
| 52.
|
Sizemore, R. K.,
J. J. Caldwell, and A. S. Kendrick.
1990.
Alternate gram staining technique using a fluorescent lectin.
Appl. Environ. Microbiol.
56:2245-2247[Abstract/Free Full Text].
|
| 53.
|
Spratt, B. G., and A. B. Pardee.
1975.
Penicillin-binding proteins and cell shape in E. coli.
Nature
254:516-517[Medline].
|
| 54.
|
Spratt, B. G.,
J. Zhou,
M. Taylor, and M. J. Merrick.
1996.
Monofunctional biosynthetic peptidoglycan transglycosylases.
Mol. Microbiol.
19:639-647[Medline].
|
| 55.
|
Suzuki, H.,
Y. van Heijenoort,
T. Tamura,
J. Mizoguchi,
Y. Hirota, and J. van Heijenoort.
1980.
In vitro peptidoglycan polymerization catalyzed by penicillin binding protein 1b of Escherichia coli K12.
FEBS Lett.
110:245-249[Medline].
|
| 56.
|
Taschner, P. E. M.,
P. G. Huls,
E. Pas, and C. L. Woldringh.
1988.
Division behavior and shape changes in isogenic ftsZ, ftsQ, ftsA, pbpB, and ftsE cell division mutants of Escherichia coli during temperature shift experiments.
J. Bacteriol.
170:1533-1540[Abstract/Free Full Text].
|
| 57.
|
van Heijenoort, Y.,
M. Gómez,
M. Derrien,
J. Ayala, and J. van Heijenoort.
1992.
Membrane intermediates in the peptidoglycan metabolism of Escherichia coli: possible roles of PBP1b and PBP3.
J. Bacteriol.
174:3549-3557[Abstract/Free Full Text].
|
| 58.
|
Wang, L.,
M. K. Khattar,
W. D. Donachie, and J. Lutkenhaus.
1998.
FtsI and FtsW are localized to the septum in Escherichia coli.
J. Bacteriol.
180:2810-2816[Abstract/Free Full Text].
|
| 59.
|
Weiss, D. S.,
J. C. Chen,
J.-M. Ghigo,
D. Boyd, and J. Beckwith.
1999.
Localization of FtsI (PBP3) to the septal ring requires its membrane anchor, the Z ring, FtsA, FtsQ, and FtsL.
J. Bacteriol.
181:508-520[Abstract/Free Full Text].
|
| 60.
|
Weiss, D. S.,
K. Pogliano,
M. Carson,
L.-M. Guzman,
C. Fraipont,
M. Nguyen-Distèche,
R. Losick, and J. Beckwith.
1997.
Localization of the Escherichia coli cell division protein FtsI (PBP3) to the division site and cell pole.
Mol. Microbiol.
25:671-681[Medline].
|
| 61.
|
Wientjes, F. B., and N. Nanninga.
1991.
On the role of high molecular weight penicillin-binding proteins in the cell cycle of Escherichia coli.
Res. Microbiol.
142:333-344[Medline].
|
| 62.
|
Yanouri, A.,
R. A. Daniel,
J. Errington, and C. E. Buchanan.
1993.
Cloning and sequencing of the cell division gene pbpB, which encodes penicillin-binding protein 2B in Bacillus subtilis.
J. Bacteriol.
175:7604-7616[Abstract/Free Full Text].
|
| 63.
|
Yousif, S. Y.,
J. K. Broome-Smith, and B. G. Spratt.
1985.
Lysis of Escherichia coli by -lactam antibiotics: deletion analysis of the role of penicillin-binding proteins 1A and 1B.
J. Gen. Microbiol.
131:2839-2845[Abstract/Free Full Text].
|
| 64.
|
Yu, X.-C.,
A. H. Tran,
Q. Sun, and W. Margolin.
1998.
Localization of cell division protein FtsK to the Escherichia coli septum and identification of a potential N-terminal targeting domain.
J. Bacteriol.
180:1296-1304[Abstract/Free Full Text].
|
Journal of Bacteriology, May 1999, p. 3201-3211, Vol. 181, No. 10
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Abstract
Introduction
