The bacterial flagellum and flagellar motor: structure, assembly and function.

The Bacterial Flagellum and Flagellar Motor: Structure. Assembly and Function CHRISTOPHER J . JONES and SHIN-ICHI AIZAWA ERA TO. Research Development Corporation of Japan. 5-9-5 Tokodai. Tsukuba. Ibaraki 300-26. Japan

I. Introduction . . . . . . . . A . Chemotaxis . . . . . . . B . Flagellation . . . . . . . C . Motility . . . . . . . . D . Survival advantages of flagellation . . I1. Genetics . . . . . . . . . 111 Structure and function . . . . . . A . Filament . . . . . . . . B . Hook and hook-associated proteins . . C . Basalbody . . . . . . . D . Motandswitchcomplexes . . . . IV . Assembly . . . . . . . . . A . Filament . . . . . . . . B . Hook and hook-associated proteins . . C. Basalbody . . . . . . . D . Motandswitchcomplexes . . . . E . Other systems affecting flagellar assembly V . Motor function . . . . . . . . A . Parametersofmotorfunction . . . B . Models . . . . . . . . C . Recenttechnicaladvances . . . . VI . Summary . . . . . . . . . VII . Acknowledgements . . . . . . . References . . . . . . . . .

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ADVANCES IN MICROBIAL PHYSIOLOGY VOL .32 ISBN C-12-0277324

Copyright0 1991 .by Academic Press Limited All rightsof reproduction in any form reserved

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I. Introduction

One of the most striking features of bacteria is that they move. This movement is not random, but is modified in response to environmental stimuli such as light (Harayama and Iino, 1976; Taylor, 1983; Armitage et al., 1985), oxygen (Laszlo and Taylor, 1981; Taylor, 1983; Shioi etal., 1987), chemicals (Mesibov and Adler, 1972; Seymour and Doetsch, 1973; Tso and Adler, 1974), pH value (Tso and Adler, 1974), temperature (Maeda et al., 1976; Imae, 1985) and pressure (Li et al., 1988), and is called taxis. This tactic response enables bacteria to migrate continually to more favourable environments. Most motile species use one of two modes of motility, namely gliding and swimming. Gliding bacteria move over surfaces by a mechanism which is still not understood, but is thought to involve both motility organelles in the cell wall and a layer of extracellular slime (Burchard, 1981; Lapidus and Berg, 1982; Wolkin and Pate, 1984, 1986; Godchaux et al., 1990); the presumed motility organelles have yet to be identified. Bacterial swimming motility is much better understood. Many species of bacteria swim in liquid environments by means of rotating flagella, distinctive organelles which have been extensively studied. The bacterial flagellum, so-called on the basis of its resemblance to the eukaryotic organelle, differs fundamentally from the latter: the bacterial flagellum has a diameter of only about 20 nm, and consists primarily of a single protein, flagellin. It does not beat, but rotates, and is powered not by ATP but by the transmembrane proton gradient. In this review, we discuss the bacterial flagellum and flagellar motor, focusing on the structure, assembly and function of the organelle. Because much of the relevant work has been carried out using Escherichia coli and Salmonella typhirnurium, most of our discussion will revolve around the flagella of these species. There is some inconsistency in the use of the term “flagellum” in the context of the bacterial organelle. We use “flagellum” to refer to the entire organelle as currently defined morphologically (i.e. the filament, hook and basal body), and reserve the use of “filament” for that portion of the flagellum distal to the hook. The “flagellar motor” is necessarily defined only operationally, as the motor has not yet been described biochemically or morphologically. A. CHEMOTAXIS

Salmonella species and E . coli are enterobacteria which can colonize the human intestine. Observation of these cells by dark-field microscopy shows (Macnab and that they swim very fast, at speeds of about 25 l m s

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Koshland, 1972; Macnab and Ornston, 1977), interrupted every second or so by brief episodes (about 0.1 second) of chaotic tumbling. After each tumbling episode, the cell swims off in a new direction (Fig. l(a)). The path that the cell follows is a random walk in three dimensions (Berg and Brown, 1972; Berg, 1988). The application of positive stimuli (such as the addition of an attractant or removal of a repellent) suppresses the tumbling episodes, so that the cell swims in a relatively straight trajectory for longer periods (Berg and Brown, 1972). The trajectory of the cell in a stimulus gradient thus becomes a biased random walk (Fig. l(b)). Drastic increases in negative stimuli (removal of an attractant or addition of a repellent) cause an increase in the frequency of tumbling episodes, randomizing the direction of travel of the cell more often (Macnab and Koshland, 1972). (a1

FIG. 1. Paths followed by swimming bacterial cells. (a) In the absence of a stimulus gradient, a bacterium swims in straight paths called runs (lines) interrupted by brief tumbles (vertices) which re-orient the cell. (b) In an attractant gradient, runs in a positive direction (towards higher concentrations of attractant) are extended, whereas those in a negative direction are not. The cell thus executes a biased random walk in three dimensions, migrating towards more favourable environments. The open arrow indicates the direction of the gradient. From Macnab (1979a).

However, in the sorts of relatively shallow gradients likely to be encountered by the cell, it does not increase its tumbling frequency when swimming in a “negative” direction; under these conditions the cell shows roughly the same pattern of swimming and tumbling as it does in an isotropic environment (Berg and Brown, 1972; Brown and Berg, 1974). Increasing its frequency of tumbling, hence re-orienting, would do the cell no good as it

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would not have enough time to “decide” whether its new direction of travel was better or worse than the previous one (Berg and Purcell, 1977). This modulation of tumbling frequency forms the basis of taxis. While heading in a favourable direction, cells tend to swim straight ahead; when heading in an unfavourable direction, cells make frequent course changes, eventually finding a better direction of travel. Bacteria are too small to be able to make use of spatial gradient information. Escherichia coli and S. typhimurium instead integrate the information temporally, comparing their previous and current circumstances; bacteria are thus capable of remembering, albeit for a very short time (Macnab and Koshland, 1972; Brown and Berg, 1974). For a more detailed treatment of the physics of chemoreception, see the article by Berg and Purcell (1977). Our understanding of the molecular basis of chemotaxis has increased greatly in recent years; for further information regarding this system, see the reviews by Macnab (1987b), Stewart and Dahlquist (1987), Hazelbauer (1988) and Koshland (1988). Specific aspects of bacterial chemotaxis have recently been reviewed by Ames et af. (1988), Hess et al. (1988), Koshland et al. (1988), Stewart etal. (1988) andstocketal. (1988). For areview of earlier work in bacterial chemotaxis, see Berg (1975). Briefly, during chemotaxis, attractant and repellent molecules are bound

FIG. 2. Flow of information in the bacterial chemotaxis system. Binding of stimulus molecules by receptors in the inner membrane causes activation of CheA by Chew. CheA in turn phosphorylates the methyltransferase CheB, which completes a feedback loop to “reset” the receptor, and CheY which binds to the switch complex of the flagellar motor and causes clockwise rotation. CheZ antagonizes CheY action at the motor, probably by dephosphorylating CheY. The diagram is modified from Hess et al. (1988), based on data from Borkovich et al. (1989) and Liu and Parkinson (1989).

THE BACTERIAL FLAGELLUM AND FLAGELLAR MOTOR

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a

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FIG. 3. Bacterial flagella and hook-basal body (HBB) complexes. (a) A Salmonefia typhimuriurn cell, showing peritrichous arrangement of the flagella. (b) The cellproximal portion of a single isolated flagellum. (c) Purified HBB complexes. Note that, in addition to the complexes, hook fragmentsare also present. All samples were stained with 2%phosphotungstic acid. The scale bars represent 1 pm, 0.5 pm and 100 nm, respectively.

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either by receptor molecules in the inner membrane of the cell or by periplasmic binding proteins which themselves interact with the receptors. This binding probably induces a conformational change in the cytoplasmic domain of the receptor. This change then sets off a chain of phosphorylation reactions affecting many of the chemotaxis proteins (Fig. 2). The ultimate phosphorylated species, CheY, is the signal which binds to the flagellar motor, causing it to rotate clockwise and the cell to tumble (see below). B. FlAGELLATlON

Under the microscope, flagellated cells of E. coli or S. typhimurium can be seen to have five to ten long, thin, sinusoidal filaments extending from random points on the cell surface; this pattern of flagellation is called “peritrichous” (Fig. 3(a)). Other species may have single or multiple flagella at one or both ends of the cell (see, for example, Leifson, 1960; Buchanan and Gibbons, 1974); some species can have either a single polar flagellum or numerous lateral flagella, depending on conditions (Allen and Baumann, 1971; Ulitzer and Kessel, 1973). The flagella of Spirochaeta spp. are located entirely in the periplasmic space, between the inner and outer membranes (Canale-Parola, 1978; Holt, 1978). Some species have “sheathed” flagella, which are external flagella consisting of an inner core, similar to the plain flagella of bacteria such as E. coli and S . typhimurium, encased in an extension of the outer membrane (Fuerst and Hayward, 1969; Yang et al., 1977; Thomashow and Rittenberg, 1985b; Fuerst and Perry, 1988; see also Fuerst, 1980). Electron-microscope examination of the flagellum has shown that it appears to consist of three sections, namely the complex basal body, which is embedded in the cell wall, the long, helical filament and the short, curved hook connecting the two (Fig. 3(b)). The basal body is composed of several rings threaded onto a short rod (Fig. 3(c)). Although the size and number of these rings vary among different bacterial species (DePamphilis and Adler, 1971b; Coulton and Murray, 1978; Johnson et al., 1979; Swan, 1985; Thomashow and Rittenberg, 1985a; Kalmokoff et al., 1988; Kupper et al., 1989), this general structure for the flagellum is conserved in all flagellated bacteria in which it has been examined. Although the hook and filament appear to be directly connected, there are actually two small sections composed of the proteins HAP1 and HAP3 joining them; additionally, a cap of HAP2 protein is at the distal end of the filament (“HAP” stands for “hook-associated protein”). Even though the HAP proteins are part of the flagellar filament, we will discuss their structure and assembly together with those of the hook. Most of the flagellum lies beyond the inner membrane of the cell. The


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component proteins of those basal body rings which are outside this membrane are exported by the conventional protein-export pathway, but the proteins which make up the axial components of the flagellum are exported by a flagellar-specific pathway which involves their passage through the hollow core of the flagellum itself. C. MOTILITY

Bacterial flagella rotate. The torque for this rotation is generated by a rotary motor at the base of the structure. This motor is powered by the flow of protons across the cytoplasmic membrane and can rotate clockwise and counterclockwise. The flagellar motor thus converts the chemical energy of the transmembrane proton potential to mechanical work performed on the environment. In alkalophilic bacteria, sodium ions power the motor, but the mechanism of motor function is presumed to be essentially identical. Flagellar filaments are rigid left-handed helices (Macnab and Koshland, 1974); counterclockwise rotation of these helices (viewing along the filament axis toward the cell body) imparts translational motion to the cell. The competing forces generated on the cell body by different flagella will cause it to begin to move in some direction; hydrodynamic forces will then sweep all of the filaments into alignment, usually along the cell axis (Macnab and Ornston, 1977). This alignment is facilitated by the flexible hook which connects the filament to the motor. The flagellar bundle thus formed appears as a tail behind the swimming cell (Fig. 4(a,b)). When a sufficient number of flagella reverse their sense of rotation to clockwise, this bundle flies apart and the cell tumbles chaotically (Macnab and Koshland, 1974). In response to the reversal of torque on the filament caused by reversal of the rotation sense of the motor, the filament adopts a new, right-handed helical shape with a smaller pitch and helical diameter (Fig. 4(c,d)). This new conformation propagates distally along the filament and prevents the bundle from jamming; it probably also facilitates dispersal of the bundle. For reviews of bacterial motility, see the articles by Macnab and Aizawa (1984), Eisenbach (1990) and Khan (1988, 1990); for a review of earlier work in the field, see the article by Berg (1975). Flagellar structure has been reviewed recently by Macnab and Aizawa (1984), Iino (1985), Macnab (1987a) and Macnab and DeRosier (1988). D. SURVIVAL ADVANTAGES OF FLAGELLATION

There is an increasing body of evidence that flagella and chemotactic ability confer decided survival advantages on the cell (see, for example, Chet and Mitchell, 1976; Walsh and Mitchell, 1978). This has been demonstrated for a

FIG. 4. Swimming and tumbling Salmonella typhimurium. (a) Dark-field micrograph of swimming cells; note the bundles of flagellar filaments which appear as “tails” behind the cells. (b) Schematic diagram of a swimming cell; the solid arrow indicates the direction of wave propagation in the bundle and the open arrow indicates the direction of swimming. (c) Dark-field micrograph of a bacterium showing a heteromorphous filament which has a curly segment (with a smaller pitch and helical diameter than normal; see Fig. 7) proximal to a normal segment, with a sharp angle in the helical axis at the junction. Such heteromorphous filaments are crucial to the tumbling response. (d) Schematic diagram of a tumbling cell. The solid arrows indicate the direction of wave propagation for the individual filaments, and the open arrow indicates the tumbling motion of the cell body. The bar represents 5 pm. From Macnab and Ornston (1977) and Khan et al. (1978).


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number of Pseudomonas species, where motility or chemotactic capabilities have been shown to provide benefits in competition for nitrogenous soil compounds (Kennedy and Lawless, 1985), in growth on leaf surfaces (Haefele and Lindow, 1987) and in invasive virulence in mice (Drake and Montie, 1988). Flagellation also enhances the pathogenicity and colonizing ability of Vibrio cholerae (Yancey et al., 1978; Attridge and Rowley, 1983; Pierce et al., 1988). In S. typhimurium, on the other hand, flagella appear not to play a role in virulence (Lockman and Curtiss, 1990). The importance of the flagellar system to the bacterium may be estimated by looking at the cost of its maintenance. For S . typhimurium, a typical cell in exponential growth has about eight flagella, each 5-10 pm long. These flagella will contain of the order of 140,000 copies of flagellin alone (Namba et al., 1989), about 8% of total cell protein (Neidhardt, 1987). Approximately 2% of the genome is devoted to flagellation and motility. More than half the species of all bacterial families are flagellated (Buchanan and Gibbons, 1974). The prevalence of such an expensive system argues strongly for its utility in the survival of these organisms. 11. Genetics

There are currently about 40 genes known to code for components of the flagellar and motility systems of E . coli (Komeda et al., 1977b, 1980; Bartlett and Matsumura, 1984; Kuo and Koshland, 1986) and S. typhimurium (Yamaguchi etal., 1972,1984b; Kutsukake etal., 1980; Hommaetal., 1988), with another ten or so coding for components of the chemotaxis system (see, for example, Parkinson, 1977). These genes are listed in Table 1. Functional homology between almost all of the corresponding genes in E. coli. and S . typhimurium has been demonstrated (DeFranco et al., 1979; Kutsukake et al., 1980; Yamaguchi et al., 1984b). Genes which are required for formation of the flagellum are collectively called flagellar @a) genes.The products of two genes (motA and motB) function solely in energizing the motor. The flagella of Mot- cells appear normal and rotate freely in response to external forces, showing that the flagellar motors are not simply jammed and that the defects must therefore be in the energy transduction process (Tshihara et al., 1981; Yamaguchi et al., 1986a). Genes which encode components of the chemotaxis system are designated che; the flagellar motors of che mutants are functional, but have altered switching probabilities, and the cells are therefore unable to respond properly to stimuli (see, for example, Parkinson, 1977; Macnab, 1987b). The flu, mot and che genes (excepting some of the chemotactic receptor genes) comprise about 13 operons which are clustered in three regions of the

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TABLE 1. Flagellar and motility genes and gene products of Escherichia coli and Salmonella typhimurium Gene symbol"

v

PgB PgC PgD PgE PgF PgG PgH Pgl PgJ

Gene product Function and comments Necessary for addition of P ring Rod structural protein Rod structural protein Completes rod structure Hook structural protein Rod structural protein Distal rod structural protein L ring structural protein P ring structural protein Unknown HAP1 structural protein; junction protein between hook and filament HAP3 structural protein; junction protein between hook and filament Unknown Unknown Unknown Antagonizes CheY function at the flagellar motor, probably by dephosphorylation of CheY, restoring counterclockwise rotation Causes clockwise rotation of the flagellar motor, probably by binding as a phosphorylated species Methylesterase; modifies receptors to adapt cell to current stimulus level Methyltransferase; modifies receptors to adapt cell to current stimulus level Receptor for dipeptides; not present in S. typhimurium Receptor for aspartate, maltose, Co2+ and NiZ+ Enhances CheA autophosphorylation; necessary for transduction of information from receptors to CheA Autophosphorylates; transfers phosphate to CheY and CheB, presumably activating them Necessary for motor rotation; site limited when overproduced Necessary for motor rotation; not site limited when overproduced Sigma factor for transcription of level-I1 operons Sigma factor for transcription of level-I1 operons Sigma factor for transcription of level-I11 operons N-Methylation of certain lysine residues of flagellin; not present in E . coli Filament structural protein (flagellin) HAP2 structural protein; caps the distal end of the filament Unknown (RflA?) Unknown (RflA?) Basal-body component


Gene symbol" PiF fliC fli H fli K fli L

fliN fli0 fli R

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Gene product Function and comments M-ring structural protein Switch component of motor Unknown Unknown Unknown Hook-length control Unknown Switch component of motor Switch component of motor Unknown Unknown Unknown Unknown

Salmonella @-region genes Repressor of fliC transcription Filament structural protein (flagellin) Invertase controlling flagellar phase variation; hin transcription 3'T hin independent of FlhD expression

1%

a The arrows to the left of the gene symbols indicate the extent and direction of transcription of the operons containing these genes. The direction of transcription of the r f B and piB operons is not known; hin is transcribed in either direction, depending on the orientation of the invertible segment on which it resides (see Section 11). * In E. coli,region I is at chromosomal map position 24 min and regions 11 and 111 are at positions 41 min to 43 min (Bachmann, 1983); in S. typhimurium, region I is at map position 23 and regions I1 and 111 are at position 40 rnin (Sanderson and Roth, 1988). Theflj genes are at position 56 min in the genome of S. typhimurium (Sanderson and Roth, 1988). ' In addition to tar (and rap) in region 11, genes for other receptors are located elsewhere on the chromosome; see, for example, Macnab (198%).

chromosome (Table 1; Komeda et al., 1980; Kutsukake and Iino, 1985; Kutsukake et al., 1988). Flagellar genes in region I are identified with the gene designationfig, those in region I1 with@, and those in region I11 with pi. Region IT also contains motA and motB as well as all of the che genes. (The gene designations for the flagellar systems of E. coli and S. typhimurium have recently been unified and brought into accordance with accepted conventions (Iino et af., 1988), so that most of the literature in this field uses a different nomenclature.) The flagellar system of Salmonella spp. differs from that of most E. coli strains in having one additional region, which contains genes designated fEj, that are involved in phase variation (see Table 1;Lederberg and Iino, 1956). The existence of phase variation in some strains of E. coli has recently been demonstrated (Ratiner, 1982, 1985, 1987) and seems to operate by a mechanism similar to that in Salmonella spp.

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The genesflic andfljI3 encode distinct flagellins, each of which can form a functional flagellar filament. Inversion of a small (995 bp) segment of DNA upstream of fljB aligns a promoter and J j B , allowing synthesis of the FljB flagellin and of FljA, which represses fliC transcription (Zieg et al., 1977; Silverman et al. , 1979; Zieg and Simon, 1980). This inversion is mediated by the product of hin (Silverman and Simon, 1980; Kutsukake and Iino, 1980), which is on this segment (Zieg and Simon, 1980). Re-inversion of this region turns off transcription of bothflj genes, allowing the cell again to synthesize FliC flagellin. The result of this switching is that any wild-type population of Salmonella sp. will contain cells displaying either of two distinct flagellar serotypes; this presumably aids in survival of the population in the host. Because the flagellum is a complex structure, and the biosynthetic cost of its construction of the organelle is high, it is not surprising to find that there is

c AM P/C A P

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fliF

fliC

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FIG. 5. Positive and negative transcriptional control in the flagellar regulon. Each operon is denoted by the first gene transcribed in that operon (see Table 1). Transcription of thePhD operon (flagellar master operon) is activated by cAMPvia a complex with CAMP-binding protein (CAP). Transcription of the level-I1 operons requires the products of the master operon, which are believed to act as a flagellar gene-specific sigma factor (Helmann et al., 1988). Transcription of the level-111 operons, in turn, requires transcription of all of the level-I1 operons; the level-I1 product FliA is a sigma factor required for level-111operon transcription, but how the effect of the other level-I1 genes is mediated is unknown (Kutsukake et al., 1990). One of the downstream genes in thePiD operon @is orfiiT) probably codes for the factor responsible for repression of all level-111 operons (RflA).


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a hierarchy of control of flagellar synthesis at the transcriptional level preventing wasteful synthesis of components which cannot be used (Fig. 5 ) . The transcription of all flagellar and motility genes, as well as at least some chemotaxis genes, depends on the products of flhC and JlhD, which together constitute the flagellar master operon (Silverman and Simon, 1974b; Komeda, 1982). FlhC and FlhD are believed to act as an alternative sigma factor for RNA polymerase (Bartlett et al., 1988; Helmann et al., 1988; Arnosti and Chamberlain, 1989). Using the operon-fusion technique, Komeda (1982) and Kutsukake et al. (1990) have shown that transcription of most flagellar genes is under the direct control of the master operon. Transcription of these “level-11” genes requires only FlhC and FlhD, but transcription of the genes for components which function only relatively late in flagellar formation, the HAP proteins, Mot proteins and flagellin, depends on the product of the level-I1 genefliA. FliA is a sigma-factor specific for these late, “level-111”, genes (Kutsukake et al., 1990; Ohnishi et al., 1990). The level-I11 genes probably require FliA only for efficient transcription, as the HAP proteins were originally purified from afliA mutant (Homma et al., 1984a). In addition, transcription of level-I11 genes requires transcription of all of the level-I1 genes, not justfEiA (Komeda 1982,1986; Kutsukake ef al., 1990). Komeda (1986) suggested that this was a result of the protein FlgA serving both as a repressor of late operon transcription and as a structural component of the flagellum; normally FlgA would be sequestered in the flagellum, but a mutation in any level-I1 gene which prevented assembly would leave FlgA free to repress the late operons. The data of Kutsukake et al. (1990) conflict with this interpretation, however, so that the question of how so many gene products exert an effect on level-111 transcription is unresolved. Comparison of the promoter sequences for the various flagellar operons supports the arrangement of the flagellar genes into this simple three-level transcriptional cascade (Kutsukake et al., 1990). Although the cascade originally proposed for E. coli is somewhat different (Komeda, 1982,1986), it is likely that the scheme proposed for S. typhimun’urnalso applies in E. coli (Kutsukake et al., 1990). Negative transcriptional control is mediated by RflA (“Rfl” stands for “repression of flagellar operons”; Kutsukake et al., 1990). Mutations in @A result in enhanced transcription of level-I11 operons (Kutsukake et al., 1990); $A may correspond to either JEiS or fliT, newly-identified genes in thefliD operon (I. Kawagishi and R. M. Macnab, personal communication; see Table 1). The master operon itself is positively regulated by CAMP (Yokota and Gots, 1970; Silverman and Simon, 1974b; Komeda etal., 1975), mediated by

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a single CAP-binding site upstream offlhD (Bartlett et al., 1988). Because E. coli (although not S. typhimurium) is subject to catabolite repression, regulation by CAP and CAMPmeans that flagellar synthesis is abolished in nutrient-rich environments, where motility is not needed. This repression of flagellar synthesis can be overcome by the presence of chemotactic repellents (Vorobyeva et al., 1982). There is evidence for post-transcriptional control in the flagellar system. Ribonuclease I11 has been shown to be involved in flagellar regulation (Apirion and Watson, 1978), and DNA sequencing of flagellar genes has revealed putative regulatory features, both transcriptional and translational (Dean et al., 1984; Stader et al., 1986; Macnab, 1988; Jones et al., 1989; Kihara et al., 1989; Malakooti et al., 1989; Homma et al., 1990a,b; Kutsukake et al., 1990). With the cloning and ongoing sequencing of the flagellar genes, more precise investigations into the genetic regulation of this system should be forthcoming. 111. Structure and Function A. FILAMENT

The flagellar filament is a rigid helical tube, typically 5-10 pm long and with an apparent outer diameter of about 20 nm (Fig. 3(b)). In most bacterial species it is composed of a single protein species, flagellin. Some species possess flagella composed of multiple flagellins (see, for example, Alam and Oesterholt, 1984; Thomashow and Rittenberg, 1985a; Brahamsha and Greenberg, 1988; Driks et al., 1989; Gerl et al., 1989). Flagellin varies in molecular weight, both between and within species, ranging from 25,000 to 69,000 (Lagenaur and Agabian, 1976; Lawn, 1977; Ibrahim et al., 1985; Joys, 1988); not surprisingly, the differences within species are reflected in the antigenic determinants of each flagellin and form the basis of Hserotyping (Lawn, 1977; Ibrahim et al., 1985; Joys, 1988). Filaments can be purified by shearing them from the cells and pelleting them by high-speed centrifugation. Such a preparation is sufficiently pure for many purposes. For purer preparations, such filaments can be depolymerized by treating them with heat or acid (Kobayashi et al., 1959; Ada et al., 1963; Asakura et al., 1964) and the resulting flagellin monomers purified by high-performance liquid chromatography and repolymerized (Vonderviszt et al., 1989; see Section 1V.A). The resulting repolymerized structures are morphologically and functionally identical to native filaments (Abram and Koffler, 1964; Asakura et al., 1964; Iino et al., 1972). Under normal conditions the filament helix is left-handed, with a pitch of


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about 2.5 pm and a helical diameter of about 0.6 pm (see, for example, Kamiya et al., 1982). Anticlockwise rotation of this left handed helix exerts force on the cell body, resulting in propulsion. Note that a tube constructed from identical elements in identical interaction cannot be other than a straight cylinder. In order to construct a helical tube, the flagellin monomers from which it is assembled must be in non-equivalent interaction, even though their primary structure is identical (Caspar and Klug, 1962). 1. Filament Polymorphism

The filament can adopt any one of several discrete helinal shapes under appropriate conditions, a phenomenon called polymorphism (Asakura, 1970; Asakura and Iino, 1972; Iino et al., 1974). Although the normal filament helix is left-handed, many of its polymorphs are right-handed and all of them have different values for pitch and helical diameter (see, for example, Hotani, 1976; Kamiya and Asakura, 1976; Kamiya et al., 1982). On the basis of their appearances, these polymorphic forms are called, for example, “curly” and “semi-coiled” (Leifson, 1960). Reversible transitions may be initiated in vitro by changes in solution pH value, ionic strength or temperature, by application of viscous flow or an electric field, or by addition of organic solvents (Kamiya and Asakura, 1976, 1977; Kamiya et al., 1979; Hotani, 1980, 1982; Hasegawa et al., 1982; Washizu et al., 1989), and in vivo by reversal of the direction of flagellar rotation, and hence twist on the filament (Macnab and Ornston, 1977). This latter shape change occurs as a distally propagating zone of right-handed helix whose axis is at an angle to that of the remaining (original) left-handed helix (Macnab and Ornston, 1977). This drastic change in quaternary structure probably explains why filaments in the bundle do not jam upon reversal of rotation, as might be expected (Macnab, 1977), and serves to help disperse the bundle (Macnab and Ornston, 1977). Changes in flagellin structure, either by mutation (Iino, 1962; Iino and Mitani, 1966,1967; Martinez et al., 1968; Iino et al., 1974;Yamaguchi et al., 1984a) or the use of amino-acid analogues (Kerridge, 1959, 1960; Iino, 1969), can result in the filament having one of the other polymorphic shapes under normal conditions. Filaments constructed by copolymerizing different types of monomers in vitro (see Section 1V.A) also have different shapes, depending on the ratio of monomers used (Asakura and Iino, 1972; Kamiya et al., 1980). Such filaments can undergo polymorphic transition (Kamiya and Asakura, 1976,1977; Matsuura et al., 1978; Hasegawa et al., 1982).

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1-Start

0

FIG. 6. Diagram showing the surface lattice of the flagellar filament. Each flagellin monomer is represented as a solid circle. The major helical families are indicated; note the basic (1-start) helix, along which monomers are assembled sequentially, and the 11-start helix, which is nearly parallel to the filament axis and comprises the “protofilament .” Co-operative conformational changes propagated along these protofilaments are responsible for filament polymorphism. Note that in this figure the filament is not “opened out”, but is cylindrical, so that not all of the monomers are visible.

2. Surface Lattice The flagellin monomers are aranged in a pseudohexagonal lattice on the filament; there are about 5.5 monomers per turn of the basic (1-start) helix and a rise per subunit of about 0.5 nm. The term “start” denotes the number of identical monomer strands which together can account for all of the subunits in the filament. F o r example, a single 1-start helix or five parallel 5start helices can describe the entire filament (Fig. 6). The subunits can also be described as organized on 6-start or 11-start helices, with this last family nearly parallel to the filament axis (Fig. 6; O’Brien and Bennett, 1972; Kamiya et a f . , 1979). This packing arrangement is nearly identical for the filaments from species as diverse as Cuulobacrer crescenfus (Trachtenberg and DeRosier, 1988), Rhizobium fupini (Trachtenberg et af., 1986), E. coli (Kondoh and Yanagida, 1975) and S . typhimurium (O’Brien and Bennett, 1972). The molecular masses of the corresponding flagellins range from approximately 25 kDa to nearly 70 kDa (Lagenaur and Agabian, 1976; Schmitt et ul., 1974; Lawn, 1977; Ibrahim et a f . , 1985).

THE BACTERIAL FXAGELLUM AND FLAGELLAR MOTOR

125

The conservation of this lattice in R. lupini is especially interesting. Like other species of soil bacteria, R. lupini has what are termed “complex” flagella, so-called because they appear in electron micrographs to be helically corrugated (Krupski et al., 1985). Despite their difference in appearance from typical “plain” filaments, the monomers of complex filaments are arranged on a lattice very similar to that of plain filaments, but with pairwise interactions between subunits (Trachtenberg et al., 1986). Each pair of subunits may consist of two distinct flagellins (Pleier and Schmitt, 1989), in contrast to Bdellovibrio bacteriovorus and C. crescentus where the different flagellins comprising the filament are segregated (Thomashow and Rittenberg, 1985a; Driks el al., 1989). 3. Model for Filament Polymorphism

The model for filament structure invokes two distinct states for the subunit. The version of Asakura and his coworkers (Asakura, 1970; Kamiya et al., 1979) suggests two conformations for the subunit, whereas that of Calladine (1978,1983) suggests alternative intersubunit binding sites. Both versions of the model require some distortion of the subunits to accommodate their packing. Whether the specific residues involved in binding are the same for both forms of the monomer is not known, but this may be decided once the molecular structures of filaments consisting of each of the two subunit forms have been determined. All of the subunits in each 11-start “protofilament” are in one state (R form) or the other (L form). Co-operative transitions along one or more protofilaments result in changes in the overall filament shape (Asakura, 1970; Wakabayashi and Mitsui, 1972). In order to minimize strain in the filament, all of the L- and R-form protofilaments are adjacent (Calladine, 1978, 1983; Kamiya et al., 1979). There should therefore be 12 different polymorphs (containing 0,1,2, . . ., 11protofilaments in the R conformation), including two distinct straight polymorphs consisting solely of flagellin in one conformation or the other (Fig. 7). Most of these polymorphs have been observed, including both straight forms (Asakura and Iino, 1972; O’Brien and Bennett, 1972; Kamiya et al., 1979,1980,1982). Yamaguchi has isolated 43 S. typhimurium mutants with straight filaments, all of which can be classified as either L- or R-form straight filaments on the basis of their X-ray diffraction patterns (Kamiya et al., 1982), supporting the idea that there are only two forms of subunit in any flagellar filament. The necessary structural change between the R and L forms of the flagellin subunit is probably quite small (Calladine, 1978; Kamiya et al., 1979). Both forms can be copolymerized in various ratios into single filaments, forming different filament polymorphs (Kamiya et al., 1980,

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C. 1. JONES AND S.A. A I Z A W A

-6

r(pm-')

-4

A o ,

-2

s

I\_ -

0

P

2

%c7

Normal

Coiled

- Semi-coiled

4

6

8 0

10

12

+

0

0

14,

Straight lum (a)

( b)

(C)

FIG. 7. Model for filament polymorphism. (a) Plot of curvature (K) versus twist (T) for the 12 theoretical helices (open circles) and several observed flagellar filaments (closed symbols; see Calladine, 1978 and Kamiya et af., 1982). Curvature is a function of the number of longitudinal protofilaments in the alternative configuration, while twist is a function of the tilting of the protofilament relative to the filament axis. (b) The shapes of the 12 theoretical waveforms, all drawn with a contour length of 4 pm, aligned with the corresponding circle in (a). (c) Names given to some of the waveforms which have been observed. Several more waveforms have been seen since this diagram was made. Modified from Calladine (1978).


127

1982). Circular dichroism and UV spectral analyses have shown no significant change upon filament transformation (Hasegawa et al. , 1982). Also, Kanto and her coworkers have identified the mutations in the flagellins of 17 flagellar-shape mutants (one R-type straight, five L-type straight, nine curly and two coiled filaments) by amino-acid sequencing (S. Kanto, S.-I. Aizawa, and S. Yamaguchi, unpublished observation). Many mutations involved only a minor change in a single amino-acid residue. For example, one straight-mutant flagellin was the result of a single alanine-forglycine substitution. Similarly, a single valine-for-alanine substitution in BacilZus subtilis flagellin results in straight filaments (Martinez et al., 1968). Thus, presumably subtle changes in flagellin structure can drastically affect filament shape. 4. Filament Structure

Three-dimensional models for filament structure have been derived from electron micrographs of negatively stained R-form and frozen-hydrated Lform straight filaments (Shirakihara and Wakabayashi, 1979; Trachtenberg and DeRosier, 1987). Currently, the most detailed description of the structure of the flagellar filament is based on X-ray fibre-diffraction analysis of oriented sols of filaments reconstructed from the latter L-form flagellin (Fig. 8; Namba et al., 1989). Both models based on L-form filaments give a filament diameter of about 24 nm and show the presence of 11columns of density roughly parallel to the filament axis, with large outer knobs protruding outward from these columns. Based on the results of the latter study, the flagellar filament appears to have a hollow central channel with a diameter of about 6 nm (Fig. 8; Namba et al., 1989). Previous estimates of the channel diameter, based on negative staining or three-dimensional image reconstructions, were smaller than this, and were difficult to reconcile with the idea of flagellin molecules being transported through the filament (see Section 1V.A.). This is the major difference between the L-form models. Only that from the fibrediffraction study shows a large, well-defined central channel in the filament. Part of this difference probably results from the fact that the earlier studies used filaments detached from cells, whereas the fibre-diffraction study used filaments reconstructed from purified flagellin. The central channel is clearly visible in end-on views of sonicated filaments (Asakura et al. ,1968, plate V), and in transverse ultrathin sections of native (Kerridge et al., 1962; Swan, 1985) and reconstructed filaments (T. Akiba, C. J. Jones and K. Namba, unpublished data), lending confidence to our belief in the presence of such a large channel. The model derived from the R-form filaments differs in many details from

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C. J. JONES AND S.-I. AIZAWA

FIG. 8. Stereo image of the three-dimensional structure of the flagellar filament from Salmonella typhimurium determined by X-ray fibre diffraction. Above, a view along the filament axis. The cross-section is approximately 5.5 nm deep, and shows two complete turns of the basic helix. Below, a side view of the filament. The segment shown is 50 nm long. The photograph is reproduced by courtesy of K. Namba.


129

those of the L-form filament. How much of this is a result of true differences between the two filament structures, and how much is from differences in sample preparation, are not known. Studies are now underway to construct a model for the R-form filament using frozen-hydrated samples and oriented fibre sols (Trachtenberg and DeRosier, 1987; K. Namba, personal communication). Negatively stained filaments show a definite polarity, being pointed at the cell-proximal end and having a complementary notch or “fishtail” at the end distal to the cell (Asakura et al., 1968; O’Brien and Bennett, 1972). To account for this, the flagellin subunits are generally thought to be inclined at an angle of about 45” to the filament axis (but see Trachtenberg and DeRosier, 1987). Because no model of filament structure has yet been determined to sufficient resolution to identify clearly the flagellin subunit within the filament, the question of subunit orientation is still unresolved. It should be noted in this regard that the ends of unstained, freeze-dried filaments have a similar pointed or notched appearance, whereas those of frozen-hydrated filaments are blunt, suggesting that this feature may be an artefact of drying (Trachtenberg and DeRosier, 1987; see also O’Brien and Bennett, 1972). Each subunit is elongated and appears to consist of three (or four) domains, with intersubunit contacts mediated by the flagellin domain(s) at inner radii (Shirakihara and Wakabayashi, 1979; Trachtenberg and DeRosier, 1987; Namba etal., 1989). The fibre-diffraction pattern indicates that each monomer contains a bundle of a-helices near the centre of the filament, aligned parallel to the filament axis (Namba et al., 1989). 5. Flagellin Structure

Attempts to make three-dimensional crystals of flagellin have failed, because flagellin polymerizes into filaments under these conditions (our laboratory, unpublished observations), so it has not yet proved possible to apply conventional X-ray diffraction analysis to this protein. Studies of flagellin structure have therefore used a variety of other techniques. Proteolysis of flagellin from S. typhirnurzurn (with a molecular mass of 52 kDa) results in production of a moderately stable fragment of about 40 kDa (F40); further digestion results in a more stable fragment of about 27 kDa (F27). These fragments each result from removal of the terminal regions of the preceding molecule; €27 is internal to F40, which is itself internal to flagellin (Vonderviszt et al., 1989). Parallel results have been obtained with flagellin from E. coli (Kostyukova et al., 1988). Scanning microcalorimetric and circular dichroism studies of filament, monomeric flagellin and these fragments indicate that the flagellin monomer

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C J JONES AND S - l

AlZAWA

is composed of three compact domains, and that one more domain is formed upon polymerization of the monomer into filament (Uratani et al., 1972; Fedorov et al., 1984; Kostyukova et al., 1988; Vonderviszt et al., 1989). This fourth domain is composed of the terminal regions of flagellin (Kostyukova et al., 1988; Vonderviszt et al., 1989; Aizawa et al., 1990). Secondarystructure predictions and circular dichroism analysis suggest that F27 consists of two P-sheeted domains with very little a-helix, the third, discontinuous domain consists of approximately equal amounts of sheet and helix, and that the terminal regions of the molecule are primarily ahelical (Fedorov et d . ,1988; Vonderviszt et d . ,1990). This is consistent with the results of circular-dichroism studies (Klein ef al., 1969; Uratani et al., 1972). The similarity of filament subunit lattices (see above), the fact that flagellins from different strains and species can be copolymerized into single filaments (see Section IV.A), and the similarity of polymorphs formed by different flagellins (Kamiya et al., 1982) all suggest that the structural basis for polymerization of different flagellins may be identical. Mutations which affect the ability of flagellin to polymerize into filament are localized to the terminal regions of the molecule (Horiguchi et al., 1975; Iino, 1977; Yamaguchi et al., 1984a, Homma et al., 1987a), suggesting that only the termini are important for polymerization and that the central region may be responsible for the antigenic determinants on the filament surface. These predictions have been verified in a number of ways. (i) DNAsequence analysis of various flagellin genes has revealed that, while there is a significant degree of sequence conservation in the N - and C-terminal regions of the encoded proteins, the sequences of the central regions vary considerably (Joys, 1985; Wei and Joys, 1985; Kuwajima et al., 1986; Martin and Savage, 1988; Harshey et al., 1989; Logan et al., 1989; Smith and Selander, 1990). This variation accounts for almost all of the differences in molecular mass between different flagellins. (ii) Kuwajima (1988b) has shown that flagellin consisting of only the N-terminal40% and C-terminal 25% of the molecule can still form a functional filament. (iii) Removing the termini of flagellin with proteases destroys its ability to polymerize (Fedorov etal., 1988;Vonderviszt etal., 1989). These termini cannot be removed from monomers incorporated into filament, which is very stable to proteolytic digestion (Kostyukova et al., 1988; Vonderviszt et al., 1989). (iv) After removing its termini, the remaining portion of the flagellin molecule retains all of the antigenic determinants of the polymer (Parish et a f . ,1969; Fedorov et al., 1988). (v) Deletions in the central region of flagellin alter the antigenicity of the molecule (Kuwajima, 1988a). (vi) The flagellin of C. crescentus is only about half the size of that from S. typhimurium, essentially lacking a major portion of the central region from the latter, and the


131

filament of C. crescentus lacks the outer domains seen in the filament from S. typhimurium (Trachtenberg and DeRosier, 1988). The picture that emerges from all of this work is that at least part of the central portion of the flagellin molecule is exposed on the surface of the filament and both of the termini are presumably located inside. This arrangement is consistent with the fact that, in the filament, intersubunit contacts (and therefore, presumably, the regions of flagellin essential for polymerization) are found at inner radii (Namba et af., 1989). The ability of flagellin to function despite major changes in its central region has recently inspired some promising experiments. At least two laboratories have successfully inserted foreign peptides into the central region of flagellin without seriously impairing polymerization; the mutant filaments display the epitopes of the inserted peptides (Kuwajima et a f . , 1989; Newton et al., 1989; Wu et a f . , 1989). Since epitopes exposed on polymerized flagellin evoke strong immune responses (Newton et al., 1989; Wu et al., 1989), this system is being examined as a possible source for vaccines. In some Salmonella spp., approximately half of the lysine residues in flagellin are converted to N-methyllysine by FliB (Parish and Ada, 1969; S.-I. Aizawa, unpublished data). No proteins other than flagellin contain Nmethyllysine (Stocker et a f . ,1961). Consistent with this, fiiB transcription is controlled by the flagellar master operon (Suzuki et af., 1988). BecausefiiB is not necessary for flagellar assembly (Stocker et a f . , 1961; Konno et a f . , 1976), its maintenance indicates that it must confer some advantage on the bacterium. Given that the outer domain of flagellin is composed of the central region of the molecule, which contains almost all of the Nmethylated lysine residues (Parish and Ada, 1969; S.-I. Aizawa, unpublished data), it is possible that such modification may make the flagellum more resistant to tryptic enzymes in the gut (Parish and Ada, 1969) or may alter flagellar antigenicity. Neither of these possibilities has been investigated. B . HOOK AND HOOK-ASSOCIATED PROTEINS

1. Hook

The hook appears in electron micrographs as a curved structure about 50 nm long with a diameter of about 20 nm (Fig. 3(c); DePamphilis and Adler, 1971b). It is a short segment of a right-handed helix (Kato et a f . , 1984) composed of roughly 120 copies of a single protein (Silverman and Simon, 1972; Hilmen et a f . , 1974; Kagawa et a f . , 1976) arranged on a pseudohexagonal lattice with lattice parameters almost identical to those of the

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C. J. JONES AND S.A. AIZAWA

filament (Wagenknecht et af., 1982; see Section 1II.A). Also, like the filament, the subunit monomers of the hook are elongated and are tilted about 45" relative to the hook axis, being more cell-distal at outer radii (Wagenknecht et al., 1982). Three-dimensional image reconstructions of the hook show that it has deep grooves running in the 6-start direction (Wagenknecht et al., 1982). These grooves may allow the hook to bend without steric interference between the subunit monomers at outer radii; such bending is crucial to the presumed function of the hook as a universal joint, allowing the filaments to form the flagellar bundle (Berg and Anderson, 1973). The flexibility of the hook is supported by the convoluted appearance in electron micrographs of extended hook polymers (see, for example, PattersonDelafield et al., 1973; Kutsukake et al., 1979; Jones et al., 1987). Such polymers can result in vivo from mutation of fliK, whose product regulates hook length (see Section 1V.B); these extended hook structures are called polyhooks. Like the filament, polyhooks also have different polymorphic forms (Kutsukake et al., 1979) and undergo polymorphic transitions (Kagawa et al., 1979; Kato et al., 1984). However, whereas the protofilaments along the 11-start helix co-operate in filament transitions, those along the 16-start helix appear to do so in the hook (Kato et al., 1984). If true, this difference may contribute to the need for two junction proteins (HAPl and HAP3) between the hook and filament. 2. Hook-Associated Proteins

The HAP proteins were first identified as minor components which copurified with the hook (Homma et al., 1984a). They have since been shown to be necessary for filament assembly. HAPl and HAP3 form the junction between the hook and the filament, with a zone composed of HAPl proximal to one composed of HAP3 (Fig. 9; Homma and Iino, 1985a;Ikeda et al., 1987). HAP2 caps the distal end of the filament (Fig. 9; Ikeda et al., 1987). The presence of HAP2 is necessary to enable flagellin monomers synthesized within the cell to assemble onto the filament, but it co-incidentally prevents addition of exogenously added monomers (Homma et al., 1986; see Section 1V.A). Estimates of HAP stoichiometries based on Coomassie-blue binding and normalized to the number of hook monomers (using estimates of hook length and its known lattice structure) indicate that the flagellum contains 10-15 copies of HAPl, 10-30 copies of HAP3 and 6-12 copies of HAP2 (Ikeda et al., 1987, but see Jones et al., 1990). By analysing radiolabelled hook-basal body complexes, we estimate that there are about 13 molecules of HAPl in the flagellum (Jones et al., 1990). If HAPl and HAP3 are


133

HAP2 ffliD} 10 nm

HAP1fffgK)

\ Filament ff7iC)

/

/

I

,Distal

rod fffgGI Outer membrane Peptidoglycan layer

Rod fflg8, ffgC, ffgF1 Cytoplasmic membrane

FIG. 9. Schematic diagram of the flagellum of Salmonella typhimurium. The various substructures of the flagellum are indicated with the gene which encodes the corresponding structural protein, where known, in parentheses. The Mot and switch proteins are not shown; although they must be part of the motor (see Section V.A), their locations relative to the flagellum are not known. Dimensions are those measured from electron micrographs of negatively stained samples. The approximate positions of the layers of the cell envelope are shown. For clarity, both sections of the rod are shaded. Only the proximal and distal ends of the filament, which is typically 5-10 pm long, are shown. HAP denotes hook-associated protein.

present in equimolar amounts (Homma et al., 1984a), this implies that both could form two turns of the basic helix in the flagellum if they assemble on the same lattice as the flanking hook and filament. This assumption seems particularly reasonable in view of the homology between the termini of these proteins and the termini of flagellin (Homma et al., 1990a), which are necessary for filament polymerization (see above). Four such turns would form a segment about 10 nm long, consistent with the length of the junction sometimes observed in electron micrographs (Ikeda et al., 1987). C. BASALBODY

Purified flagella consist of the filament, hook and basal body (DePamphilis and Adler, 1971a; Dimmitt and Simon, 1971). After depolymerization of

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C. J. JONES AND S.4. AIZAWA

the filament, usually by incubation of the flagella at pH 2.5, a structure called the hook-basal body (HBB) complex remains (DePamphilis and Adler, 1971b; Aizawa et af.,1985). Hook-basal body preparations are at the heart of most investigations into basal-body structure. The flagellar basal body of E. coli and S. typhimurium appears as a complex of four rings connected by a central rod which is itself connected to the curved hook (Fig. 9). Although the basal body cannot be the entire flagellar motor, as it lacks both the Mot proteins and the switch complex necessary for motor function (Macnab and Aizawa, 1984; Aizawa ef al., 1985; Clegg and Koshland, 1985), its location in the cell wall and connection to the hook and filament argue that it is a key part of the motor. The basal body consists of at least seven proteins, for which the corresponding genes and substructures have been identified (see Fig. 9 and Table 1). Additional flagellar proteins copurifying with the HBB complex have been identified (Jones and Macnab, 1990), but their relationship to the basal body has not been determined.

I . Rings The basal-body rings interact with the various layers of the cell envelope (DePamphilis and Adler, 1971~).The M ring is integral to the cytoplasmic membrane and the L ring to the outer (lipopolysaccharide)membrane while the P ring is thought to interact with the peptidoglycan layer, although this remains to be demonstrated. The S (supramembrane) ring lies immediately distal to the M ring (Fig. 9). The subunit proteins and their corresponding genes have been identified for the M, P and L rings (Komedaet al., 1978;Aizawa et af.,1985;Homma et af., 1987c,d; Jones ef af., 1987). It is still not known whether fhe S ring is separate from or a projection of the M ring, but extensive searches have failed to suggest a candidate subunit protein or encoding gene for the S ring. The M and S rings appear strongly connected in three-dimensional image reconstructions (Stallmeyer et al., 1989), also supporting the latter possibility. The P and L rings together form the outer cylinder, which is thought to function simply as a bushing, allowing free rotation of the rod (Berg, 1974). Indirect support for this putative function comes from several observations. (i) Overproduction of the hook protein can override mutations in eitherJlgH orJlgZ(the genes for the L- and P-ring proteins), resultingin the formation of functional flagella lacking part or all of the outer cylinder (Jones et al., 1987; Ohnishi ef al., 1987). (ii) The flagella of Gram-positive bacteria lack any structure corresponding to the outer cylinder (DePamphilis and Adler, 1971b; Dimmitt and Simon, 1971). (iii) The axial position and orientation of the outer cylinder in isolated HBB complexes is slightly variable (Stallmeyer


135

et al., 1989). How the position of the outer cylinder along the rod is maintained is not known. The distal flange (corresponding to the L ring) and wall of the outer cylinder are contributed by the FlgH protein, with the proximal disk built of FlgI molecules (Jones et al., 1987). These rings are about 26 nm in diameter and about 10 nm apart, with about 17 nm separating opposite sides of the cylinder wall (Stallmeyer et al., 1989). Given the relatively harsh conditions used in purifying the basal body (pH extremes of 2.5 and 11, detergent, and caesium chloride banding; Aizawa et al., 1985),the terminal M ring must be firmly attached to the rod, indicating that it probably turns with the rod during flagellar rotation. This, the fact that the M ring is integral to the cytoplasmic membrane (DePamphilis and Adler, 1971c) and it is the flow of protons across this membrane that drives the motor (see Section V.A), and the proximity of the M ring to presumed Mot complexes in the membrane (see Section 1II.D) all suggest that the M ring may be the rotor of the flagellar motor. The M r i n d s ring complex has a complicated structure in threedimensional image reconstruction, resembling a wing-nut in cross-section, with diameters of about 29 nm at the M-ring “ears” and about 25 nm at the S ring (Stallmeyer et al., 1989). Each of the rings contains approximately 26 subunits (Jones et al., 1990); it is not clear whether this is significant in terms of structure. This identity in the number of subunits is not surprising in the case of the P and L rings, since they interact directly. The M ring probably has a similar number of subunits simply because it is a structure similar in size to the P and L rings and its subunits are of a comparable size. The genes for the subunit proteins of the M, P and L rings have been sequenced (Jones et al., 1989), but there is as yet insufficient structural data to enable us to relate their primary and higher-order structures. The M-ring protein appears to have, at most, three membrane-spanning a-helices but, in view of the apparent thickness of this ring in the plane of the inner membrane, there are probably other membrane-spanning segments. The Lring protein is predicted to be relatively rich in p-structure, a common motif in other outer-membrane proteins (Vogel and Jahnig, 1986; Sass et al., 1989). 2. Rod

The rod passes through all four rings, with a convex terminus facing the cytoplasm (Fig. 9; Stallmeyer et al., 1989). It can be split into two portions at a point close to the plane of the P ring, either by incubating the basal body at low pH values (Sosinsky et al., 1988) or by growing cells of a mutant in the

136


fliF (M-ring) gene in high-viscosity media (Okino et al., 1989). In the latter study, the proximal and distal portions of the rod were determined to have diameters of about 13 and 18 nm, respectively. Stallmeyer and her coworkers (1989), on the other hand, describe the distal portion of the rod as being slightly narrower than the proximal portion, and cite an estimate of 13 nm for the diameter of the former. The reason for this discrepancy is unclear, but the boundary between the two sections of the rod may reflect a structural difference which serves to restrict movement of the outer cylinder along the rod. Four subunit proteins of the rod, and their encoding genes, are known (Komeda et al., 1978; Aizawa et al., 1985; Homma et al., 1990b). The distal portion of the rod consists of approximately 26 copies of FlgG, and the proximal portion contains five or six copies of each of FlgB, FlgC and FlgF (Okino et al., 1989; Jones et al., 1990). Assuming that the rod proteins assemble on a lattice similar to that of the hook and filament (see Section III.A), these stoichiometries cannot account for the observed length of the rod, suggesting the existence of additional rod components. The assumption of a common assembly lattice is supported by the presence of sequence homologies in the N - and C-terminal regions of the rod, hook, HAP and filament proteins, as deduced from gene sequences (Homma et al., 1990a,b) . Although rotation of the rod has never been directly demonstrated, it must serve to transmit the torque generated at the cytoplasmic membrane to the hook, and thence to the filament.

3. Additional Basal- Body Components

There are several additional proteins present in preparations of HBB complexes from S. typhimurium that are synthesized under the control of the flagellar master operon (see Section II), and are therefore presumably true component proteins of the basal body (Jones and Macnab, 1990). The functions of these proteins are not known, but one possible role is as part of the machinery to control export of the axial components of the flagellum (see Section 1V.A). Sequencing of these proteins and identification of their encoding genes may help to determine their roles in the basal body. The basal bodies of several species with polar flagella have an additional structure, consisting of one or a pair of very large discs, 80-170 nm in diameter, which are sensitive to proteases and are apparently associated with the outer membrane (Coulton and Murray, 1977, 1978; Curry ef al., 1984; Ferris et al., 1984; Kupper et al., 1989). The function of these structures is not known, but it may have to do with the location of these


137

flagella at the cell pole; such discs have never been reported associated with flagella from peritrichously flagellated species. D. MOT AND SWITCH COMPLEXES

The only portion of the flagellar motor which has been characterized biochemically is the basal body. The Mot and switch proteins must form part of the motor, since they are all necessary for energy transduction, but their relative locations and structures have not been demonstrated. Evidence for the existence of Mot and switch complexes is convincing, but indirect. All of the genes encoding components of these putative complexes have been sequenced, and the sequences of the proteins deduced (Dean et a f . , 1984; Kuo and Koshland, 1986; Stader et a f ., 1986;Kihara et al., 1989;Malakooti et al., 1989). 1. Motor Elements

Like any rotary motor, the flagellar motor must have both rotor and stator elements which mutually rotate against each other. The stator must in turn be attached to a rigid structure, presumably the peptidoglycan layer, in order to be able to apply the force the motor generates and propel the cell. If there were no such anchoring, the motor would still function, passing protons and generating torque, but the viscous resistance of the medium acting on the filament would result in rotation of the stator in the cell envelope. Indeed, some Mot- mutations (perhaps those in motB, see below) may turn out to be the result of just such a disruption of anchoring. It is interesting to note in this regard that Thermoplasma acidophilum, an organism probably related to the archaebacteria (Razin, 1985), appears to lack a cell wall but is nonetheless flagellated and motile (Black et al., 1979). Because at least some of the cells were non-spherical, they must possess some rigid surface structure upon which the flagella could be anchored. Several bacterial species have been shown to possess rings of intramembrane particles, or studs, surrounding a space large enough to accommodate the basal-body M ring (Chalcroft et al., 1973; Coulton and Murray, 1978; Khan et al., 1988). Khan and his coworkers (1988) have shown that these particles are absent from E. coli strains lacking either MotA or MotB, suggesting that these two proteins comprise at least part of these particles. The fact that the Mot proteins can reversibly assemble into the motor after its construction (Block and Berg, 1984; Blair and Berg, 1988) supports a location for these proteins circumferential to the M ring. The MotA and MotB proteins presumably form a major part of the stator and the M ring, the rotor (see Section 111.C); whether the switch proteins form part of the rotor or the stator is not known.

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C. I. JONES AND S.-I. AIZAWA

2. Mot Complex

MotA is an integral cytoplasmic membrane protein (Ridgway et al., 1977; Wilson and Macnab, 1988) which conducts protons across the membrane and is therefore probably the proton channel of the motor (Blair and Berg, 1990). Sequence analysis of the motA gene suggests that the protein may span the membrane with four a-helices; two highly charged regions may also cross the membrane and function in proton transport (Dean et al., 1984). The data of Blair and Berg (1990) indicate that MotA functions as a monomer in a larger assembly, presumably including MotB and possibly one or more of the switch proteins. Unlike MotB (see below), large amounts of MotA can insert into the membrane, although there are a limited number of functional sites it can occupy (Wilson and Macnab, 1988; Blair and Berg, 1990). MotA is normally present at levels of about 600 molecules per cell (Wilson and Macnab, 1988); as there are only about five flagella on the average E. coli cell, it is likely that many copies of MotA are normally adrift in the membrane. MotA when overproduced to levels of about 30,000 copies per cell is still localized to the cytoplasmic membrane (Wilson and Macnab, 1988). Excess MotA not associated with the motors appears to be relatively inefficient at transporting protons, presumably because it is not in optimal interaction with other motor components (Blair and Berg, 1990), which may explain why such high levels of overproduction failed to impair cell motility or growth rate significantly (Wilson and Macnab, 1988). The MotB protein is currently the best candidate for the species anchoring the motor to the cell wall. It is integral to the cytoplasmic membrane (Ridgway et al., 1977). Using proteolysis and alkaline phosphatase fusion experiments, Chun and Parkinson (1988) have shown that MotB crosses the cytoplasmic membrane once and has a large C-terminal periplasmic domain which they suggest could interact with the peptidoglycan layer, linking the force generators of the motor (see below) to the cell wall. Incorporation of MotB into the membrane is site limited (Stader et al., 1986), and MotB appears to form part of the intramembrane particles which surround the M ring (Khan et al., 1988), consistent with this proposed role. On the other hand, motors can lose force generators containing MotB (Block and Berg, 1984; Blair and Berg, 1990), indicating that, if MotB does link the motor to the cell wall, this anchoring may be reversible. The number of MotB subunits normally present in the cell envelope is not known. Results from bacteriophage h-programming experiments suggest that the protein is synthesized at a level only about one-twentieth that of MotA (see Dean et al., 1984), but this may not be representative of relative amounts of these proteins synthesized under normal conditions.

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In the resurrection experiments of Berg and his coworkers (Block and Berg, 1984; Blair and Berg, 1990), flagellar torque increased in equally spaced steps, which they ascribed to the stepwise addition of force generators, containing the plasmid-encoded MotA and MotB, to the paralysed motors. Since decreases in rotation (of equal size) were also seen, these force generators must not be irreversibly associated with the motor. Blair and Berg (1990) conclude that there are probably eight such generators associated with a fully functional motor, and that motors from exponentially growing cells may not possess the full complement of generators. Because freeze-fracture preparations of E. cofi (Khan et af., 1988) show intramembrane rings of ten to 12 particles surrounding depressions large enough to accommodate the M ring, they suggest that the full complement of such particles could be 16, so that each force generator would consist of two such particles. The rings appear fairly uniform, however, so how four or five more particles might squeeze in without greatly altering the ring diameter (and thus the distance between the M ring and the generators) is not obvious. Another possibility is that, at any one time, a maximum of only eight of the particles are properly aligned so as to be able to function. Freeze-fracture analysis of cells with “fully-loaded” motors might help to resolve the question of the relationship between the morphologically described intramembrane particles and the functionally described force generators.

3. Switch Complex FliG, FliM and FliN are all peripheral membrane proteins (Ridgway et af., 1977; Ravid and Eisenbach, 1984a; Homma et al., 1988; Kihara et al., 1989; Malakooti et a f . , 1989). Because mutants in any of the corresponding genes may be non-flagellate (Fla-), non-motile (Mot-) or non-chemotactic (Che-), depending on the allele, these proteins are required for flagellar assembly, conversion of the transmembrane proton-motive force into rotational work, and modulation of switching of the direction of rotation of the motor (see, for example, Yamaguchi et af., 1972, 1986a; Tsui-Collins and Stocker, 1976; Warrick et af., 1977; Dean et al., 1983; Parkinson et al., 1983). These different types of mutation are localized to different regions of these genes (Yamaguchi et af., 1986a). Nothing is known of the stoichiometries of FliG, FliM and FliN in the cell. The amount of FliM synthesized is greater than that of FliN when both are expressed from a plasmid (Malakooti et af., 1989) but, as is true for MotA and MotB (see above), this may not accurately reflect the normal levels of synthesis in vivo. By isolating allele-specific pseudoreverting mutations in these genes

140


which complemented lesions in one of the others, Yamaguchi and his coworkers (1986b) provided evidence that these proteins are in physical interaction in the cell, forming a “switch complex’’ (Parkinson et al., 1983; Macnab and Aizawa, 1984). Certain switch-complex mutations could also be partially compensated for by mutations in the MotB, CheA, CheY and CheZ proteins (Parkinson and Parker, 1979; Parkinson et al., 1983; Yamaguchi et al., 1986b), consistent with the fact that the switch complex functions in both the motility and chemotaxis systems, and with the idea that it is the interface between them. Several lines of evidence suggest that the switch complex is located at the cytoplasmic face of the basal body. (i) The switch proteins are required for the earliest stages of flagellar assembly, which begins at the cytoplasmic membrane (Suzuki et al., 1978; Suzuki and Komeda, 1981; Jones and Macnab, 1990). (ii) The switch complex physically interacts with CheY and CheZ, both cytoplasmic proteins (Parkinson and Parker, 1979; Parkinson et al., 1983; Clegg and Koshland, 1984; Ravid et al., 1986; Yamaguchi et al., 1986b). (iii) FliG and FliM are associated with the cell envelope (Ravid and Eisenbach, 1984a; Homma et al., 1988; Malakooti et al., 1989). (iv) Because they have Mot- alleles, the switch proteins are involved in the process of energy transduction, which occurs across the cytoplasmic membrane (see Section V. A). Since all five of these proteins are necessary for energy transduction, they may all interact, presumably at the circumference of the M ring where the Mot proteins are thought to be located. It may turn out that the switch complex is in fact a multiple of complexes, and they may form an interface between the force generators containing MotA and MotB and the basalbody M ring. IV. Assembly A . FILAMENT

The bacterial flagellar filament can self-assemble from monomeric flagellin in witro in a process which has been compared to crystallization (Ada et al., 1963; Abram and Koffler, 1964; Asakura et al., 1964; Asakura, 1970). Although the resulting filament is identical with one constructed in wiwo (Asakura etal., 1966; Wakabayashi etal., 1969), there are several important differences between these two processes, the most important being that addition of flagellin monomers to the filament in wiwo requires the presence of an accessory protein, HAP2 (Homma et al., 1984a, 1986).


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I . Assembly in vitro Filaments can be depolymerized into monomers by any of a number of treatments. The most commonly used are incubation at low pH values (around 2.5) or high temperature (60°C for ten minutes; e.g., see Wakabayashi et al., 1969; Kuroda, 1972; Kondoh and Hotani, 1974). Either treatment results in a stable, super-saturated monomer solution. Polymerization can be induced under physiological conditions by adding short fragments (seeds) of filament or by adding a precipitant such as ammonium sulphate or polyethylene glycol (e.g. see Ada et al., 1963; Abram and Koffler, 1964; Asakura et al., 1964, 1966; Wakabayashi et al., 1969; Novikov et al., 1982). Filament elongation in vitro proceeds at a constant average rate (Hotani and Asakura, 1974) as long as there is a supply of monomer available and occurs only at one end of the growing filament, the end that corresponds to the distal end of the filament on the intact cell (Asakura et al., 1968). Heatinduced melting of the filament also occurs preferentially at this end (Hotani and Kagawa, 1974). The maximum rate of elongation observed in vitro is only about half that of short (i.e. rapidly growing, see below) filaments in vivo (Iino, 1974; Kondoh and Hotani, 1974), but varies with the medium and flagellin used (Asakura et al., 1964; Kuroda, 1972; Kondoh and Hotani, 1974). A noteworthy advantage of polymerization in vitro is that it allows us to copolymerize different flagellins (Asakura et al., 1966; Lawn, 1977). Copolymerization of flagellins from different species has indicated that the molecular interactions underlying the process of polymerization have been conserved through evolution (Kuroda, 1972; Kondoh and Hotani, 1974). Copolymerization of L- and R-form flagellins in different ratios results in formation of many of the helical polymorphs (Kamiya et al., 1980), lending support to the model of filament structure proposed by Calladine (1978) and Kamiya et al. (1979) (see Section 1II.A). 2. Assembly in vivo Filament assembly in vivo requires the presence of HAP2 protein capping the distal end of the filament (Homma et al., 1986), where monomers are added (Iino, 1969; Emerson et al., 1970). Details of the subsequent interaction are not known but, since flagellin is capable of self-assembling (albeit at relatively high monomer concentrations) in vitro, the HAP2 structure probably acts simply to prevent arriving monomers from diffusing away until they assume the conformation necessary for addition to the filament (see below). Consistent with this, flagellin can be polymerized in vitro onto the

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filaments of living cells provided that HAP2 is lacking, either as a result of mutation (Kagawa efa f . ,1981,1983) or shearing of the filaments (Iino et al., 1972). In the latter case, HAP2 synthesis must be prevented, either by starving the cells (Iino ef al., 1972) or by adding chloramphenicol (Iino, 1974), before exogenous flagellin can be successfully polymerized onto the portions of the flagella remaining. While elongation in vitro occurs at a constant average rate regardless of the length of the filament (Hotani and Asakura, 1974), the elongation rate in vivo decreases exponentially with increasing filament length (Iino, 1969, 1974); no plausible explanation of this phenomenon has been presented. Filaments broken mechanically grow at the same rate as new filaments which have reached the same length, ruling out aging of the flagellum as a factor in this decrease in growth rate (Iino, 1969). A number of studies indicate that, for E. coli and S. typhirnuriurn, there is not a significant pool of flagellin monomers inside the cell from which the filament can assemble (Kerridge, 1963; Dimmitt et al., 1968; Silverman and Simon, 1974a,b; Iino ef al., 1975). This may not be true for other species (Martinez and Gordee, 1966). a. Ffagelfinfransporf.Flagellin monomers reach their assembly site at the distal end of the filament by travelling through the hollow core of the filament. Although this mode of transport seems somewhat implausible, several lines of evidence support it. There is essentially no flagellin in the culture medium of flagellated cells (Ikeda ef al., 1983; Homma ef al., 1984b), arguing against simple export of the monomer and its subsequent diffusion to the end of the filament. Consistent with this, flagellin lacks a signal sequence required for the conventional export pathway (Joys, 1985; Wei and Joys, 1985; Kuwajima ef al., 1986). Co-cultivation of two strains with different filament shapes does not result either in heteromorphous filaments or in cells bearing both kinds of filaments, which would be expected if flagellin were exported into the culture medium and assembled (Iino, 1969). Also, the decrease in filament elongation rate with increasing filament length already referred to (Iino, 1969) is difficult to reconcile with diffusion of monomer from the medium to the tip of the filament. Little is known about this flagellar-specific export pathway. There must be some sort of gating mechanism, presumably at the cytoplasmic face of the basal body, to distinguish flagellin and component proteins of the other axial structures of the flagellum (see Sections 1V.B and 1V.C) from other cytoplasmic proteins, and possibly to facilitate their export. What serves as a signal for this export pathway is unknown.

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Yamaguchi and his coworkers mapped the mutations in a number of export-deficient flagellins and discovered that they were in the highly conserved N- and C-terminal regions responsible for flagellin polymerization (Homma et al., 1987a; see below). Some deletions in the C-terminal portion of flagellin from B. subtilis can also affect export (LaVallie and Stahl, 1989). The N-terminal third of flagellin is itself competent for export, suggesting that the C-terminal mutations block flagellin export by their effects on the conformation of the molecule (Kuwajima et al., 1989). Comparison of the primary sequences of proteins comprising the axial flagellar structures has revealed no obvious similarities which might serve as a signal for this export pathway (Homma et al., 1990a,b). However, there must be some control mechanism because these proteins are exported and assembled in an ordered fashion; perhaps a combination of conformational and sequence aspects serve as export signals in these proteins. Some other species appear to use different pathways for flagellin transport. A flagellin gene from Spirochaeta auranfiahas a signal sequence, suggesting that the flagellum, which is located in the periplasmic space of the cell (Canale-Parola, 1978; Holt, 1978), is assembled from monomers using the conventional export pathway (Brahamsha and Greenberg, 1989). The archaebacterium Halobacterium halobium has five flagellin genes; the predicted products of these genes share extensive homologies with each other but are unrelated to eubacterial flagellins (Gerl and Sumper, 1988). It is interesting to note that the terminal regions of these flagellins have the greatest similarity, while the central region is the most divergent, as found in flagellins from eubacteria, suggesting a similar assembly mechanism. The Ntermini of flagellins from two other, very different, archaebacteria are highly homologous to an internal segment of the conserved N-terminal region of flagellins from H. halobium, suggesting that archaebacterial flage!lins may have an approximately 12-residue leader peptide (Kalmokoff et al., 1990), pointing to a flagellin-transport pathway different from that used by E. coli and S. typhimurium. b. Conformational changes in flagellin structure upon polymerization. The terminal regions of flagellin are essential for filament assembly. This has been confirmed using a variety of approaches. (i) Flagellin mutations affecting polymerization o r filament shape are preferentially found in the Nand C-termini of the molecule (Yamaguchi et al., 1984a; Homma et al., 1987a; S. Kanto, S.4. Aizawa and S. Yamaguchi, unpublished observation). (ii) Unlike that of the central region of the molecule, the sequences of the terminal regions are highly conserved in different flagellins (Joys, 1985; Wei and Joys, 1985; Kuwajima et al., 1986; Martin and Savage, 1988; Harshey et

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al., 1989; Logan et al., 1989). (iii) Flagellins with various deletions in the central region are capable of polymerizing (Kuwajima, 1988b), while proteolytic fragments lacking the termini are not (Fedorov et al., 1988; Vonderviszt et al., 1989). Secondary-structure prediction strongly suggests that the terminal regions of flagellin are a-helical (Fedorov et al., 1988; Vonderviszt et al., 1990), but scanning microcalorimetric and circular dichroism studies on flagellin and its tryptic fragments (Kostyukova ef al., 1988; Vonderviszt et al., 1989) have indicated that, although the terminal regions in the monomer may have some marginally stable a-helical character (Vonderviszt et al., 1989), they do not have any stable tertiary structure (Kostyukova et al., 1988; Vonderviszt et al., 1989). Nuclear magnetic resonance analysis of filament, flagellin and its tryptic fragments (see Section 1II.A) has confirmed that the termini of flagellin are highly mobile in solution but are stable in the polymer (Aizawa et al., 1990). It is rather surprising that the terminal regions, although indispensable for assembly, are disordered in solution; we suspect that this serves to prevent flagellin monomers assembling in the cytoplasm. The terminal regions together are believed to form the innermost flagellin domain in the filament (Namba et al., 1989). The X-ray fibre-diffraction analysis already described shows that at least part of this domain of flagellin does consist of a-helices in the polymer (Namba et al., 1989). Circular dichroism studies have shown that the a-helical content of flagellin doubles upon polymerization (Klein et al., 1969; Uratani et al., 1972). Taken together, these data suggest that flagellin monomers are transported, by an unknown mechanism, to the distal end of the filament where their Nand C-termini become organized into a-helices upon polymerization into filament. The HAP2 protein serves to maintain an extremely high local concentration of flagellin at the filament tip. B . HOOK AND HOOK-ASSOCIATED PROTEINS

Studies of the assembly of the hook and HAP structures have generally been hampered by the limited size of these polymers. On the basis of their locations and the sequence similarities they share with flagellin (Homma ef al., 1990a), the hook protein and the HAPS are presumed to be exported by the same flagellar-specific pathway used by flagellin (see Section 1V.A). I. Hook

Unlike the filament, the length of the hook is closely regulated, being about 50 nm (DePamphilis and Adler, 1971b; Ikeda et al., 1987; Jones et al., 1990).


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Length regulation of the hook is controlled by FliK. Mutants in fliK assemble hook polymers of various lengths, termed polyhooks, onto the basal body (Silverman and Simon, 1972; Patterson-Delafield et af., 1973). Reconstitution of polyhooks in vitro has shown that, like flagellin (see Section IV.A), hook monomers polymerize at the distal end of the structure (Kato et al., 1982). This is probably also true of assembly in vivo. Mutants inPiK are non-motile because they assemble filaments onto their polyhooks only at very low levels, presumably because polyhooks lack HAPs (Homma et a f . , 1984a). Motile revertants of one fliK mutant assembled filaments onto their polyhooks. Detailed mapping of one such revertant revealed that the second mutation was also infliK, indicating that the length-regulation and filament-addition functions of FliK can be separated (Suzuki and Iino, 1981). The mechanism of FliK function is unknown. The hook protein is exported via the flagellum-specific pathway (Homma et af., 1990a; see Section 1V.A) and is not generally excreted from flagellated cells (Homma and Iino, 1985b), suggesting that its export is somehow restricted once hook assembly is completed. On the other hand, it does appear to be exported from fliD (HAP2) mutants, but not from flgK (HAPl),flgL (HAP3) orfliC (flagellin) mutants (Homma and Iino, 1985b). The significance of this is unclear.

2. Hook-Associated Proteins Reconstitution experiments in vitro with HAP proteins have shown that they assemble onto the hook in discrete zones in the order HAP1-HAP3HAP2(Hommaetaf., 1986; Ikedaetal., 1989), consistent with theirorderin the flagellum (Fig. 10; Homma and Iino, 1985a; Ikeda et al., 1987). Whether the monomers assemble in a proximal-to-distal fashion within each zone, as do the hook and filament, is not known. The inability to form extended H A P l or HAP3 polymers in vitro, even with very high monomer concentrations (Ikeda et al., 1989), suggests that H A P l and HAP3 self-regulate the lengths of their respective zones. The HAPs do not possess signal sequences (Homma et af., 1990a) and do not appear to be processed (Homma et af., 1985) except for removal of the N-terminal methionine residues in H A P l and HAP2 (Homma et af., 1990a). Their high degree of sequence similarity to flagellin and the other axial proteins indicates that they are exported by the flagellar-specific pathway (Homma et al., 1990a,b; see Section 1V.A). Strains with mutations in the flagellin genes excrete all three HAPs, while mutants with defects in any of the three HAP genes excrete flagellin and the two non-mutant HAPs (Homma et af., 1984b; Homma

flbC flbD

FliF (switch)

f/bB f/iZ FlgB (rod)

FlgC (rod)

6

Rivet

A

-@--

(HAP11 FlgK

(HAP31 FlgL

HBB complex

(HAP21 FliD __t

- ---

(f ilament) Flit f G L r -a n Je

Peptidoglycan layer

C;toplasmic

membrane

FIG. 10. Pathway for flagellar assembly. Proteins necessary for individual steps are indicated above the respective arrows; where only a gene is indicated (i.e. in italics), the gene product has not yet been identified biochemically. Structures corresponding to the proteins and genes are given in parentheses, where known. The set of genes in the square brackets are necessary for formation of the M rinds rindrod complex (“Rivet”), but their roles in assembly have not been characterized further. The M and S rings are arbitrarily shown as assembling together. Whether the S ring is a separate structure has not been established. Based on data from Suzuki et al. (1978), Suzuki and Komeda (1981), Homma et al. (1986) and Jones and Macnab (1990). From Jones and Macnab (1990).


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and Iino, 1985b), indicating that ordered assembly of these components is not controlled at the level of subunit export. The HAPs are also exported from wild-type flagella (Homma and Iino, 1985b).Unlike the subunits of the hook and rod (see above and Section IV.C), there may be no control over export of HAPs and flagellin other than their relative levels of synthesis. Although all three HAPs are excreted into the medium, flagellin is not (Homma et al., 1984b), despite the fact that it is exported at levels at least several hundred times that of the HAPs (Homma and Iino, 1985b). This indicates that the terminal HAP2 cap must somehow be capable of efficiently distinguishing between flagellin and the HAPs, and allows passage only of the latter. The continued export of HAP2 monomers from wild-type flagella may indicate that the cap formed by HAP2 is a dynamic structure, its component subunits being continuously replaced during filament growth (Ikeda et al., 1987). It is much more difficult to envisage continued export of HAP1 and HAP3 in this way, however. Because the presence of HAP2 at the distal end of the filament is required for filament elongation (see Section IV.A), continued export of HAP2 may simply serve to salvage a flagellum whose external portion has been broken off, by enabling regrowth of the filament. Alternatively, it may be preferable for the cell to waste a few hundred copies of the HAPSrather than having to develop an export-control system capable of distinguishing them from flagellin. C. BASALBODY

Examination of basal-body precursors produced by various Fla- mutants (Suzuki et af.,1978; Suzuki and Komeda, 1981) and analysis of basal bodies assembled by temperature-sensitive Fla- mutants radiolabelled in vivo (Jones and Macnab, 1990) show that the basal body is assembled in a proximal-to-distal fashion (Fig. 10). On the basis of these studies, basalbody assembly is posited to consist of four major stages. In the first, the M ring assembles in the cytoplasmic membrane in conjunction with two proteins with apparent molecular weights of 23,000 and 26,000. The genes encoding these two proteins are not known, but their synthesis is controlled by the flagellar master operon, indicating that they are flagellar proteins (Jones and Macnab, 1990). They may serve to nucleate M-ring assembly or to plug the hole in the cytoplasmic membrane formed by this ring. Concurrent with this and subsequent stages, monomers of the P and L rings are exported, presumably to their respective destinations in the periplasm and outer membrane. These proteins, FlgI and FlgH, are unique among the known flagellar proteins in having canonical signal sequences

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(Homma et al., 1987b; Jones et al., 1989). They are exported via the standard signal peptidase-dependent pathway (Homma et al., 1987b,c), unlike other flagellar components which are external to the cytoplasmic membrane (i.e. the rod, hook, HAPs and filament). The second stage is presumed to involve assembly of the switch complex and of the export machinery necessary for controlling the ordered construction of the axial components of the flagellum, namely the rod, hook, HAPs and filament. The third stage of assembly involves construction of the rod. The labelling patterns of the four proteins identified as rod components in mutants in several rod genes (flgC,fEgF,flgG) indicate that the rod is only a metastable structure at this stage (Jones and Macnab, 1990). Consistent with this, no basal-body precursor structures were observed from strains with mutations in these genes (Suzuki et al., 1978; Suzuki and Komeda, 1981). The rod proteins are presumably exported via the flagellar-specific export pathway used by the other axial flagellar components (Homma et al., 1990b; see Section 1V.A). Like the hook protein (see Section IV.B), the rod proteins are probably no longer exported once the rod is completed. The final stage of basal-body assembly consists of nucleation of the P and Lrings onto the rod. The labelling patterns of the rod proteins inflgZ(P-ring) mutants indicates that they are not stably assembled in the absence of the P ring (Jones and Macnab, 1990). On the other hand, some particles consisting of the M and S rings and an apparently normal rod have been observed inflg1 mutants (Suzuki et al., 1978; Suzuki and Komeda, 1981). Resolving this discrepancy must await isolation of the precursor structures from these mutants in quantities sufficient for biochemical analysis. As the P- and L-ring proteins probably do not exist in the cell envelope as heterodimers (Jones and Macnab, 1990), the L ring presumably assembles onto the completed P ring. AlthoughflgH (L-ring) mutants fail to assemble structures distal to the rod (Suzuki et al., 1978; Suzuki and Komeda, 1981), the L ring appears to contact directly only the P ring (Stallmeyer et al., 1989), so it is not clear how failure of L-ring assembly blocks further construction. One possibility is that assembly of the L ring alters the structure of the rod, perhaps by conformational changes via the P ring, to allow assembly of later structures. Evidence exists for propagation by the rod of the effect of a mutation in the M-ring protein (Okino et al., 1989). While both the P- and L-ring proteins are normally necessary for flagellar assembly, overproduction of the hook protein of S.typhimurium in E. coli fEgH (L-ring) and fEgZ (P-ring) mutants can override the block caused by these mutations, resulting in formation of weakly functional flagella lacking the L ring or the entire outer cylinder, respectively (Jones et al., 1987;


149

Ohnishi etal., 1987). Similar defective flagella are also formed, albeit rarely, in the absence of elevated hook-protein levels (Suzuki et al., 1978; Suzuki and Komeda, 1981). FlgD appears to modify the distal end of the rod, making it competent for hook assembly (Suzuki etal., 1978). It is not known whether FlgD functions enzymically or as the terminal structural component of the rod, nor is it known whether FlgD normally acts before or after assembly of the outer cylinder. There are still a number of flagellar gene products, about 11, whose roles and placement in the assembly pathway have not yet been defined (Suzuki et al., 1978; Suzuki and Komeda, 1981; Jones and Macnab, 1990). All that is known is that they appear to function early in the pathway, being required prior to rod assembly. These proteins may be regulatory or scaffolding proteins, or form part of the export-control machinery. In B. subtilis, synthesis of flagella and autolysins (enzymes which cleave bonds in the peptidoglycan layer) are linked, suggesting that limited degradation of the peptidoglycan may be necessary for passage of the basal body through this layer (Fein, 1979). No such function has been described for E. coli or S . typhimurium, but the product of one or more of the flagellar genes not examined in the assembly studies may serve a homologous function in these species. D. MOT AND SWITCH COMPLEXES

1. Mot Complex

The motility proteins, MotA and MotB, can assemble after the flagellum is in place, as shown by experiments in which addition of MotA or MotB, supplied from bacteriophage or plasmids, brought about rotation of preexisting, paralysed flagella on appropriate Mot- mutant cells (Silverman et al., 1976; Block and Berg, 1984; Blair and Berg, 1988). Whether the Mot proteins normally assemble at a specific point in the flagellar assembly pathway in vivo is not known but seems unlikely. In view of these results, and because only the incorporation of MotB into the inner membrane is sitelimited (Stader et d., 1986; Wilson and Macnab, 1988), it is likely that the two proteins normally assemble separately into the membrane and subsequently combine to form a complex.

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2. Switch Complex Because fia alleles of all three switch genes have been found (Yamaguchi et al., 1972,1986a; Kutsukake et al., 1980), all three proteins are necessary for assembly of the flagellum. All three function very early in the assembly pathway, but may assemble at different points along it: FliG is required for stabilization of the M ring, while FliN may function somewhat later, before initiation of rod assembly (Fig. 10; Jones and Macnab, 1990). FliM probably assembles at some point prior to completion of the rod (Suzuki and Komeda, 1981), although this is not certain, since the lesions of both of the putativefliM mutants examined could have been in the recently discovered adjacent gene fliL (Kuo and Koshland, 1986; Kihara et al., 1989). E. OTHER SYSTEMS AFFECTING FLAGELLAR ASSEMBLY

1. Flagellar Assembly and the Cell Cycle Linkage of flagellar formation to the cell cycle has been extensively studied in the dimorphic bacterium Caulobacter crescentus (for reviews see, for example, Champer et al., 1986; Newton, 1989). Motile C. crescentus has a single polar flagellum which is shed at a specific point in the cell cycle and replaced by a stalk with which the cell anchors itself to a surface. This stalked cell proceeds to divide, producing one daughter cell with a new flagellum and another with the stalk from the parent cell. The relationship between flagellar assembly and cell cycle in bacteria lacking such a well-defined programme of synthesis and division has been more difficult to demonstrate. Using E. coli temperature-sensitive mutants defective in various cell-division genes, Nishimura and Hirota (1989) showed that flagellar assembly is linked to the cell cycle by the mechanism that controls cell division. This linkage is mediated at the transcriptional level, presumably by effects on the flagellar master operon (see Section 11). By comparison of cell length and the number of flagella per unit length, Nishimura and Hirota (1983) have proposed that initiation of flagellar formation occurs preferentially just prior to septum formation. However, these authors counted flagella on cells by electron microscopy, so that only flagella of a certain minimum length would have been detected. While the precise timing of flagellar assembly relative to the cell cycle is not known, it is clear that there must be some connection, as the number of flagella observed per unit cell length is not constant (Nishimura and Hirota, 1983), as would be expected if cell growth and initiation of flagellar formation were not coupled.


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Continuous elongation of flagellar filaments on S. typhimurium over several generations of growth has been observed by dark-field microscopy (T. Noguchi, S.-I. Aizawa and K. Namba, unpublished observation). The rate of filament elongation decreased rapidly with increasing filament length, confirming observations by Iino (1969). There was no cessation of filament elongation before or after cell division, indicating that filament elongation continues independently of the cell cycle. 2. Flagellar Assembly and the Cell Surface Very little is known about the relationship between flagellar assembly and assembly of other cell-surface structures. Flagellar formation is dependent on synthesis of the peptidoglycan layer (Vaituzis and Doetsch, 1965,1966). Sphaeroplasts formed by penicillin treatment, or strains incapable of synthesizing diaminopimelic acid (a constituent of the peptidoglycan) grown in the absence of the acid, fail to form new flagella, suggesting that flagellar assembly requires an actively growing cell wall (Vaituzis and Doetsch, 1965, 1966). Old flagella are restricted to regions of the sphaeroplast retaining some portion of the cell wall (Vaituzis and Doetsch, 1965). Flagellin is not found in penicillin-treated cells (Vaituzis and Doetsch, 1966; McGroarty et al., 1973), but the level at which its synthesis is blocked has not been determined. Mutations in galU result in changes in the composition of the outer membrane (Fukusawa et al., 1962; Sundararajan et al., 1962); the number of flagella on galU mutant cells is also greatly decreased (Komeda etal., 1977a). Deep rough mutants of S. ryphimurium, which have an altered cell surface, lack flagellin as well as flagella (Ames et al., 1974; Irvin et al., 1975). Defects arising from galU mutations can be suppressed by mutations in flhA (Komeda et al., 1977a); FlhA- cells allow assembly of the M ring, which is located in the inner membrane, but not of the rod or subsequent components (Jones and Macnab, 1990), so how the FlhA protein suppresses effects arising from changes in the outer membrane is not clear. The compensating mutations may be in PhE, a newly discovered gene downstream ofjlhAin the same operon (P. Matsumura, personal communication; see Table 1).

3. Other Factors Affecting Flagellar Assembly A single mutation (probably in the gene encoding the p-subunit of RNA polymerase) prevents cell growth at high temperatures and decreases

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flagellation at the permissive temperature (Yamamori et al., 1977). These cells are somewhat longer than those of the parent strain, indicating that the mutation may also have an effect on cell division. The transmembrane proton-motive force is necessary for flagellar assembly (Galperin et al., 1982). An intact electron-transport chain is also required for flagellar formation in E. coli, whether the cells are grown aerobically or anaerobically (Bar Tana et al., 1977; Hertz and Bar-Tana, 1977); this requirement is probably an indirect result of effects on CAMP levels (Hertz and Bar-Tana, 1982). Growth at high temperatures inhibits flagellar formation in several bacterial species, including S. typhimurium (Quadling and Stocker, 1962) and E. coli (Morrison and McCapra, 1961). This effect is a result of blockage of flagellar gene transcription at higher temperatures (McGroarty et al., 1973), apparently at the level of the flagellar master operon and friA (Silverman and Simon, 1977), and may be a result of a diminished protonmotive force (Galperin et al., 1982). V. Motor Function

In general, motor function can be characterized by parameters such as the power source, maximum speed and the amount of torque it generates (Table 2). Because of the small size of the bacterium, the sorts of measurement techniques useful for characterizing motor function in automobiles, for example, are of no use in analysing the bacterial motor. For the bacterium, inertial forces are essentially non-existent and viscous forces dominate (see, for example, Berg, 1974; Berg and Purcell, 1977; Purcell, 1977; Yates, 1986), allowing us to deduce some of the operating parameters of the flagellar motor using less direct techniques. The most widely used approach involves the use of tethered cells, which are flagellated cells attached to a glass microscope slide by an antibody directed against the hook or filament (see Fig. 12(a); Larsen et al., 1974a; Silverman and Simon, 1974~).Because the tethering flagellum is no longer free to rotate, the torque generated by the motor rotates the cell body, which is easily observed. Because they lack endogenous energy reserves and have no outer membrane, a Gram-positive Streptococcus sp. (Manson et al., 1977) and Bacillus subtilis (Matsuura et al., 1977) have been used for most of the mechanistic studies to be described. Although the characteristics of flagellar rotation in these species and in E. coli and S. typhimurium are in all probability essentially identical, this remains to be demonstrated. For further discussions of motor function, see the articles by Macnab (1987b), Khan (1988, 1990) and Eisenbach (1990).


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TABLE 2. Parameters of flagellar motor function Energy source

Proton-motive force across the cytoplasmic membrane

Flagellar rotation speed

Load-dependent until motor saturates; on free-swimming cells, maximum observed speed is about 300 Hz

Number of protons per revolution

About loo0

Efficiency

Variable; less than 5% at low load, but 5&100% at high load

Torque output per motor

About lo-" dyne cm

Power output per motor

About lo-'" W at 20 Hz

Power per cell

About lo-'' W under normal swimming conditions

Cost to the cell of flagellar operation

About 0.1%of total energy expenditure under growth conditions

Cost to the cell of flagellar synthesis

About 2% of biosynthetic energy expenditure

The table is modified from Macnab (1987b), using data from Berg (1974), Lowe etal. (1987) and Meister et nl. (1987, 1989).

A . PARAMETERS OF MOTOR FUNCTION

1. Power Source Flagellar rotation in bacteria is not driven by consumption of ATP (Larsen et al., 1974b). In most species it is powered by the transmembrane protonmotive force (Belyakova et af.,1976; Manson et af.,1977,1980; Matsuura et af., 1977; Glagolev and Skulachev, 1978; Shioi et af., 1978; Eisenbach and Adler, 1981; Ravid and Eisenbach, 1984b). In alkalophilic species, sodium ions are used instead of protons (Hirota et al., 1981; Imae and Atsumi, 1989). The proton-motive force has two components, the membrane potential (Ay) and the transmembrane pH gradient (ApH). Either component can be specifically eliminated, either by adjusting the pH value of the medium to match that of the cytoplasm or by collapsing the membrane potential with ionophores (see, for example, Larsen et af., 1974b; Manson et af., 1977; Matsuura et al., 1977; Glagolev and Skulachev, 1978; Khan and Macnab, 1980b). Such experiments have shown that flagellar rotation can be driven by either component alone, that the threshold for rotation is between -8 and -18mV in Streptococcus sp. (Manson et af., 1980; Berg et al., 1982), but about

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-3OmV in B. subtilis (Khan and Macnab, 1980b; Shioi et al., 1980), and that rotation speed is proportional to the proton-motive force imposed, up to some saturating value (Khan and Macnab, 1980b). Tethered cells of wild-type Streptococcus sp. rotate in response to an applied proton-motive force of either sign (Manson et al., 1980). Their direction of rotation is the same with either sign as a result of cytoplasmic pH-value effects on the chemotaxis system. Mutant cells that are insensitive to cytoplasmic pH-value changes rotate in opposite directions in response to proton gradients of opposite polarity (Khan and Berg, 1983). In contrast, the motors of E. coli and S. typhirnuriurn cell envelopes (cells which have been lysed and resealed; Eisenbach and Adler, 1981) fail to function upon reversal of direction of the applied proton-motive force (Ravid and Eisenbach, 1984a; Ravid et al., 1986). This may reflect species differences or the possibility that cell envelopes lack some cytoplasmic component, present in whole cells, which might be necessary for enabling rotation in response to an inverted proton-motive force. A fixed number of protons probably drives each rotation of the motor (Manson et al., 1980); this number appears to be of the order of lo00 and is independent of motor speed (Berg, 1974; Meister et al., 1987, 1989).

2. Speed and Torque The maximum rotation speed of tethered cells is only about 15 Hz (Berg, 1974; Berg and Turner, 1979; