Microbiology Unit, Biochemistry Department,
University of Oxford, Oxford OX1 3QU,1
School of Biological Sciences, Sutton Bonnington Campus,
University of Nottingham, Sutton Bonnington, Leicestershire LE12
5RD,3 and Immunology
Division4 and Institute of
Genetics,5 University of Nottingham, Queens
Medical Centre, Nottingham NG7 2UH, United Kingdom, and
Department of Bioscience, Teikyo University, 1-1 Toyosatodai, Utsunomiya 320, Japan2
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INTRODUCTION |
Bacteria swim through liquids by
means of a propeller-like, rotating flagellum (23). The
major component of the flagellum is the long, extracellular filament, a
polymer of flagellin protein. The antigenicity of flagellin, its
variability, its property of self-assembly into filaments, and the ease
with which it may be purified have resulted in an extensive study of
these proteins and the gene(s) encoding them in many bacterial genera
(reviewed in reference 15). Electron microscopic
studies have revealed two distinct types of filaments called
"plain" and "complex" (32). Plain filaments have a
smooth appearance, whereas complex filaments have ridges and grooves on
the surface (32, 38, 39). Most bacteria including
Salmonella possess plain filaments. Complex filaments have
been observed for three species of soil bacteria: Pseudomonas
rhodos (32), Sinorhizobium meliloti
(10), and Sinorhizobium lupini (33).
Previously, it was suggested that filament type might correlate with
the mode of flagellar rotation. Bacteria with plain filaments can
switch their rotation from clockwise (CW) to counterclockwise, whereas
bacteria with complex filaments can rotate their flagella only in the
CW direction and do not switch direction of rotation but stop rotation
periodically (so-called unidirectional, intermittent rotation
[11]). Complex flagella are brittle and form
left-handed helices with little or no structural polymorphism
(11). Plain filaments are flexible and have distinct polymorphic forms with different helical characteristics. Filaments rotating in the counterclockwise direction (swimming cells) are normally left-handed helices (normal shape), and a change in direction of rotation to CW (tumbling cells) converts them into right-handed helices (curly shape [24]). This polymorphic ability
is required for the swimming and tumbling of bacterial taxis.
Plain filaments can also be induced to change shape in vitro
(polymorphic transitions) by changing environmental conditions such as
pH, temperature, salt concentration, organic solvent, viscous flow, or
sugar concentrations (18, 34). Although it is not clear
whether such transitions have a physiological effect on swimming
behavior in the wild, they may be a useful way of modulating motility
under different conditions in the absence of tactic stimuli. A
comparison of the amino acid sequences from plain filament flagellin
proteins of many species reveals a relatively high degree of
conservation in the amino- and carboxyl-terminal regions, whereas the
central regions are very variable even between flagellins from
different strains of the same species (8, 14, 15).
A notable deviation from the correlation between filament types and
mode of flagellar rotation is seen in the purple, nonsulfur, aquatic
bacterium Rhodobacter sphaeroides (37). Each cell
has a single plain filament but displays unidirectional, intermittent flagellar rotation. Moreover, the filament can take on at least three
distinct polymorphic forms in vivo: a normal structure while rotating,
an apparently straight form during fast rotation, and an unusual,
loosely coiled conformation during a stop. Detached filaments with the
former and latter polymorphic forms can also be observed (2,
3). In light of these interesting differences between R. sphaeroides and other species, we chose to study the filament of
this organism in greater detail. In this paper, we describe polymorphic
forms of the filament induced in vitro by changes in pH and ionic
strength. We also report sequence analysis and expression studies of
the fliC gene, which encodes the flagellin protein of this organism.
| |
MATERIALS AND METHODS |
Strains, plasmids, and media.
The bacterial strains and
plasmids used in this work are described in Table
1. Growth media and antibiotic selection
were as described previously (35). Motility analysis was
carried out by direct observation of exponentially growing cells by
phase-contrast microscopy or by point inoculation of semisolid agar
(0.3% [wt/vol] agar, 0.03% [wt/vol] yeast extract, 0.03%
[wt/vol] NaCl, 0.03% [wt/vol] tryptone) to test for swarming.
Isolation of flagella and observation of filament
polymorphs.
