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Journal of Bacteriology, July 2001, p. 4167-4175, Vol. 183, No. 14
Department of Biological Sciences, University
of Wisconsin-Milwaukee, Milwaukee, Wisconsin 53201
Received 4 January 2001/Accepted 24 April 2001
Cells of Flavobacterium johnsoniae move over
surfaces by a process known as gliding motility. The mechanism of this
form of motility is not known. Cells of F. johnsoniae
propel latex spheres along their surfaces, which is thought to be a
manifestation of the motility machinery. Three of the genes that are
required for F. johnsoniae gliding motility,
gldA, gldB, and ftsX, have
recently been described. Tn4351 mutagenesis was used to
identify another gene, gldD, that is needed for gliding.
Tn4351-induced gldD mutants formed
nonspreading colonies, and cells failed to glide. They also lacked the
ability to propel latex spheres and were resistant to
bacteriophages that infect wild-type cells. Introduction of wild-type gldD into the mutants restored motility,
ability to propel latex spheres, and sensitivity to bacteriophage
infection. gldD codes for a cytoplasmic membrane protein
that does not exhibit strong sequence similarity to proteins of known
function. gldE, which lies immediately upstream of
gldD, encodes another cytoplasmic membrane protein that
may be involved in gliding motility. Overexpression of
gldE partially suppressed the motility defects of a
gldB point mutant, suggesting that GldB and GldE may
interact. GldE exhibits sequence similarity to Borrelia
burgdorferi TlyC and Salmonella enterica serovar
Typhimurium CorC.
Gliding motility is a trait shared
by a large number of bacteria belonging to different branches of the
eubacterial phylogenetic tree (21, 30, 42). Gliding
bacteria move over surfaces by mechanisms that do not involve flagella.
The rates of cell movement vary from approximately 0.05 µm/s for
Myxococcus xanthus (43) to 10 µm/s for some
filamentous cyanobacteria (21). Gliding cells travel
individually or in swarms, resulting in the formation of colonies that
have thin spreading edges.
Since the initial observations of gliding motility nearly 200 years ago
(47) a number of mechanisms to explain gliding have been
proposed (10, 30, 33, 42, 53). It is likely that the
mechanisms of movement employed by diverse gliding bacteria are not all
closely related. M. xanthus, a member of the Flavobacterium johnsoniae (formerly Cytophaga
johnsonae) (4) is a common gliding bacterium that
belongs to the CFB group. Cells of F. johnsoniae glide at
rates of up to 10 µm/s over wet glass, although speeds of 2 to 4 µm/s are more typical (33). Techniques to genetically
manipulate F. johnsoniae were recently developed
(31) and have been used to identify three of the genes, gldA, gldB, and ftsX, that are
required for gliding motility (1, 23, 26). GldA exhibits
sequence similarity to ATP-binding cassette transport proteins. It
presumably functions as part of a transporter complex, but its exact
role in gliding motility is not yet understood. GldB is not similar in
sequence to any proteins of known function, and its exact role in
gliding motility is unknown. FtsX is needed for both cell division and
gliding motility. ftsX mutants form filamentous, nongliding
cells. It is not known whether the defects in gliding are a direct
result of the mutations in ftsX or an indirect effect of the
defects in cell division. Here we report the identification and
characterization of two additional genes involved in F. johnsoniae gliding motility.
Bacterial and bacteriophage strains, plasmids, and growth
conditions.
F. johnsoniae UW101 (ATCC 17061) was the
wild-type strain used in these studies. All mutants were derived from
this strain. The 50 nongliding F. johnsoniae mutants that
were obtained from J. Pate were previously described (11, 23,
50). The bacteriophages active against F. johnsoniae
that were used in this study (
0021-9193/01/$04.00+0 DOI: 10.1128/JB.183.14.4167-4175.2001
Copyright © 2001, American Society for Microbiology. All rights reserved.
Cloning and Characterization of the
Flavobacterium johnsoniae Gliding Motility Genes
gldD and gldE
ABSTRACT
Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References
INTRODUCTION
Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References
subdivision of the proteobacteria, appears to have two independent systems of
gliding motility, "S" or social gliding motility and "A" or adventurous gliding motility (20, 42). S motility requires type IV pili and is probably related to the bacterial twitching motility of Pseudomonas aeruginosa and other bacteria
(25, 39, 48, 51). Extension and retraction of pili appear
to be responsible for this type of cell movement (32, 44).
The mechanism of M. xanthus A motility is not known, but it
does not involve pili (42). Some cyanobacteria employ type
IV pili to move over surfaces (5), whereas others may rely
on polysaccharide secretion to power cell movement (22).
Numerous bacteria belonging to the Cytophaga-Flavobacterium-Bacteroides (CFB) branch of the
eubacterial phylogenetic tree exhibit rapid gliding motility (4,
37). This form of movement does not appear to involve pili
(19, 37) and is unlikely to be powered by polysaccharide
secretion (23, 28). Lapidus and Berg performed extensive
behavioral studies of Cytophaga sp. strain U67
movements and proposed that cells have a system of tracks anchored to
the peptidoglycan. In their model outer membrane components are driven
along these tracks by periplasmic and cell membrane proteins that
obtain energy from the proton motive force (28). Other
models to explain gliding of members of the CFB group include
contraction or expansion of fibrils in the periplasm or cytoplasm
(9), the functioning of rotary motors (34),
the generation of waves in the outer membrane (13), and
the movement of conveyor belts made of polysaccharide or protein along
the cell surface (30).
