INTRODUCTION

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Motile bacteria are propelled by helical flagellar filaments connected by a proximal hook to the basal body holding the flagellar rotary motor (for reviews, see references 1, 20, and 41). Traditionally, flagellar filaments are classified by their electron microscopical appearance into two types, named plain and complex (22, 33). The plain filaments of Escherichia coli and Salmonella spp. have a smooth surface structure with faint striations, whereas the complex filaments of soil bacteria, like Rhizobium lupini H13-3 and Sinorhizobium meliloti, exhibit a prominent helical pattern of alternating ridges and grooves (16, 32, 38). Unlike the flexible plain filaments, which are capable of switching from left-handed to right-handed helicity (21), the complex filaments are more rigid and do not switch handedness. Pairwise helical perturbations result in a subunit composed of a dimer of flagellin (39). Interflagellin bonds are believed to lock the complex filament in a rigid, right-handed helical conformation suitable for propulsion in viscous media (4, 8). Concomitantly, the flagellar motor of Rhizobium rotates entirely clockwise and does not reverse its sense of rotation (9). It has been shown that swimming S. meliloti cells respond to tactic stimuli by modulating their flagellar rotary speed (37) and that two novel motor proteins may be essential players in speed control (27). Hence, directional changes in the tracks of swimming S. meliloti cellsimperative for any chemotactic responseare a consequence of individual flagella rotating at different speeds (31). It thus appears that complex flagellar filaments and the new mode of directional control of swimming cells have evolved in response to the specific condition of swimming in viscous fluids prevailing in the soil biotope.

The flagellar filament consists of an assembly of about 20,000 flagellin subunits, whose molecular mass typically ranges from 40 to 60 kDa (20). Flagellins are three-domain proteins, with the N- and C-terminal domains being responsible for the quarternary interactions between subunits and the central, surface-exposed domain performing no obvious structural role but containing all of the potent antigenic epitopes. We have previously shown that the S. meliloti genome contains four genes, flaA, flaB, flaC, and flaD, encoding four related flagellin subunits (28, 36), and we report here three flagellin genes, flaA, flaB, and flaD, as constituents of the R. lupini H13-3 flagellar regulon. The latter strain was chosen because the first 13-Å-resolution, three-dimensional density map has been generated from its complex filament using low-dose electron micrographs of negatively stained specimens (4). This constellation may provide specific handles for future sequence-structure analysis.

In an effort to understand the process of assembling the complex filaments of the related soil bacteria R. lupini H13-3 and S. meliloti and to elucidate the contribution of single subunits to the filament structure, we have taken a genetic approach. Mutational analyses revealed that in both strains flagellin A is the principal, absolutely essential subunit but that, in addition, at least one of the secondary flagellin species is needed for assembling a functional filament. We also report that flagellin A biosynthesis is subject to control by transcriptional regulation.

	MATERIALS AND METHODS

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Bacteria and plasmids. Derivatives of E. coli K-12, R. lupini H13-3 (7), and S. meliloti MV II-1 (15) and the plasmids used are listed in Table 1.

Media and growth conditions. E. coli strains were grown in Luria broth (19) at 37°C. R. lupini and S. meliloti strains were grown in TYC (0.5% tryptone, 0.3% yeast extract, 0.13% CaCl₂ · 6H₂O [pH 7.0]) at 30°C (27). Motile cells prepared for functional tests were grown for 2 days in TYC, diluted in 15 ml of RB minimal medium (8) to an optical density at 600 nm (OD₆₀₀) of 0.05, layered on Bromfield agar plates (37), and incubated on a slowly rotating platform at 30°C for 16 h to an OD₆₀₀ of 0.2 to 0.5. The following antibiotics were used at the indicated final concentrations: for E. coli, ampicillin at 100 mg/liter, kanamycin at 50 mg/liter, and tetracycline at 10 mg/liter; for R. lupini and S. meliloti, neomycin at 120 mg/liter, streptomycin at 600 mg/liter and tetracycline at 10 mg/liter.

Gene replacement and reporter gene assay. Deletions (listed in Table 1) were generated in vitro by overlap extension PCR as described by Higuchi (12). Constructs containing the deletions were cloned into the mobilizable suicide vector pK18mobsacB, used to transform E. coli S17-1, and then conjugally transferred to R. lupini or S. meliloti by filter matings according to the method of Simon et al. (34). Allelic replacement was achieved by sequential selections on neomycin and 10% sucrose as described previously (37). Confirmation of allelic replacement and elimination of the vector was obtained by gene-specific primer PCR and Southern blotting. The broad-host-range plasmid pPHU234 and its derivatives pPHU235 and pPHU236 served as vectors for translational fusions of the four S. meliloti fla promoters. The resulting lacZ fusion plasmids were used to transform E. coli S17-1 and were then conjugally transferred to R. lupini RU12/001 or S. meliloti RU11/001 by streptomycin-tetracycline double selection, as described by Labes et al. (17).