A modified version of the method of Aizawa and
coworkers (1) was used for the large-scale isolation of
intact flagella for R. sphaeroides. Photosynthetic culture
(1.5 liters) at an optical density at 660 nm of 0.75 was harvested, and
the cells were lysed and treated with DNase as described previously
(1). The pH was increased to 10 by the dropwise addition of
5 M NaOH in order to eliminate contaminating membrane fragments.
Cellular debris was removed by centrifugation at 15,000 rpm for 45 min in Beckman JA 21 rotor. Flagella were collected from the supernatant by
centrifugation at 30,000 rpm for 30 min in a Beckman L7
ultracentrifuge. The flagellar pellet was resuspended in TET buffer (10 mM Tris-HCl [pH 8], 1 mM EDTA, 0.1% [wt/vol] Triton X-100). A
density gradient was set up by the addition of 0.43 g of CsCl per
ml of suspension and centrifugation at 20,000 rpm at 14°C overnight
in a Beckman L7 ultracentrifuge. The flagella formed a cloudy white
band, which was collected. CsCl was removed by a wash in 4 volumes of
TET buffer. The final pellet was resuspended in 0.5 ml of TET buffer. The flagella were observed by high-intensity dark-field microscopy. Broken, detached flagellar filaments were also harvested from stationary-phase cultures by a 30-min 30,000-rpm centrifugation step
and CsCl gradient (as detailed above). These were also used in
polymorphic studies to preclude any effects of flagellar basal bodies
(present in the intact preparation) on filament polymorphisms.
The buffers for observations of filament polymorphism at pH 2 to 8 were
prepared with citric acid and Na2HPO4 as
described previously (6). A second set of buffers from pH 7 to 11 were prepared by mixing 12.5 ml each of 0.2 M Tris and 0.2 M
glycine. The pH was adjusted with either 0.2 M HCl or 0.1 M NaOH before deionized water was added to a final volume of 50 ml. In all cases, an
appropriate volume of 5 M NaCl was added to give the desired concentration of NaCl. Equal volumes of buffer and flagellar solution were mixed and incubated at room temperature for 5 min before observation by high-intensity dark-field microscopy. The results were
recorded on videotape.
Recombinant DNA techniques.
Recombinant DNA techniques were
carried out as described by Sambrook and coworkers (31).
Restriction and modification enzymes were obtained from Northumbria
Biochemicals, New England Biolabs, and Boehringer Mannheim. DNA
fragments were purified using the Geneclean kit (Bio 101), and the
Photogene detection kit (Life Technologies) was used for Southern
blotting. R. sphaeroides genomic DNA isolations, DNA
sequencing, and conjugation protocols for complementation analysis were
as described previously (7, 27, 35).
Western blotting.
Sodium dodecyl sulfate-polyacrylamide gel
electrophoresis of whole cells or sheared filaments was carried out
according to the method of Laemmli as described by Sambrook and
coworkers (31). Sheared filaments were prepared from motile
bacterial cultures by 10 passages through a 3FG cannulum (Portex UK)
held between two 10-ml syringes. The proteins from the gels were
transferred to Hybond-C super nitrocellulose (Amersham) for 1 h at
a constant 100 mA. After being blocked overnight in PBS-T (140 mM NaCl,
2.7 mM KCl, 10 mM Na2HPO4, 1.8 mM
KH2PO4, 0.3% [vol/vol] Tween 20), containing
1% (wt/vol) milk (Marvel nonfat dried milk), the membrane was washed
in PBS-T for 5 min. It was incubated in a 1/1,000 dilution of antiserum
raised against R. sphaeroides flagellar filaments (36) in PBS-T, containing 1% (wt/vol) milk and 15%
(wt/vol) bovine serum albumin for 2 h. The membrane was washed
three times with PBS-T, containing 0.3% (wt/vol) milk, and then
incubated with a 1/1,000 dilution of anti-rabbit immunoglobulin
G-alkaline phosphatase conjugate (Sigma) in PBS-T containing 0.3%
(wt/vol) milk. After five washes in phosphate-buffered saline, the blot was developed by incubation in detection buffer (2.5 mg of
5-bromo-4-chloro-3-indolylphosphate [BCIP] ml
1, 5 mg of
nitroblue tetrazolium ml1, 100 mM Tris-HCl [pH 9.5],
100 mM NaCl, 5 mM MgCl2) until color development was
observed. The reaction was stopped by washing the blot in excess water.
Bioluminescence measurements.
Stationary-phase, nonmotile,
photosynthetic cultures (30-ml volume) were diluted by mixing them with
70 ml of fresh medium to introduce flagellar gene expression. Samples
were taken immediately and then every hour. Readings for light emission
were taken on a Turner luminometer. The light intensity per unit of
cell mass was calculated by dividing the luminometer reading by the
optical density of the culture sample at 600 nm.
Nucleotide sequence accession number.
The DNA sequence was
submitted to EMBL under accession no. Y14687.
| |
RESULTS |
Polymorphic ability of detached R. sphaeroides
flagella.
Figure 1 is a phase
diagram of the dominant forms observed under different conditions of pH
and ionic strength. Four forms for R. sphaeroides filaments
could be distinguished: straight, normal, open coils, and curly.
Examples are illustrated in Fig. 2. Both
intact flagella and broken detached filaments from R. sphaeroides gave the same results. The normal, straight, and
curly types resembled morphological types previously observed for the Salmonella filament (17). The coiled structures,
seen in this study and by Armitage and Macnab (2), are
referred to as "open coils" because they are distinct from the
coiled forms of Salmonella filaments observed by Kamiya and
Asakura (16). The difference is that the diameter of the
R. sphaeroides open coils is greater and the
Salmonella coil appears as a long cylinder when viewed from
the side whereas no such structures were visible for the R. sphaeroides open coils. They were found to resemble rope lasso structures rather than cylinders.
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FIG. 1.
Phase diagram showing the predominant polymorphic forms
of isolated R. sphaeroides flagellar filaments in
buffers of different pHs and ionic strengths.
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FIG. 2.
High-intensity dark-field microscope images of typical
polymorphic forms of R. sphaeroides flagella. (A)
Straight; (B) normal with some open coils; (C) open coils; (D) curly
(the inset shows greater magnification of the curly form). The open
coils and the normal forms were faint and highly susceptible to
Brownian motion, and therefore it was difficult to obtain good
micrographs of these from the videotape; some examples are shown.
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Interestingly, the dominant form under physiological conditions is the
open-coil form, which was also observed for cells with stopped flagella
(2, 3). This is in contrast to Salmonella i-flagella, which have a predominantly normal conformation, which undergoes a transition to curly at pH values lower than 4 in the presence of 0.1 M NaCl (16).
As found previously for Salmonella filaments
(16), the lines drawn between the types of
Rhodobacter filament conformations in Fig. 1 are not firm
boundaries. In many cases, different morphological types could coexist
under the same conditions (Fig. 2B), and the lines represent boundaries
where one form becomes dominant over all others. When normal filaments
were the predominant forms, a small proportion of open coils and
straight forms were observed. The curly and straight forms were never
observed together. At pH values lower than 3, the filaments were no
longer visible owing to the depolymerization of the filament into
flagellin monomers. Prolonged incubation (>15 min) at pH 3 also
resulted in depolymerization, and this effect was enhanced with
increasing NaCl concentration.
The range of forms obtained for the R. sphaeroides
filament is different from that obtained for the Salmonella
filament by Kamiya and Asakura (16). In order to determine
whether this is a result of gross differences between the amino acid
sequences of the flagellin proteins of the species, the fliC
gene of R. sphaeroides was cloned and sequenced as
described below.
Cloning, sequencing, and expression of R. sphaeroides
fliC.
The R. sphaeroides fliC gene was isolated by
complementation of a nonmotile, filament-minus TnphoA
mutant, R. sphaeroides Nm15 (37), with a
clone from a wild-type R. sphaeroides cosmid library.
Two kilobases of wild-type DNA from the cosmid clone that flanked the
site of TnphoA insertion in the mutant was localized by
Southern hybridization and sequenced on both strands. It was found to
encode one long open reading frame (ORF) starting at nucleotide 241, just after the HincII site (Fig.
3) and ending at nucleotide 1722. Eleven
bases upstream from the ATG start codon was the sequence AGGAGGG,
which matches the consensus sequence for ribosome-binding sites
in bacteria (9, 20). Upstream from that was a potential
promoter region with a version of the 
28 consensus
sequence TAAA(N14)GCCGTTGA. The DNA sequence was submitted to EMBL (accession no. Y14687), and the predicted amino acid sequence
from the ORF was used to search the EMBL database. The R. sphaeroides ORF showed homology with numerous flagellin
proteins, particularly flagellins from the plain flagella of
Bacillus subtilis (43% identity), Escherichia
coli and Salmonella enterica serovar Typhimurium (36%
identity), and Pseudomonas aeruginosa (41% identity) (Fig.
4). The amino acid sequence of R. sphaeroides flagellin, in common with those of other bacteria,
lacks cysteine and tryptophan (S. meliloti FlaA and FlaB are
the only ones that contain tryptophan), but it is unusual that it also
lacks proline. R. sphaeroides flagellin has very low
homology with the S. meliloti flagellins FlaA (27% identity) and FlaB (24% identity) from complex flagella. These low
levels of homology are not unexpected given that R. sphaeroides has plain flagella and that the unidirectionality
of flagella from both species is likely due to motor and not filament
properties, although 16S ribosomal DNA phylogenies show that S. meliloti and R. sphaeroides are closely related.
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FIG. 3.
Map of the region encoding R. sphaeroides
FliC. The putative 28 promoter region and the site of
cloning of the luxCDEAB reporter cassette used in expression
studies are shown.
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FIG. 4.
Prettybox multiple sequence alignment of R. sphaeroides FliC (flic_rsph) with FliCs from enteric and soil
bacteria. For reasons of space, much of the variable central region is
not shown. The regions predicted to form coiled coils are shown using
serovar Typhimurium coordinates (heavy bars) or R. sphaeroides coordinates (dotted line). Accession numbers are
as follows: S. meliloti FlaA (flaa_rhime), P13118; S. meliloti FlaB (flab_rhime), P13119; P. aeruginosa FlaA
(flaa_pseae), P21184; B. subtilis FliC (flic_bacsu), P02868;
serovar Typhimurium FliC (flic_salty), P06179; E. coli FliC
(flic_ecoli), P04949.
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To test for promoter activity, in the 600-bp region upstream from the
fliC ORF, a transcriptional fusion containing a promoterless Photorhabdus luminescens luxCDABE cassette (41)
was constructed. The cassette from pSB390 (G. S. A. B. Stewart and M. Winson, unpublished data) was cloned as a 5.8-kb
BamHI fragment into the BglII site of the
fliC gene in plasmid pRK415 to give pTP11. A second plasmid, pSB395 (41), which contains the lux cassette in
the same orientation and same vector as pTP11 but without any R. sphaeroides DNA, was used as a background control. The
bioluminescence levels of R. sphaeroides WS8N cultures,
containing each plasmid, were monitored over a time course after
dilution from stationary phase (as detailed in Materials and Methods).
The results are shown in Fig. 5. It is
clear that the region upstream from the fliC ORF has
promoter activity in R. sphaeroides. Interestingly,
although the (wild-type) R. sphaeroides WS8N cells
containing pSB395 were motile, the cells containing pTP11 were
nonmotile throughout the experiment, although the growth rates of the
two were similar (data not shown). There are two possible reasons for
this: (i) significant overexpression of the products of the
lux genes somehow interferes with motility, or (ii) the
promoter fragment of the plasmid sequesters some essential transcription factor(s) and prevents or reduces expression of chromosomally encoded fliC. The only identifiable promoter
consensus in the upstream 600-bp region is a sequence similar to the
28-dependent promoters of other species (Fig.
6), and so it may be 28
that is sequestered by the plasmid-borne promoter. Plasmid pTP11, containing the R. sphaeroides fliC promoter region, did
not give luminescence activity in E. coli (data not shown).
This may be due to slight differences between the 28
consensus in R. sphaeroides and that in E. coli (Fig. 6) (n = 14 bases and penultimate base
is G in R. sphaeroides, n = 15 bases and
penultimate base is A in E. coli).
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FIG. 5.
Promoter activity of the region upstream from the
fliC ORF as indicated by the luxCDEAB reporter
system. OD600, optical density at 600 nm.
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FIG. 6.
Consensus 28 recognition sequences
compared to the putative promoter sequence of R. sphaeroides
fliC. The B. subtilis 28 is also known
as D, and that of E. coli is known as
F (4, 12, 21).
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To test whether the polymorphic, plain filament R. sphaeroides flagellin could substitute for the flagellin in
E. coli plain filaments, the R. sphaeroides
fliC gene was cloned into pUC19 such that it was under the control
of the lac promoter. This plasmid, named pKa, was introduced
into two nonmotile fliC mutants of E. coli,
YK4146 and YK4516. R. sphaeroides fliC failed to
complement the E. coli mutations, and although R. sphaeroides FliC protein was detectable by Western blotting in
the cytoplasm of E. coli, it was not present in sheared
fractions or in the medium (data not shown). This result prevented us
from studying any polymorphic properties of hybrid E. coli-R.
sphaeroides flagella. It appears that the R. sphaeroides flagellin may not be exported by the E. coli flagellar export system.
| |
DISCUSSION |
The flagellar filament of R. sphaeroides
shows an interesting range of polymorphic transitions under varying
conditions of pH and ionic strength in vitro. Although the curly,
normal, and straight shapes are also seen with Salmonella
filaments (16, 17), the open-coil form is, as a naturally
occurring form, unique to R. sphaeroides filaments. The
open-coil form is the most common under physiological conditions and is
distributed across a broad range of conditions. Interestingly, in vivo,
stopped flagella have an open-coil shape whereas fast-rotating flagella
have a normal helical or apparently straight shape (2, 3).
It is thought that slow rotation of the coiled flagellum after a stop facilitates cell reorientation (3). This ability to change direction is essential during a tactic response since R. sphaeroides cells are unable to tumble in the manner of
bacteria with switching flagella. Thus, it appears that the filament of
R. sphaeroides has intrinsic properties that allow
polymorphic transitions adapted to the mode of swimming.
In serovar Typhimurium, mutations that affect the polymorphic ability
of the filament are found to cause amino acid substitutions in the
highly conserved N- and C-terminal regions of flagellin (19)
and flagellins with small deletions at either terminus are also
affected in their polymorphic ability (40). Direct interactions within the termini of flagella are important for the
polymorphic ability of the flagellar filament (25). Thus, terminal portions of flagellin are a determinant for polymorphic ability in serovar Typhimurium. In addition, a deletion in the variable
central domain of serovar Typhimurium flagellin also resulted in
alteration of polymorphic ability of the filament, possibly because a
change in the overall charge of the region altered repulsive or
attractive forces between subunits (26, 43). Such studies
suggest that a combination of interactions between the ordered termini
at the filament core and interactions between the outer domains of
flagellin molecules is responsible for polymorphic changes.
In order to test whether the differences in polymorphic abilities of
the filaments from R. sphaeroides and serovar
Typhimurium were reflected in differences at the molecular level, the
flagellin sequences of these two organisms were compared. The
hydropathy profiles of the two proteins are similar throughout, despite
the divergence of the R. sphaeroides sequence in the
central, nonconserved domain (data not shown). The terminal regions of
both proteins are predicted to have an alpha-helical character. The
serovar Typhimurium flagellin forms coiled-coil structures at the
termini (13, 28, 42). We analyzed both flagellins using the
COILS program (22) (Fig. 7).
As expected, serovar Typhimurium flagellin is predicted to have coiled
coils close to either terminus whereas R. sphaeroides
flagellin has a much lower probability of coiled coils in these
regions. This difference may be the key reason for the different
polymorphic abilities of filaments from these two species.
Interestingly, most of the mutations affecting polymorphic ability in
serovar Typhimurium flagella (19) map within, or close to,
these regions. Furthermore, many of these mutant sequences are
predicted to have altered coiled-coil probabilities compared to the
wild-type sequence according to the COILS program (data not shown).
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FIG. 7.
Prediction of coiled-coil structures at the termini of
flagellin proteins. The predictions were carried out using the COILS
program (22) with a window size of 28 residues. Solid line,
serovar Typhimurium FliC; dashed line, R. sphaeroides
FliC.
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The overall amino acid compositions of the nonconserved central regions
of FliCs (corresponding to amino acids 204 to 292 of serovar
Typhimurium), which comprise the outer domains in the assembled
flagellar filament, are similar in flagellins of R. sphaeroides and Salmonella. Therefore, the
intersubunit interactions in the outer parts of the filaments of these
species might be expected to be similar. This general assertion does
not seem to be borne out by the differences that we observed in
filament polymorphisms for the two bacteria. Mimori-Kiyosue and
coworkers (26) have found that the outer domain (D3) plays a
significant role in determining the polymorphic abilities of serovar
Typhimurium flagellar filament. It is possible that the central domain
plays no role in the differences in polymorphic ability seen here, or
subtle changes involving just a few amino acids in the D3 domain of
R. sphaeroides flagellin, rather than the overall amino
acid composition, are responsible for altered intersubunit interactions
that account for the unusual polymorphic abilities of R. sphaeroides flagellar filaments. It would be interesting to
exchange the central domains of Salmonella and R. sphaeroides flagellins and test whether the resulting
filaments had altered polymorphic abilities.
We have discussed some features that may be responsible for the
polymorphic properties of the R. sphaeroides filament.
The R. sphaeroides FliC is predicted to differ in its
secondary structure at the termini from the serovar Typhimurium
flagellin. It is also possible that a few key amino acid changes in the
nonconserved region of R. sphaeroides FliC cause changes
in intersubunit interactions. It would be interesting to see if these
predictions are borne out by a detailed structural study of the
R. sphaeroides filament.
It is difficult to speculate on the biological significance of the
unusual properties of the R. sphaeroides filaments. It is likely that the properties of the filament are adapted to its mode
of swimming motility, which in turn is adapted to its environment. R. sphaeroides inhabits aquatic or terrestrial
environments, whereas serovar Typhimurium is found mainly in intestinal
mucous surfaces. It may be that tumbling facilitates directional
changes in highly viscous mucous surfaces whereas in relatively
low-viscosity aquatic environments a combination of Brownian motion and
slow rotation of the filament open coil is sufficient to achieve a
change in orientation. Thus, the flagellar filaments of the two species may be adapted to these different requirements. Additionally, for
R. sphaeroides, having a single flagellum, which forms a
coil close to the cell body when stopped, may reduce the risk of being caught by protozoal predators in the wild. The single flagellum presents only a single receptor site for protozoa like
Acanthamoeba which bind to bacterial flagella
(30). There could be other explanations for the unusual
properties of R. sphaeroides FliC that will be
illuminated by a greater understanding of its natural history in the wild.
This study has provided some interesting insights into the properties
of the R. sphaeroides flagellum and a comparison of it
with the well-studied serovar Typhimurium flagellum. Understanding polymorphic transitions in different flagella and how they vary with
the chemical environment sheds light on the nature of FliC subunit
interactions in filaments. Such knowledge may have implications for the
design of nanofilaments with predictable physical properties.
This work was supported by grant no. F.114L from the Leverhulme
Trust to R.E.S. and by a Royal Society study visit grant to D.S.H.S.
and a BBSRC ISIS study visit grant to R.E.S. S.M.S. was supported
by an EC technical training scheme.
We thank Takuya Gotou for assistance with image processing and members
of the Aizawa and Sockett labs for useful discussions.
| 1.
|
Aizawa, S.-I.,
G. E. Dean,
C. J. Jones,
R. M. Macnab, and S. Yamaguchi.
1985.
Purification and characterization of the flagellar hook-basal body complex of Salmonella typhimurium.
J. Bacteriol.
161:836-849[Abstract/Free Full Text].
|
| 2.
|
Armitage, J. P., and R. M. Macnab.
1987.
Unidirectional, intermittent rotation of the flagellum of Rhodobacter sphaeroides.
J. Bacteriol.
169:514-518[Abstract/Free Full Text].
|
| 3.
|
Armitage, J. P.,
T. P. Pitta,
M. A.-S. Vigeant,
H. L. Packer, and R. M. Ford.
1999.
Transformations in flagellar structure of Rhodobacter sphaeroides and possible relationship to changes in swimming speed.
J. Bacteriol.
181:4825-4833[Abstract/Free Full Text].
|
| 4.
|
Arnosti, D. N., and M. J. Chamberlin.
1989.
Secondary factor controls transcription of flagellar and chemotaxis genes in Escherichia coli.
Proc. Natl. Acad. Sci. USA
86:830-834[Abstract/Free Full Text].
|
| 5.
|
Davis, J.,
T. J. Donohue, and S. Kaplan.
1988.
Construction, characterization, and complementation of a Puf mutant of Rhodobacter sphaeroides.
J. Bacteriol.
170:320-329[Abstract/Free Full Text].
|
| 6.
|
Dawson, R. M. C.,
D. C. Elliott,
W. H. Elliott, and K. M. Jones.
1986.
Data for biochemical research, 3rd ed.
Clarendon Press, Oxford, United Kingdom.
|
| 7.
|
Donohue, T. J.,
A. G. McEwan, and S. Kaplan.
1986.
Cloning, DNA sequence, and expression of the Rhodobacter sphaeroides cytochrome c2 gene.
J. Bacteriol.
168:962-972[Abstract/Free Full Text].
|
| 8.
|
Dons, L.,
O. F. Rasmussen, and J. E. Olesen.
1992.
Cloning and characterisation of a gene encoding flagellin of Listeria monocytogenes.
Mol. Microbiol.
6:2919-2929[Medline].
|
| 9.
|
Gold, L.,
D. Pribnow,
T. Schneider,
S. Shinedling,
B. S. Singer, and G. Stormo.
1981.
Translation initiation in prokaryotes.
Annu. Rev. Microbiol.
35:365-403[CrossRef][Medline].
|
| 10.
|
Gotz, R.,
N. Limmer,
K. Ober, and R. Schmitt.
1982.
Motility and chemotaxis in two strains of Rhizobium with complex flagella.
J. Gen. Microbiol.
128:789-798.
|
| 11.
|
Gotz, R., and R. Schmitt.
1987.
Rhizobium meliloti swims by unidirectional, intermittent rotation of right-handed flagellar helices.
J. Bacteriol.
169:3146-3150[Abstract/Free Full Text].
|
| 12.
|
Helmann, J. D.
1991.
Alternative sigma factors and the regulation of flagellar gene expression.
Mol. Microbiol.
5:2875-2882[Medline].
|
| 13.
|
Homma, M.,
D. J. DeRosier, and R. M. Macnab.
1990.
Flagellar hook and hook-associated proteins of Salmonella and their relationship to other axial components of the flagellum.
J. Mol. Biol.
213:819-832[Medline].
|
| 14.
|
Joys, T. M.
1985.
The covalent structure of the phase-1 flagellar filament protein of Salmonella typhimurium and its comparison with other flagellins.
J. Biol. Chem.
260:15758-15761[Abstract/Free Full Text].
|
| 15.
|
Joys, T. M.
1988.
The flagellar filament protein.
Can. J. Microbiol.
34:452-458[Medline].
|
| 16.
|
Kamiya, R., and S. Asakura.
1976.
Helical transformations of Salmonella flagella in vitro.
J. Mol. Biol.
106:167-186[CrossRef][Medline].
|
| 17.
|
Kamiya, R.,
S. Asakura, and S. Yamaguchi.
1980.
Formation of helical filaments by copolymerization of two types of `straight' filaments.
Nature (London)
286:628-630[CrossRef][Medline].
|
| 18.
|
Kamiya, R.,
H. Hotani, and S. Asakura.
1982.
Polymorphic transition in bacterial flagella, p. 53-76.
In
W. B. Amos, and J. D. Duckett (ed.), Prokaryotic and eukaryotic flagella. Cambridge University Press, Cambridge, United Kingdom.
|
| 19.
|
Kanto, S.,
H. Okino,
S.-I. Aizawa, and S. Yamaguchi.
1991.
Amino acids responsible for flagellar shape are distributed in terminal regions of flagellin.
J. Mol. Biol.
219:471-480[CrossRef][Medline].
|
| 20.
|
Kozak, M.
1983.
Comparison of initiation of protein synthesis in prokaryotes, eukaryotes and organelles.
Microbiol. Rev.
47:1-45[Free Full Text].
|
| 21.
|
Kutsukake, K.,
Y. Ohya, and T. Iino.
1990.
Transcriptional analysis of the flagellar regulon of Salmonella typhimurium.
J. Bacteriol.
172:741-747[Abstract/Free Full Text].
|
| 22.
|
Lupas, A.,
M. Van Dyke, and J. Stock.
1991.
Predicting coiled coils from protein sequences.
Science
252:1162-1164[CrossRef][Medline].
|
| 23.
|
Macnab, R. M.
1996.
Flagella and motility, p. 123-145.
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, 2nd ed. ASM Press, Washington, D.C.
|
| 24.
|
Macnab, R. M., and M. K. Ornston.
1977.
Normal-to-curly flagellar transitions and their role in bacterial tumbling. Stabilization of an alternative quaternary structure by mechanical force.
J. Mol. Biol.
112:1-30[Medline].
|
| 25.
|
Mimori-Kiyosue, Y.,
F. Vonderviszt,
I. Yamashita,
Y. Fujiyoshi, and K. Namba.
1996.
Direct interaction of flagellin termini essential for the polymorphic ability of the flagellar filament.
Proc. Natl. Acad. Sci. USA
93:15108-15113[Abstract/Free Full Text].
|
| 26.
|
Mimori-Kiyosue, Y.,
I. Yamashita,
Y. Fujiyoshi,
S. Yamaguchi, and K. Namba.
1998.
Role of the outermost subdomain of Salmonella flagellin in the filament structure revealed by electron cytomicroscopy.
J. Mol. Biol.
284:521-530[CrossRef][Medline].
|
| 27.
|
Moore, M. D., and S. Kaplan.
1989.
Construction of TnphoA gene fusions in Rhodobacter sphaeroides: isolation and characterization of a respiratory mutant unable to utilize dimethyl sulfoxide as a terminal electron acceptor during anaerobic growth in the dark on glucose.
J. Bacteriol.
171:4385-4394[Abstract/Free Full Text].
|
| 28.
|
Namba, K.,
I. Yamashita, and F. Vonderviszt.
1989.
Structure of the core and central channel of flagella.
Nature (London)
342:648-654[CrossRef][Medline].
|
| 29.
|
Pleier, E., and R. Schmitt.
1989.
Identification and sequence analysis of two related flagellin genes in Rhizobium meliloti.
J. Bacteriol.
171:1467-1475[Abstract/Free Full Text].
|
| 30.
|
Preston, T. M., and C. A. King.
1984.
Binding sites for bacterial flagella at the surface of the soil amoeba Acanthamoeba.
J. Gen. Microbiol.
130:1449-1458.
|
| 31.
|
Sambrook, J.,
E. F. Fritsch, and T. Maniatis.
1989.
Molecular cloning: a laboratory manual, 2nd ed.
Cold Spring Harbor Laboratory, Cold Spring Harbor, N.Y.
|
| 32.
|
Schmitt, R.,
I. Raska, and F. Mayer.
1974.
Plain and complex flagella of Pseudomonas rhodos: analysis of fine structure and composition.
J. Bacteriol.
117:844-857[Abstract/Free Full Text].
|
| 33.
|
Schmitt, R.,
I. Bamberger,
G. Acker, and F. Mayer.
1974.
Fine structure analysis of the complex flagella of Rhizobium lupini H13-3.
Arch. Microbiol.
100:145-162[CrossRef].
|
| 34.
|
Seville, M.,
T. Ikeda, and H. Hotani.
1993.
The effects of sugar on the morphology of the bacterial flagellum.
FEBS Lett.
322:260-262.
|
| 35.
|
Shah, D. S. H., and R. E. Sockett.
1995.
Analysis of the motA flagellar motor gene from Rhodobacter sphaeroides, a bacterium with a unidirectional, stop-start flagellum.
Mol. Microbiol.
17:961-969[CrossRef][Medline].
|
| 36.
|
Sockett, R. E., and J. P. Armitage.
1991.
Isolation, characterization, and complementation of a paralyzed flagellar mutant of Rhodobacter sphaeroides WS8.
J. Bacteriol.
173:2786-2790[Abstract/Free Full Text].
|
| 37.
|
Sockett, R. E.,
J. C. A. Foster, and J. P. Armitage.
1990.
Molecular biology of the Rhodobacter sphaeroides flagellum.
FEMS Symp.
53:473-479.
|
| 38.
|
Trachtenberg, S.,
D. J. DeRosier,
S.-I. Aizawa, and R. M. Macnab.
1986.
Pairwise perturbation of flagellin subunits. The structural differences between plain and complex bacterial flagellar filaments.
J. Mol. Biol.
190:569-576[CrossRef][Medline].
|
| 39.
|
Trachtenberg, S.,
D. J. DeRosier, and R. M. Macnab.
1987.
Three-dimensional structure of the complex flagellar filament of Rhizobium lupini and its relation to the structure of the plain filament.
J. Mol. Biol.
195:603-620[CrossRef][Medline].
|
| 40.
|
Voderviszt, F.,
H. Uedaira,
S. Kidokoro, and K. Namba.
1990.
Structural organisation of flagellin.
J. Mol. Biol.
214:97-104[CrossRef][Medline].
|
| 41.
|
Winson, M. K.,
S. Swift,
P. J. Hill,
C. M. Sims,
G. Griesmayr,
B. W. Bycroft,
P. Williams, and G. S. A. B. Stewart.
1998.
Engineering the luxCDEAB genes from Photorhabdus luminescens to provide a bioluminescent reporter for constitutive and promoter probe plasmids and mini-Tn5 constructs.
FEMS Microbiol. Lett.
163:193-202[CrossRef][Medline].
|
| 42.
|
Yamashita, I.,
F. Vonderviszt,
Y. Mimori,
H. Suzuki,
K. Oosawa, and K. Namba.
1995.
Radial mass analysis of the flagellar filament of Salmonella: implications for the subunit folding.
J. Mol. Biol.
253:547-588[CrossRef][Medline].
|
| 43.
|
Yoshioka, K.,
S.-I. Aizawa, and S. Yamaguchi.
1995.
Flagellar filament structure and cell motility of Salmonella typhimurium mutants lacking part of the outer domain of flagellin.
J. Bacteriol.
177:1090-1093[Abstract/Free Full Text].
|
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Abstract
Introduction