MATERIALS AND METHODS
Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References
Cj1, Cj13, Cj23, Cj29,
Cj42, Cj48, and Cj54) have been previously described
(11, 35, 50). The Escherichia coli strains used were DH5
MCR (Gibco/BRL Life Technologies, Gaithersburg, Md.), HB101
(8), and S17-1 (40). E. coli
strains were grown in Luria-Bertani medium at 37°C, and
F. johnsoniae strains were grown in Casitone-yeast extract
(CYE) medium at 30°C, as previously described (31). To
observe colony spreading, F. johnsoniae was grown on PY2
medium (2 g of peptone, 0.5 g of yeast extract, and 10 g of
agar/liter, pH 7.3) at 25°C. Antibiotics were used at the following
concentrations when needed: ampicillin, 100 µg/ml; chloramphenicol,
30 µg/ml; erythromycin, 100 µg/ml; tetracycline, 20 µg/ml; and
kanamycin, 30 µg/ml. Plasmids and primers used in this study are
listed in Table 1 and Table
2, respectively.
TABLE 1.
Plasmids used in this studya
TABLE 2.
Primers used in this study
Transposon mutagenesis and cloning of
Tn4351-disrupted gldD.
Tn4351 was introduced into wild-type F. johnsoniae as described previously (23).
Tn4351-disrupted gldD was cloned from nongliding mutant CJ282 essentially as previously described (26).
Briefly, chromosomal DNA was digested with HindIII,
ligated into pBC SK(+), and transferred by electroporation into
E. coli DH5MCR. Cells were plated on Luria-Bertani medium
containing tetracycline to select clones carrying pMM122, which
contained 3.5 kb of Tn4351 DNA and 2.5 kb of adjacent
F. johnsoniae DNA.
Cloning of gldD and surrounding DNA from wild-type F. johnsoniae. A library of wild-type F. johnsoniae DNA was constructed in LambdaGEM-11 essentially as previously described (26). Lambda clones containing the regions of interest were detected by hybridization with radiolabeled DNA prepared using the 6-kb HindIII fragment of pMM122 and the Prime-a-Gene labeling kit (Promega, Madison, Wis.).
DNA from one of the lambda clones was isolated, and a 4.6-kb SalI fragment, which contained gldD and adjacent genes, was subcloned into pBC SK(+) to generate pMM129. For complementation of CJ282, the 4.6-kb fragment was isolated from pMM129 as a KpnI-BamHI fragment and cloned into shuttle vector pCP23 to generate pMM144. Subclones were generated to determine the regions needed for complementation. To remove the region downstream from gldD, pMM144 was digested with Bst98I and XhoI, treated with a DNA polymerase Klenow fragment to make the ends blunt, and treated with T4 DNA ligase to generate pMM166. pMM168, which carries gldD and gldE, was generated by digesting pMM166 with NsiI and KpnI and inserting the 2.3-kb fragment into pCP23 which had been digested with PstI and KpnI. pMM209, which carries gldD and 1.15 kb of upstream DNA, was generated by removing the 1- and 0.8-kb ClaI fragments from pMM166. pMM210 was generated by inserting the 2.1-kb kanamycin resistance cassette from pHP45
kan, which carries transcription termination signals on each
end, into the BamHI site of pMM209. pMM213 was identical to
pMM210 except that the kanamycin resistance cassette was inserted in
the opposite orientation. pMM169, which carries gldE but not gldD, was generated by inserting the 2.1-kb
BglII-PstI fragment of pMM166 into pCP23 which
had been digested with BamHI and PstI. Plasmid
DNA was transferred to nongliding mutants by conjugation or
electroporation as previously described (23, 31).
Nucleic acid sequencing. Nucleic acid sequencing was performed by the dideoxy nucleotide procedure using an automated (Applied Biosystems) sequencing system. Sequences were analyzed with MacVector and AssemblyLign software (Oxford Molecular Group Inc., Campbell, Calif.), and comparisons to database sequences were made using the BLAST (2) and FASTA (36) algorithms. Preliminary sequence data for Porphyromonas gingivalis were obtained from The Institute for Genomic Research website at http://www.tigr.org. Preliminary sequence data for Cytophaga hutchinsonii were obtained from The DOE Joint Genome Institute at http://jgi.doe.gov/.
RNA analysis. Total RNA was isolated from overnight cultures of F. johnsoniae UW101 using the AquaPure RNA isolation kit (Bio-Rad, Hercules, Calif.). Northern blots were performed using the NorthernMax kit (Ambion, Austin, Tex.). Twenty micrograms of RNA was separated by electrophoresis and transferred to nylon membranes. RNA was fixed to the membranes by UV cross-linking. A 446-bp fragment of DNA internal to gldD was radiolabeled using the Prime-a-Gene system (Promega) and used as a probe.
Promoter analysis was conducted by reverse transcription-PCR (RT-PCR) using the 5'-RACE kit (Gibco/BRL Life Technologies). RNA was treated with 50 U of RNase-free DNase (Ambion)/ml for 30 min at 37°C to remove residual DNA. Samples were assumed to be DNA free if a 35-cycle PCR with primers 70 and 377 produced no visible product on ethidium bromide-stained agarose gels. cDNA was synthesized using primer 73 or 378 for gldE and primer 59 or 474 for gldD. Following cDNA synthesis and oligo(C) tailing with terminal deoxytransferase, nested PCRs were carried out using the Gibco/BRL Life Technologies "anchor" primer and primers internal to the expected cDNAs (Table 2). The PCR products were sequenced to determine the transcription start sites.Overexpression of recombinant GldD and GldE and antibody
production.
A fragment coding for the C-terminal 166 amino acids
of GldD was amplified by PCR using Vent polymerase (New England
Biolabs) and primers 415 and 409. The amplified fragment was cloned
into pCR-Script Amp SK(+) to produce pDH256. The gldD
fragment was isolated from pDH256 as an
EcoRI-PstI fragment and was ligated into
expression vector pMAL-c2 (New England Biolabs) to generate pDH263. A
fragment which codes for the C-terminal 296 amino acids of GldE was
amplified as described above using primers 214 and 212 and cloned into
pCR-Script Amp SK(+) to generate pDH247. gldE was isolated
from pDH247 as a PstI-HindIII fragment and
ligated into pMAL-c2 to produce pDH249. All fusion constructs were
confirmed by DNA sequencing. Expression of GldD and GldE fusion
proteins was induced in E. coli DH5MCR by the addition of
0.3 mM IPTG (isopropyl-
-D-thiogalactopyranoside). Fusion
proteins were partially purified by preparative sodium dodecyl
sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) and recovered as
previously described (23). Antibodies against GldD and
GldE were generated and affinity purified essentially as previously
described (23).
Cell fractionation and Western blot analyses. F. johnsoniae cells were fractionated into soluble, inner membrane (Sarkosyl-soluble), and outer membrane (Sarkosyl-insoluble) fractions as described previously (23). Proteins were separated by SDS-PAGE, and Western blot analyses were performed as previously described (23).
Microscopic observations of cell movement. Wild-type and mutant cells of F. johnsoniae were examined for movement over glass and agar surfaces by phase-contrast microscopy as previously described (23). The ability of cells to bind and propel 0.2- and 0.4-µm polystyrene latex spheres (Seradyn, Indianapolis, Ind.) was also examined. Cells were grown to late exponential phase (1 × 109 to 3 × 109 cells/ml) in CYE broth, and latex spheres in distilled water were added to a final concentration of approximately 0.08 mg/ml. The cell suspension (0.02 ml) was spotted on a microscope slide, covered with an O2-permeable membrane (Yellow Springs Instrument Co., Inc., Yellow Springs, Ohio), and examined by phase-contrast microscopy on a heated (30°C) microscope stage.
Measurements of phage sensitivity. Sensitivity to F. johnsoniae phage was determined by spotting 1 to 10 µl of phage lysates (2 × 107 phage/ml) onto lawns of cells in CYE overlay agar followed by incubation for 24 h at 25°C (23).
Genetic nomenclature. Genes involved in gliding motility were given the name gld followed by a letter. Open reading frames (ORFs) that exhibited strong sequence similarity to genes of known function were named after the corresponding genes. ORFs of unknown function that did not exhibit strong similarity to previously described genes were given the provisional name fjo (F. johnsoniae ORF) followed by a number.
Nucleotide sequence accession number. The sequence reported in this paper has been deposited in the GenBank database under accession no. AF287009.
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RESULTS |
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Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References
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Isolation of nongliding mutants by Tn4351 mutagenesis. F. johnsoniae was mutagenized with Tn4351, and 280 nonspreading mutants were isolated from approximately 20,000 erythromycin-resistant transconjugants. Sixty-eight of the mutants were completely deficient in gliding motility as determined by microscopic analysis of individual cells in wet mounts and on agar slide cultures. Thirty-five of these exhibited filamentous morphology and were not considered further in this study. Of the remaining 33 nongliding mutants, 2 were complemented by introduction of pSA21, which contains gldA, and 8 were complemented by pDH233, which contains gldB, suggesting that these mutants had Tn4351 insertions in previously characterized genes gldA and gldB. The insertions in gldA or gldB were verified by PCR amplification and sequencing (data not shown). The remaining 23 nongliding mutants had mutations in gld genes which had not been previously characterized. One of these mutants, CJ282, was chosen for further analysis in this study.
Cloning and sequencing of gldD and adjacent
DNA.
We determined the site of insertion of Tn4351 in
CJ282 by cloning the HindIII fragment that spans 3.5 kb
of the transposon, including the tetX gene, and 2.5 kb of
adjacent DNA from the F. johnsoniae genome. This fragment
was used to probe a lambda library of wild-type F. johnsoniae DNA, and several hybridizing clones were isolated.
These clones were used to determine the nucleotide sequence of a 4.6-kb
region that spans the site of the original Tn4351 insertion.
Analysis of this region identified seven ORFs (Fig.
1).
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Complementation of strains carrying Tn4351
insertions in gldD.
Introduction of pMM144, which
carries a 4.6-kb SalI fragment spanning the gldD
gene, into F. johnsoniae mutant CJ282 restored the ability
of cells to glide and to form spreading colonies. Electroporation of
pMM144 into the other 33 Tn4351-induced nongliding mutants
resulted in the complementation of 2 additional mutants (CJ695 and
CJ758). The sites of Tn4351 insertion in CJ695 and CJ758
were identified by cloning or amplification of the disrupted genes and
determination of the nucleotide sequence (Fig. 1). Further subcloning
revealed that introduction of pMM168, which contains only
gldE and gldD, restored motility to each of these
mutants. Inspection of the DNA sequence suggested that gldE
and gldD may constitute an operon, since the stop codon of
gldE is adjacent to the start codon of gldD. To
determine whether gldE and gldD are
transcriptionally linked, we constructed pMM210 and pMM213, which each
carry gldD and the 3' end of gldE but which lack
the N-terminal coding region and the transcriptional start site of gldE. pMM210 and pMM213 also carry the kanamycin resistance
omega fragment (14), which has transcription terminators
at each end to prevent transcription of gldD from promoters
within the vector. The only difference between pMM210 and pMM213 is the
orientation of the omega fragment upstream of gldD.
Introduction of pMM210 or pMM213 into CJ282 resulted in production of
the GldD protein as determined by Western blot analysis (see Fig. 4)
and restored gliding motility (Fig. 3).
This indicates that gldD is expressed from a promoter that
lies within gldE. pMM210 or pMM213 also restored motility to
the other Tn4351-induced gldD mutants, CJ695 and
CJ758 (data not shown).