DNA methods. R. lupini and S. meliloti DNAs were isolated and purified as described previously (37). Plasmid DNA was purified with NucleoSpin (Macherey Nagel, Düren, Germany). DNA fragments or PCR products were purified from agarose gels by use of a QiaEx DNA purification kit (Qiagen, Hilden, Germany). PCR amplification of chromosomal DNA and Southern blotting were carried out according to published protocols (27, 36). R. lupini genomic fragments containing the fla genes (Fig. 1) were isolated by plasmid insertion and rescue. An 850-bp flaA fragment and a 300-bp flaD fragment were PCR amplified and ligated into pRU1993, a derivative of the broad-host-range mobilizable vector pK18mobsacB containing the tetracycline resistance cassette of 2pRU738 (40). Recombinant plasmids were conjugally transferred and allowed to insert into the homologous R. lupini gene loci by a single crossover. Cells containing the insertion were selected on TYC plates containing neomycin and streptomycin. Chromosomal DNA was digested with MluI (flaA) or HindIII (flaD), ligated, and used to transform E. coli DH10B. Plasmid DNAs isolated from tetracycline-resistant transformants were physically mapped and sequenced with a model 310 automatic sequencer (Applied Biosystems, Weiterstadt, Germany). Sequences were aligned and compared by using Genetics Computer Group sequence analysis software (6).

View larger version (23K): [in this window] [in a new window]   FIG. 1.   Aligned partial maps of the S. meliloti (S.m.) (36), R. lupini (R.l.) (this work) and A. tumefaciens (A.t.) (5) flagellar gene clusters. Solid lines signify fully sequenced genomic regions, and dashed lines signify partially sequenced genomic regions. Gene loci are not drawn to scale; relevant transcription units and their polarities are marked by small dashed arrows, and general transcription polarity is marked by large arrows. The nomenclature of the S. meliloti flaC and flaD genes (36) has been changed in accordance with their map order, and the R. lupini flaD gene has been named by analogy to its A. tumefaciens paralogue (5). An R. lupini flaC paralogue was not detected. R. lupini (and A. tumefaciens) genes translocated and inverted with respect to their positions on the S. meliloti map are connected by dotted lines.

Chemotaxis assays. Swarm plates containing Bromfield medium and 0.3% Bacto Agar were inoculated with 3-µl droplets of the test culture and incubated at 30°C for 3 days. The speed of swimming cells was measured by using the computerized motion analysis of the Hobson BacTracker system (Hobson Tracking Systems Ltd., Leicester, United Kingdom) as described by Sourjik and Schmitt (37).

Flagellar filament isolation. Flagella were detached from motile cells (from 200-ml cultures) by agitation in a Braun MX32 mixer (Braun, Melsungen, Germany) at maximum power for 20 s, separated from cells by centrifugation at 8,000 × g, and purified by serial centrifugation at 8,000 × g for 8 min and at 15,000 × g for 15 min. Purified flagella sedimented at 87,000 × g for 2 h were washed once with a solution containing 0.5 mM CaCl₂, 0.1 mM EDTA, and 20 mM HEPES (pH 7.2) and resuspended in 150 µl of the buffer. Ninety-five percent of the final preparation was flagella, as assessed by electrophoresis in 10% acrylamide gels in the presence of sodium dodecyl sulfate.

Immunoblots. Polyclonal antibodies raised against purified R. lupini and S. meliloti whole flagellar filaments were isolated from whole serum by affinity purification. One milligram of isolated flagellar filaments was resuspended in 450 µl of sodium dodecyl sulfate sample buffer and heated to 100°C for 8 min. Fla proteins were separated electrophoretically in a 10% acrylamide gel by the method of Laemmli (18) and transferred to nitrocellulose (35). Blotted flagellins were cut into four pieces (5 by 20 mm²) to fit a 2-ml Eppendorf cup. Two milliliters of undiluted antiserum was added and incubated for 16 h at 4°C. Filter chips were washed three times with phosphate-buffered saline (PBS)-0.1% bovine serum albumin (BSA), twice with PBS-0.1% BSA-0.1% Nonidet P-40, and three times with PBS-0.1% BSA (5 min each). Bound antibodies were eluted from filter chips by mixing them carefully with 750 µl of 0.2 M glycine-HCl, pH 2.5, for 1 min. The supernatant was immediately added to 375 µl of prechilled KP_i (pH 9.0). The elution procedure was repeated twice, and the combined eluates were dialyzed three times against PBS. Whole-cell extracts were separated in a 10% acrylamide gel, transferred to nitrocellulose (as before), and probed as described previously (35) using purified anti-R. lupini or anti-S. meliloti Fla polyclonal antibody at a 1:500 dilution.

Protein and -galactosidase assays. Protein concentrations were determined by the Bradford microassay (Bio-Rad, Munich, Germany). Cultures of S. meliloti containing lacZ fusions were sampled, diluted in Z buffer (24) to an OD₆₀₀ of 0.2, permeabilized with 1 drop of toluene, and assayed for -galactosidase activity by the method of Miller (24).

Electron microscopy. Complex flagella were negatively stained with uranyl acetate (3%, wt/vol) on carbon-coated copper grids. Images were taken on a transmission electron microscope (Philipps model CM12) at 120 kV with a single-stage charge-coupled-device camera (Tietz, Gauting, Germany).

Nucleotide sequence accession numbers. The R. lupini fla sequences have been deposited in the GenBank database under accession no. AY7305, AY7306, and AY7307, and the S. meliloti fla sequences have been deposited under accession no. L49337.

	RESULTS

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Flagellin genes and deduced polypeptide sequences. Similarities between the fla genes of R. lupini H13-3 and S. meliloti (28, 36) were used in the design of flaA and flaD sequence-specific PCR primers (containing suitable restriction sites) that were used to amplify two gene-specific R. lupini DNA fragments. These were used as probes for plasmid rescue of two genomic, 10.7- and 8.7-kb regions containing flaA-flaB (pRU2351) and flaD (pRU2352), respectively. The R. lupini fla genes were completely sequenced and identified by the similarity of the encoded proteins to other flagellins, in particular those of S. meliloti (28, 36) and Agrobacterium tumefaciens (3, 5). Adjacent portions of the two clones were sequenced only to the extent where flagellar and motility genes could be identified and arranged on the map by similarity to their S. meliloti paralogues (36).

View larger version (75K): [in this window] [in a new window]   FIG. 2.   Comparison of the three R. lupini (R.l.) and four S. meliloti (S.m.) deduced flagellin polypeptide sequences. Numbering refers to amino acid residues in each line. Dashes signify identical residues with respect to the R. lupini FlaA sequence, and dots signify gaps. The consensus sequence includes all residues that form a homology group with a weighted relative frequency of 0.5 or greater. Conserved N- and C-terminal domains are boxed, and black bars denote amino acid residuces unique to right-handed helical flagellar filaments (41). Identity/similarity values (percentages) relative to R. lupini FlaA are listed at the end of each sequence.

View larger version (17K): [in this window] [in a new window]   FIG. 3.   Dendrogram based on full-length flagellin sequences from A. tumefaciens (Fla A. t.), R. lupini (Fla R. l.), and S. meliloti (Fla S. m.) and constructed by using the progressive, pairwise alignment method of PileUp from the Genetics Computer Group package (6). The Salmonella enterica serovar Typhimurium flagellin (FliC S. t.) sequence was used as an outgroup marker.

Mutational analysis of flagellin genes. To study the contribution of single flagellin subunits to flagellar filament function and structure, the three fla genes of R. lupini and the four fla genes of S. meliloti were separately inactivated by introducing defined deletions using allelic replacement (30). Although our previous results (28) and gene mapping (36) (Fig. 1) indicated that each fla gene is transcribed from its own promoter, in-frame deletions were generated throughout to minimize the chance of polar effects. Single- and multiple-allelic fla mutants were tested for their motile behavior and for flagellar morphology. Motile fitness was determined on swarm plates (by comparison to that of the wild type) and by computerized motion analysis of free-swimming cells (37). The averaged data of all R. lupini and S. meliloti strains tested are listed in Table 2. The table also lists features of flagellar appearance, as inspected by electron microscopy (Fig. 4). The wild-type specimen (Fig. 4A and D) exhibited the typical cross-hatched pattern reflecting the conspicuous helical undulations of complex flagellar filaments (32, 33). The two FlaA-deficient filaments shown failed to form proper helical filaments and were quite fragile (Fig. 4B and E), although their surface fine structure appeared to be normal (Fig. 4C and F).

View larger version (224K): [in this window] [in a new window]   FIG. 4.   Electron micrographs of R. lupini and S. meliloti wild-type and mutant flagellar filaments negatively stained with uranyl acetate. Wild-type R. lupini (A) and S. meliloti (D) complex flagellar filaments are dominated by a prominent three-start helical band pattern (zigzag pattern). Mutant filaments lacking the FlaA subunit from either R. lupini (B) or S. meliloti (E) are shown. Their complex surface structures appear to be normal (C and F). Bars: 100 nm (A, C, D, and F) and 500 nm (B and E).

Western blot analysis. Approximate amounts of flagellin produced by motile wild-type and fla mutant cells of R. lupini and S. meliloti were compared by Western blot analysis. Flagellin bands were detected by purified polyclonal antiflagellin antibody, as shown in Fig. 5. Wild-type R. lupini (Fig. 5A) presented two bands of 41 kDa (FlaA) and 42 kDa (FlaB). A third band, corresponding to FlaD, was not detectable, probably owing to low gene expression, and there was no indication whatsoever of a fourth flagellin subunit. Most striking was the decrease in flagellin in two mutants (flaAD and flaBD) (Fig. 5A) that could not assemble a functional filament, because either the principal FlaA subunit or the accessory FlaB and FlaD subunits were missing. This observation was augmented and confirmed by the immunochemical analysis of S. meliloti (Fig. 5B). In this species, FlaA and FlaB comigrated at 43 kDa, FlaC appeared as a weak, slightly slower-migrating band (Fig. 5B) (flaAB and flaBD), and FlaD was not detectable. Like in R. lupini, the amounts of Fla proteins synthesized decreased dramatically if the given subunit combination prevented the assemblage of a functional filament, notably, if FlaA and one or several secondary Fla subunits were lacking. Therefore, there must be a feedback regulation that reduces the rate of flagellin biosynthesis whenever an unproductive combination of Fla subunits prevents the assemblage of functional flagellar filaments. An apparent exception to this rule is the lack of reduction in (secondary) flagellin synthesis seen in flaA mutants of both R. lupini and S. meliloti (Fig. 5A and B). This lack of reduction is considered a consequence of transcriptional down-regulation that affects only flaA and not the other fla genes, as shown below (Table 3).

View larger version (15K): [in this window] [in a new window]   FIG. 5.   Immunoblot analysis of flagellin subunit proteins present in wild-type and mutant strains of R. lupini (A) and S. meliloti (B). Equal amounts of total cell protein (of ca. 107 cells contained in 20 µl at an OD600 of 0.3) of each strain listed were separated by denaturing gel electrophoresis, blotted on a nitrocellulose membrane, and detected with polyclonal antibody raised against purified flagella from R. lupini (A) and S. meliloti (B), respectively. Approximate band intensities are listed in Table 3.

Transcriptional control of fla genes. When the general picture of a feedback control of flagellin biosynthesis by nonfunctional subunit combinations emerged, we asked whether this regulation relates to fla gene transcription or not. Transcription from the four S. meliloti fla gene promoters was probed by using lacZ fusions in wild-type and fla mutant backgrounds. Table 3 lists the -galactosidase activities expressed as fractions of the wild-type activities of the four promoter fusion proteins measured in 14 tester strains. Absolute -galactosidase activities recorded for the wild-type background (Table 3) revealed that flaA has by far the strongest promoter exceeding the flaB and flaC promoter activities 6-fold and the flaD promoter activity nearly 100-fold.

	DISCUSSION

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In an attempt to relate the structure of the complex flagellar filaments of R. lupini and S. meliloti to their subunit compositions, we have used mutational analysis to study the contributions of the various flagellins to filament structure and to the regulation of flagellin biosynthesis. Of the three (R. lupini) or four (S. meliloti) flagellins, one (FlaA) is the principal subunit, while the others (FlaB, FlaC, and FlaD), though very similar, are accessory subunits. It takes a combination of FlaA and at least one accessory flagellin to assemble a functional filament. It is also the transcription of the principal FlaA-encoding gene (flaA) that is down-regulated whenever a nonproductive combination of Fla proteins prevents the assemblage of a functional filament. This type of control adds a new twist to the regulatory devices operating in bacterial flagellar synthesis.

The complex filaments of R. lupini H13-3 and, by analogy, those of S. meliloti differ from plain enterobacterial filaments by a pairwise perturbation of the helical lattice, resulting in a subunit composed of a dimer of flagellin (38, 39). This earlier observation has been related to the presence of several distinct flagellin subunits in the complex filament by the proposal of a structure composed of flagellin heterodimers (29). This concept has been strengthened by the present finding of a principal FlaA and several secondary FlaX subunits that together constitute a functional filament. In support of a heterodimer model is the need for a combination of FlaA and at least one accessory flagellin for assembling a functional filament. This assembly requires a FlaA-FlaX stoichiometry of 1:1. However, when we compared flaA promoter activity to the combined activities of the flaB, flaC, and flaD (= flaX) promoters (Table 3), the derived flaA-to-flaX transcript ratio was about 3:1, suggesting that, on average, only every second dimer in the filament was a FlaA-FlaX heterodimer. On the other hand, an inspection of band intensities on immunoblots (Fig. 5) suggested that, upon translation, FlaA and the secondary FlaX proteins result in a 1:1 ratio. This becomes particularly obvious in R. lupini, whose FlaA and FlaB proteins migrate at different mobilities (Fig. 5A). Also, in S. meliloti, a balanced FlaA-FlaX stoichiometry is evidenced by the flaA mutant which reveals roughly half the FlaX band intensity seen in the heavy single band of the wild-type strain (Fig. 5B). Such a balanced stoichiometry does, in fact, indicate a heterodimeric FlaA-FlaX substructure of the complex filament. It also implies an as yet unknown translational control of flagellin biosynthesis that equalizes the dominance of flaA transcription.

The transcription of the single flagellin gene, fliC, of E. coli proceeds from a particular promoter sequence, recognized by a specific sigma factor, ²⁸, encoded by fliA. This class II gene (20) is part of the hierarchial flagellar control system, expressed only when flagellar basal-body components are already being assembled. The fliA-encoded ²⁸ itself is antagonized by a specific anti-²⁸ factor, FlgM, which is eventually removed from the cytoplasm by export through the completed flagellar channel. FliA, which remains behind, is now capable of binding to the flagellin promoters. As reported here, the complex rhizobial filaments differ from plain E. coli filaments in their compositions by containing more than one type of flagellin. They also differ in the negative transcriptional control of flaA exerted by abortive combinations of Fla subunits that regulate FlaA biosynthesis in addition to the global control by VisNR (35). Can we envision a regulatory mechanism consistent with the current results? Upstream of the R. lupini and S. meliloti fla genes are sequences (delineated in Fig. 6) that resemble the ²⁸ consensus sequence derived for Vibrio parahaemolyticus (23) and the enterobacterial flagellar promoters (11). As previously elaborated by primer extension analysis of the flaA and flaB mRNAs of S. meliloti (28), these ²⁸-responsive elements represent the most likely candidates for fla promoters. Very similar promoter elements have been detected by sequence alignments upstream of all R. lupini and S. meliloti fla genes (Fig. 6). Although the 35 sequences of all seven fla genes are perfectly conserved, the 10 sequences deteriorate from flaA to flaD in comparison to the consensus sequences. Diminishing promoter quality is seen as the main reason for the decrease in transcriptional activity from flaA to flaD, as listed in Table 3.

View larger version (39K): [in this window] [in a new window]   FIG. 6.   DNA sequence alignment of putative promoter sequences upstream of the R. lupini (Rl) and S. meliloti (Sm) fla genes. flaA* denotes a secondary promoter sequence further upstream from the flaA gene (28). Capital letters indicate homology with consensus residues at a given alignment position. The R. lupini and S. meliloti (Rl/Sm) flagellin promoter consensus sequence was derived from a minimum plurality of four matches (except for position 3 of the 10 box, which was adapted to the highly active flaA promoter sequences). The consensus sequences of V. parahaemolyticus polar flagellar (VPA) (23) and E. coli 28 (11) promoters are shown for comparison.

A plausible model for the unusual down-regulation of flaA transcription reported here postulates jamming of the export channel by unproductive combinations of flagellins (missing the FlaA or the FlaX subunits) that cannot be assembled at the distal end. Rather than postulating a direct inhibitory interaction of excess flagellin accumulated in the cell with flaA transcription, we think that an FlgM-like anti-²⁸ protein (yet to be identified) is being prevented from escaping and, doomed to remain inside the cell, continues to render ²⁸-dependent transcription from the primary flaA promoter inactive. In order to substantiate this model, we are presently screening for mutants able to overcome the down-regulation of flaA. One may raise the objection that, different from the data of Table 3, an anti-²⁸ device should affect all four fla genes of S. meliloti, given their transcription from ²⁸-specific promoters. In principal, this is true. However, even at a maximum reduction to about 20%, flaA transcription still runs at a level above that of the standard transcription of flaB, flaC, or flaD (Table 3). It may be noted that there are secondary flaA promoter motifs located further upstream of the strong flaA* (28) promoters of both R. lupini and S. meliloti (Fig. 6); these may well account for the residual 20% transcription. We thus infere that down-regulation by nonassembled flagellin affects and reduces high-expression flaA but does not affect the secondary flaA* promoters and those of the lowly transcribed flaX genes.