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Identification of spontaneous and chemically induced gldD mutants. Pate and colleagues isolated a large number of spontaneous and chemically induced nongliding mutants (11, 50). We introduced pMM144 into 50 of these mutants to determine if any have defects in gldD or in nearby genes. pMM144 completely restored motility to 3 of the 50 mutants (UW102-42, UW102-58, and UW102-97). pMM210, which carries just gldD, also complemented each of these nongliding mutants, suggesting that each has a mutation in gldD.
Overexpression of GldE suppresses a gldB point
mutation.
In the course of complementation analyses we discovered
that introduction of pMM168, which spans gldE and
gldD, into gldB point mutant UW102-103 resulted
in partial restoration of motility. This suppression was allele
specific, since introduction of pMM168 into three other gldB
point mutants, UW102-90, UW102-99, and UW102-154, did not restore
motility. We amplified and sequenced the gldE and
gldD genes from UW102-103 and determined that there was no mutation in either gene, indicating that the partial restoration of
motility was not due to complementation of a gldD or
gldE mutation. Further subcloning revealed that introduction
of pMM169, which contains gldE but not gldD,
suppressed the motility defect of UW102-103 (Fig. 3), whereas
introduction of pMM213, which expresses gldD but not
gldE, did not (data not shown). pMM169 has a copy number of
approximately 10 in F. johnsoniae, and Western blot analyses
demonstrated that cells carrying pMM169 produce about 10 times as much
GldE protein as wild-type cells (Fig. 4)
or cells of gldB mutant UW102-103 (data not shown).
Apparently, this limited overexpression of GldE in UW102-103 resulted
in partial restoration of motility and colony spreading.
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RNA analysis.
The complementation results described above
indicated that gldD was transcribed from a promoter within
the gldE coding sequence. Northern blot analyses of
wild-type F. johnsoniae RNA confirmed this result.
Hybridization with a probe to an internal fragment of gldD
revealed a band of approximately 650 bp (Fig.
5). A transcript of this size is large
enough to encompass gldD but could not contain gldD and gldE.
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Cellular localization of GldD and GldE. The cellular locations of GldD and GldE were determined by cell fractionation followed by Western blot analyses as described in Materials and Methods. GldD and GldE were found primarily in the cytoplasmic membrane fraction of wild-type cells (Fig. 4, lanes 2), as expected from analysis of the predicted protein sequences. GldD exhibited an apparent molecular mass by SDS-PAGE of 23 kDa, whereas GldE was approximately 50 kDa. These sizes are similar to those predicted from sequence analysis: 21.3 (GldD) and 48.6 kDa (GldE). When GldD was overexpressed, some of the protein was soluble rather than membrane associated (Fig. 4A, lanes 7 and 8). The soluble GldD protein migrated with a lower apparent molecular weight than the protein in the membrane fraction. Overexpression of GldD may result in limited proteolysis of some of the proteins, resulting in removal of the N-terminal hydrophobic tail.
Since gldA, gldB, gldD, and gldE are all involved in motility, mutations in one gene might affect the expression, localization, or stability of the other gld proteins. We used antisera raised against GldB, GldD, and GldE to determine the amount and location of each protein in various mutants. CJ288, which carries an insertion in gldA, produced normal amounts of GldB, GldD, and GldE, and these proteins localized to the cytoplasmic membrane as expected (data not shown). CJ588, which carries an insertion in gldB, failed to produce detectable levels of GldB but produced normal amounts of GldD and GldE, and these proteins localized to the cytoplasmic membrane (data not shown). Finally, CJ282, which carries a Tn4351 insertion in gldD, did not produce detectable levels of GldD but produced normal amounts of GldB and GldE, and these proteins localized to the cytoplasmic membrane (Fig. 4 and data not shown).Phage resistance of gldD mutants.
It has
previously been reported that many nonmotile F. johnsoniae
mutants are resistant to infection by a number of F. johnsoniae bacteriophages (49). We tested the
sensitivities of F. johnsoniae strains UW101, CJ282, and
CJ736 (CJ282 carrying pMM213) to F. johnsoniae
bacteriophages Cj1, Cj13, Cj23, Cj29, Cj42, Cj48, and
Cj54. F. johnsoniae UW101 was readily lysed by these
phages, whereas gldD mutant CJ282 was resistant to infection
by each of them. Introduction of pMM213 into CJ282, which restored
motility to the mutant, also restored sensitivity to each of the phages.
Movement of latex spheres by gldD mutant cells. Wild-type cells of F. johnsoniae and related bacteria bind latex spheres on their surfaces and propel these spheres along the length of the cells (28, 34). Spheres move at approximately the same speed as gliding cells, which suggests that the machinery that propels spheres is also responsible for cell gliding. We examined the ability of cells of F. johnsoniae strains UW101, CJ282, and CJ736 (CJ282 carrying pMM213) to propel latex spheres. Wild-type cells propelled the spheres, whereas cells of gldD mutant CJ282 did not. Cells of CJ736 propelled the spheres as well as wild-type cells.
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DISCUSSION |
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Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References
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The mechanism responsible for F. johnsoniae gliding motility is not known. Previously, we identified three genes, gldA, gldB, and ftsX, that are required for gliding (1, 23, 26). gldA codes for a protein that is similar in sequence to the ATP-hydrolyzing components of ATP-binding cassette-transporter complexes (1). Presumably GldA is involved in some transmembrane transport process that is directly or indirectly required for gliding motility. However, it is unlikely that ATP hydrolysis by GldA is the driving force for cell movement because others have demonstrated that the proton motive force is the likely energy source for the gliding of F. johnsoniae and related bacteria (34, 38). GldB is anchored in the cytoplasmic membrane and has a large hydrophilic domain. The exact function of GldB in gliding motility is not known. FtsX is needed for both cell division and gliding motility. The inability of ftsX mutants to glide may be an indirect effect of the defects in cell division. The results presented in this paper identify another gene, gldD, that is required for gliding. Transposon insertions in gldD eliminate gliding motility, and introduction of a wild-type copy of gldD on a plasmid restores motility to the mutants. F. johnsoniae GldD did not exhibit significant similarity to proteins of known function. The only protein that was similar to GldD was a putative protein that we refer to as C. hutchinsonii GldD. C. hutchinsonii and F. johnsoniae are distant relatives within the CFB branch of the eubacteria, and they both exhibit rapid gliding motility (4). GldD may be required for gliding in both organisms.
gldE, which lies immediately upstream of gldD, also appears to be involved in gliding motility. Overexpression of gldE resulted in partial suppression of gldB point mutant UW102-103, which suggests that GldE and GldB may interact. Suppression was allele specific, since overexpression of gldE did not restore motility to three other gldB point mutants. The presence of large amounts of GldE may stabilize the mutant GldB protein of UW102-103, resulting in partial recovery of motility. Both GldB (23) and GldE (this study) localize to the cytoplasmic membrane, making such an interaction possible. We do not know whether gldE is required for gliding motility, since we were unable to construct strains with mutations in this gene. F. johnsoniae GldE exhibits strong sequence similarity to a large number of proteins. It shares greatest amino acid sequence similarity with a putative protein of unknown function from P. gingivalis. Similarity between these two extends over essentially the entire length of each protein. Both proteins have hydrophobic regions near their N termini. Many other proteins that exhibit strong similarity to F. johnsoniae GldE lack similarity in this N-terminal domain. Instead, similarity begins after approximately amino acid 150 of F. johnsoniae GldE. The functions of most of these proteins are not known, but Serpulina hyodysenteriae TlyC may be a hemolysin (46), and S. enterica serovar Typhimurium CorC is thought to function in magnesium export (16).
The arrangement of gldE and gldD suggested that
they might be cotranscribed, since the stop codon of gldE
was adjacent to the start codon of gldD. However,
complementation analyses and Northern blot analysis indicated that
under the conditions tested, gldD was transcribed separately
from gldE, and RT-PCR analyses identified transcription
start sites upstream of each gene. Conventional E. coli
70 promoter sequences were not detected
upstream of either transcription start site. This is not surprising
given the distant relationship between E. coli and F. johnsoniae. Consensus promoter sequences have not been determined
for F. johnsoniae, but a number of promoters of another
member of the CFB branch, Bacteroides fragilis, have been
identified, and a tentative consensus has been proposed
(3). The consensus sequence (TAXXTTTG) is
generally centered 4 to 18 nucleotides upstream of the transcriptional
start site. Sequences with similarity to the B. fragilis
consensus were identified 8 nucleotides upstream of the transcriptional
start site of gldE (TAACTTTG) and 21 nucleotides
upstream of the transcriptional start site of gldD
(TAATTTTC). We do not know whether these sequences are
important for transcription of gldE and gldD.
Further studies will be needed to determine the requirements for
transcription initiation of F. johnsoniae genes.
As mentioned above, C. hutchinsonii, which exhibits gliding motility, contains homologs to F. johnsoniae gldD and gldE. C. hutchinsonii also contains a homolog to F. johnsoniae ssb. Surprisingly, the gene order ssb-gldE-gldD is conserved between these bacteria, although each of the individual genes has diverged considerably. The significance of this gene arrangement is not known. P. gingivalis, another member of the CFB group, also has a gldE homolog situated downstream from a putative ssb gene. P. gingivalis does not exhibit gliding motility and lacks a gldD homolog.
The nearly complete genome sequence of C. hutchinsonii was recently made available for analysis. Since C. hutchinsonii is distantly related to F. johnsoniae, exhibits gliding motility, and has homologs to gldD and gldE, we examined its genome for the presence of genes that are similar to other F. johnsoniae gld genes. ORFs which code for proteins that are similar to GldA (49% amino acid identity over 297 amino acids), GldB (29% amino acid identity over 346 amino acids), and GldC (44% amino acid identity over 98 amino acids) were found. (GldC is not required for F. johnsoniae gliding motility, but the absence of GldC results in reduced colony spreading [23].) The presence of homologs to each of the F. johnsoniae gld genes suggests that similar motility machinery may be used by these bacteria. gldB, gldC, and gldD lack close homologs in the databases other than those found in the C. hutchinsonii genome. The involvement of these novel genes in F. johnsoniae gliding and presumably in C. hutchinsonii gliding may indicate that Cytophaga-Flavobacterium gliding motility is not closely related to other motility systems, mediated by flagella or type IV pili, that have been well studied (6, 48).
Pate and colleagues isolated a bank of 50 independent spontaneous or chemically induced mutants which are completely deficient in gliding motility (11, 23, 50). Complementation and sequence analyses indicate that four of these mutants are nonmotile because of mutations in gldA (1), four others fail to glide because of mutations in gldB (23), and three are nonmotile because of mutations in gldD (this study). Complementation of 11 of 50 nongliding mutants by just 3 genes suggests that a relatively small number of genes, perhaps less than 20, are required specifically for gliding motility.
Gliding motility is thought to be the result of movement of cell surface components. The movement of these components can be observed by adding latex spheres, which bind to, and are propelled along, the surfaces of the cells. Many nongliding mutants fail to propel spheres (11, 18). Cells of gldD mutants were unable to propel latex spheres. Complementation of the motility defect by introduction of gldD on a plasmid restored the ability to move spheres. These results are similar to those observed for gldA, gldB, and ftsX mutants (1, 23, 26). In each case complete disruption of gliding motility was associated with loss of ability to propel spheres over the cell surface and sphere movement was restored by complementation. These results support the suggestion that the machinery that propels latex spheres along the cell surface is also involved in cell movement.
It has previously been reported that many nongliding F. johnsoniae mutants are resistant to bacteriophages that infect wild-type cells (49). Movement of the cell surface may be necessary to allow access of phages to their receptors. Others have isolated nongliding mutants which remain susceptible to infection by some bacteriophages (18). In this study we found that cells with mutations in gldD were nonmotile and were resistant to bacteriophage infection. Restoration of motility by introduction of gldD on a plasmid restored sensitivity to bacteriophage infection. These results are similar to those obtained for strains with mutations in gldA (1), gldB (23), or ftsX (26) and lend support to the observation that mutations that disrupt gliding motility generally result in phage resistance.
In addition to the nongliding mutants, which were the focus of this study, we also isolated a large number of "motile nonspreading" (MNS) mutants. MNS mutants form nonspreading colonies, but individual cells exhibit gliding motility. Cells of some MNS mutants glide as well as wild-type cells, whereas others appear to be crippled but still move over surfaces. MNS mutants retain the ability to move latex spheres and are generally sensitive to F. johnsoniae bacteriophages. Of the 280 mutants isolated in this study, 212 were MNS mutants and 68 were nongliding mutants. Similar results have been previously reported (11, 18, 26, 50). MNS mutants appear to retain a functional motility machinery but may have alterations to their cell surfaces that prevent efficient spreading on agar. Defects in cell surface polysaccharides have been observed to result in decreased colony spreading without eliminating gliding motility (17).
The mechanism of F. johnsoniae gliding motility remains a mystery. Genetic analyses have uncovered four genes (gldA, gldB, gldD, and ftsX) that are required for F. johnsoniae gliding motility. Identification of the remaining gld genes and analysis of their protein products should result in a better understanding of the F. johnsoniae gliding machinery and the mechanism of cell movement.
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ACKNOWLEDGMENTS |
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This research was supported by a grant from the National Science Foundation (MCB-9727825) and by a Shaw Scientist Award to M.J.M. from The Milwaukee Foundation.
DNA sequencing was performed by the Automated DNA Sequencing Facility at the University of Wisconsin-Milwaukee Department of Biological Sciences. Preliminary sequence data for C. hutchinsonii were obtained from The DOE Joint Genome Institute at http://jgi.doe.gov/. We thank D. Saffarini for careful reading of the manuscript.
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FOOTNOTES |
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* Corresponding author. Mailing address: Department of Biological Sciences, 260 Lapham Hall, University of Wisconsin-Milwaukee, 3209 N. Maryland Ave., Milwaukee, WI 53211. Phone: (414) 229-5844. Fax: (414) 229-3926. E-mail: mcbride{at}uwm.edu.
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REFERENCES |
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Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References
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| 1. |
Agarwal, S.,
D. W. Hunnicutt, and M. J. McBride.
1997.
Cloning and characterization of the Flavobacterium johnsoniae (Cytophaga johnsonae) gliding motility gene, gldA.
Proc. Natl. Acad. Sci. USA
94:12139-12144 |
| 2. | Altschul, S. F., W. Gish, W. Miller, E. W. Myers, and D. J. Lipman. 1990. Basic local alignment search tool. J. Mol. Biol. 215:403-410[CrossRef][Medline]. |
| 3. | Bayley, D. P., E. R. Rocha, and C. J. Smith. 2000. Analysis of cepA and other Bacteroides fragilis genes reveals a unique promoter structure. FEMS Microbiol. Lett. 193:149-154[CrossRef][Medline]. |
| 4. |
Bernardet, J.-F.,
P. Segers,
M. Vancanneyt,
F. Berthe,
K. Kersters, and P. Vandamme.
1996.
Cutting a Gordian knot: emended classification and description of the genus Flavobacterium, emended description of the family Flavobacteriaceae, and proposal of Flavobacterium hydatis nom. nov. (basonym, Cytophaga aquatilis Strohl and Tait 1978).
Int. J. Syst. Bacteriol.
46:128-148 |
| 5. |
Bhaya, D.,
N. Watanabe,
T. Ogawa, and A. R. Grossman.
1999.
The role of an alternative sigma factor in motility and pilus formation in the cyanobacterium Synechocystis sp. strain PCC6803.
Proc. Natl. Acad. Sci. USA
96:3188-3193 |
| 6. | Blair, D. F. 1995. How bacteria sense and swim. Annu. Rev. Microbiol. 49:489-522[CrossRef][Medline]. |
| 7. |
Blattner, F. R.,
G. Plunkett III,
C. A. Bloch,
N. T. Perna,
V. Burland,
M. Riley,
J. Collado-Vides,
J. D. Glasner,
C. K. Rode,
G. F. Mayhew,
J. Gregor,
N. W. Davis,
H. A. Kirkpatrick,
M. A. Goeden,
D. J. Rose,
B. Mau, and Y. Shao.
1997.
The complete genome sequence of Escherichia coli K-12.
Science
277:1453-1462 |
| 8. | Bolivar, F., and K. Backman. 1979. Plasmids of E. coli as cloning vectors. Methods Enzymol. 68:245-267[Medline]. |
| 9. | Burchard, R. P. 1984. Gliding motility and taxes, p. 301. In E. Rosenberg (ed.), Myxobacteria, development and cell interactions. Springer-Verlag, New York, N.Y. |
| 10. | Burchard, R. P. 1981. Gliding motility of prokaryotes: ultrastructure, physiology, and genetics. Annu. Rev. Microbiol. 35:497-529[CrossRef][Medline]. |
| 11. |
Chang, L. Y. E.,
J. L. Pate, and R. J. Betzig.
1984.
Isolation and characterization of nonspreading mutants of the gliding bacterium Cytophaga johnsonae.
J. Bacteriol.
159:26-35 |
| 12. | Draper, D. E. 1996. Translation initiation, p. 902-908. In F. C. Neidhardt, R. Curtiss III, J. L. Ingraham, E. C. C. Lin, K. B. Low, B. Magasanik, W. S. Reznikoff, M. Riley, M. Schaechter, and H. E. Umbarger (ed.), Escherichia coli and Salmonella: cellular and molecular biology. ASM Press, Washington, D.C. |
| 13. | Duxbury, T., B. A. Humphrey, and K. C. Marshall. 1980. Continuous observations of bacterial gliding motility in a dialysis microchamber: the effects of inhibitors. Arch. Microbiol. 124:169-175[CrossRef]. |
| 14. | Fellay, R., J. Frey, and H. Krisch. 1987. Interposon mutagenesis of soil and water bacteria: a family of DNA fragments designed for in vitro insertional mutagenesis of gram-negative bacteria. Gene 52:147-154[CrossRef][Medline]. |
| 15. | Fraser, C. M., S. Casjens, W. M. Huang, G. G. Sutton, R. Clayton, R. Lathigra, O. White, K. A. Ketchum, R. Dodson, E. K. Hickey, M. Gwinn, B. Dougherty, J. F. Tomb, R. D. Fleischmann, D. Richardson, J. Peterson, A. R. Kerlavage, J. Quackenbush, S. Salzberg, M. Hanson, R. van Vugt, N. Palmer, M. D. Adams, J. Gocayne, and J. C. Venter. 1997. Genomic sequence of a Lyme disease spirochaete, Borrelia burgdorferi. Nature 390:580-586[CrossRef][Medline]. |
| 16. | Gibson, M. M., D. A. Bagga, C. G. Miller, and M. E. Maguire. 1991. Magnesium transport in Salmonella typhimurium: the influence of new mutations conferring Co2+ resistance on the CorA Mg2+ transport system. Mol. Microbiol. 5:2753-2762[Medline]. |
| 17. |
Godchaux, W., III,
M. A. Lynes, and E. R. Leadbetter.
1991.
Defects in gliding motility in mutants of Cytophaga johnsonae lacking a high-molecular-weight cell surface polysaccharide.
J. Bacteriol.
173:7607-7614 |
| 18. | Gorski, L., W. Godchaux III, E. R. Leadbetter, and R. R. Wagner. 1992. Diversity in surface features of Cytophaga johnsonae motility mutants. J. Gen. Microbiol. 138:1767-1772. |
| 19. | Henrichsen, J., and J. Blom. 1975. Examination of fimbriation of some gram-negative rods with and without twitching and gliding motility. Acta Pathol. Microbiol. Scand. B 83:161-170[Medline]. |
| 20. | Hodgkin, J., and D. Kaiser. 1979. Genetics of gliding motility in Myxococcus xanthus (Myxobacteriales): two gene systems control movement. Mol. Gen. Genet. 171:177-191[CrossRef]. |
| 21. | Hoiczyk, E. 2000. Gliding motility in cyanobacteria: observations and possible explanations. Arch. Microbiol. 174:11-17[CrossRef][Medline]. |
| 22. | Hoiczyk, E., and W. Baumeister. 1998. The junctional pore complex, a prokaryotic secretion organelle, is the molecular motor underlying gliding motility in cyanobacteria. Curr. Biol. 8:1161-1168[CrossRef][Medline]. |
| 23. |
Hunnicutt, D. W., and M. J. McBride.
2000.
Cloning and characterization of the Flavobacterium johnsoniae gliding motility genes, gldB and gldC.
J. Bacteriol.
182:911-918 |
| 24. | Jarosik, G. P., and E. J. Hansen. 1994. Cloning and sequencing of the Haemophilus influenzae ssb gene encoding single-strand DNA-binding protein. Gene 146:101-103[CrossRef][Medline]. |
| 25. |
Kaiser, D.
1979.
Social gliding is correlated with the presence of pili in Myxococcus xanthus.
Proc. Natl. Acad. Sci. USA
76:5952-5956 |
| 26. |
Kempf, M. J., and M. J. McBride.
2000.
Transposon insertions in the Flavobacterium johnsoniae ftsX gene disrupt gliding motility and cell division.
J. Bacteriol.
182:1671-1679 |
| 27. | Kiyono, M., T. Omura, M. Inuzuka, H. Fujimori, and H. Pan-Hou. 1997. Nucleotide sequence and expression of the organomercurial-resistance determinants from a Pseudomonas K-62 plasmid pMR26. Gene 189:151-157[CrossRef][Medline]. |
| 28. |
Lapidus, I. R., and H. C. Berg.
1982.
Gliding motility of Cytophaga sp. strain U67.
J. Bacteriol.
151:384-398 |
| 29. |
Li, L.-Y.,
N. B. Shoemaker, and A. A. Salyers.
1995.
Location and characterization of the transfer region of a Bacteroides conjugative transposon and regulation of the transfer genes.
J. Bacteriol.
177:4992-4999 |
| 30. | McBride, M. J. Bacterial gliding motility: multiple mechanisms for cell movement over surfaces. Annu. Rev. Microbiol., in press. |
| 31. |
McBride, M. J., and M. J. Kempf.
1996.
Development of techniques for the genetic manipulation of the gliding bacterium Cytophaga johnsonae.
J. Bacteriol.
178:583-590 |
| 32. | Merz, A. J., M. So, and M. P. Sheetz. 2000. Pilus retraction powers bacterial twitching motility. Nature 407:98-102[CrossRef][Medline]. |
| 33. | Pate, J. L. 1988. Gliding motility. Can. J. Microbiol. 34:459-465. |
| 34. | Pate, J. L., and L.-Y. E. Chang. 1979. Evidence that gliding motility in prokaryotic cells is driven by rotary assemblies in the cell envelopes. Curr. Microbiol. 2:59-64. |
| 35. | Pate, J. L., S. J. Petzold, and L.-Y. E. Chang. 1979. Phages for the gliding bacterium Cytophaga johnsonae that infect only motile cells. Curr. Microbiol. 2:257-262[CrossRef]. |
| 36. | Pearson, W. R. 1990. Rapid and sensitive sequence comparison with FASTP and FASTA. Methods Enzymol. 183:63-98[Medline]. |
| 37. | Reichenbach, H. 1992. The order Cytophagales, p. 3631-3675. In A. Balows, H. G. Truper, M. Dworkin, W. Harder, and K. M. Schleifer (ed.), The prokaryotes. Springer-Verlag, New York, N.Y. |
| 38. |
Ridgway, H. F.
1977.
Source of energy for gliding motility in Flexibacter polymorphus: effects of metabolic and respiratory inhibitors on gliding movement.
J. Bacteriol.
131:544-556 |
| 39. |
Semmler, A. B. T.,
C. B. Whitchurch, and J. S. Mattick.
1999.
A re-examination of twitching motility in Pseudomonas aeruginosa.
Microbiology
145:2863-2873 |
| 40. | Simon, R., U. Priefer, and A. Puhler. 1983. A broad host range mobilization system for in vivo genetic engineering: transposon mutagenesis in gram negative bacteria. Bio/Technology 2:784-791. |
| 41. |
Smith, D. R.,
L. A. Doucette-Stamm,
C. Deloughery,
H. Lee,
J. Dubois,
T. Aldredge,
R. Bashirzadeh,
D. Blakely,
R. Cook,
K. Gilbert,
D. Harrison,
L. Hoang,
P. Keagle,
W. Lumm,
B. Pothier,
D. Qiu,
R. Spadafora,
R. Vicaire,
Y. Wang,
J. Wierzbowski,
R. Gibson,
N. Jiwani,
A. Caruso,
D. Bush,
H. Safer,
D. Patwell,
S. Prabhakar,
S. McDougall,
G. Shimer,
A. Goyal,
S. Pietrokovski,
G. M. Church,
C. J. Daniels,
J.-I Mao,
P. Rice,
J. Nölling, and J. N. Reeve.
1997.
Complete genome sequence of Methanobacterium thermoautotrophicum  H: functional analysis and comparative genomics.
J. Bacteriol.
179:7135-7155 |
| 42. |
Spormann, A. M.
1999.
Gliding motility in bacteria: insights from studies of Myxococcus xanthus.
Microbiol. Mol. Biol. Rev.
63:621-641 |
| 43. |
Spormann, A. M., and A. D. Kaiser.
1995.
Gliding movements in Myxococcus xanthus.
J. Bacteriol.
177:5846-5852 |
| 44. | Sun, H., D. R. Zusman, and W. Shi. 2000. Type IV pilus of Myxococcus xanthus is a motility apparatus controlled by the frz chemosensory system. Curr. Biol. 10:1143-1146[CrossRef][Medline]. |
| 45. | Takami, H., and K. Horikoshi. 2000. Analysis of the genome of an alkaliphilic Bacillus strain from an industrial point of view. Extremophiles 4:99-108[CrossRef][Medline]. |
| 46. | ter Huurne, A. A., S. Muir, M. B. van Houten, A. van der Zeijst, W. Gaastra, and J. G. Kusters. 1994. Characterization of three putative Serpulina hyodysenteriae hemolysins. Microb. Pathog. 16:269-282[CrossRef][Medline]. |
| 47. | Vaucher, J. P. 1803. Histoire des conferves d'eau douce, contenant leurs differents modes de reproduction, et la description de leurs principales especes, suivie de l'histoire des tremelles et des ulves d'eau douce. J. J. Paschoud, Geneva, Switzerland. |
| 48. | Wall, D., and D. Kaiser. 1999. Type IV pili and cell motility. Mol. Microbiol. 32:1-10[CrossRef][Medline]. |
| 49. | Wolkin, R. H., and J. L. Pate. 1986. Phage adsorption and cell adherence are motility-dependent characteristics of the gliding bacterium Cytophaga johnsonae. J. Gen. Microbiol. 132:355-367. |
| 50. | Wolkin, R. H., and J. L. Pate. 1985. Selection for nonadherent or nonhydrophobic mutants co-selects for nonspreading mutants of Cytophaga johnsonae and other gliding bacteria. J. Gen. Microbiol. 131:737-750. |
| 51. | Wu, S. S., and D. Kaiser. 1995. Genetic and functional evidence that type IV pili are required for social gliding motility in Myxococcus xanthus. Mol. Microbiol. 18:547-558[CrossRef][Medline]. |
| 52. | Yamamoto, H., S. Uchiyama, and J. Sekiguchi. 1996. Cloning and sequencing of a 27.8-kb nucleotide sequence of the 79 degrees-81 degrees region of the Bacillus subtilis genome containing the sspE locus. DNA Res. 3:257-262[Abstract]. |
| 53. | Youderian, P. 1998. Bacterial motility: secretory secrets of gliding bacteria. Curr. Biol. 8:R408-R411[CrossRef][Medline]. |
Journal of Bacteriology, July 2001, p. 4167-4175, Vol. 183, No. 14
0021-9193/01/$04.00+0 DOI: 10.1128/JB.183.14.4167-4175.2001
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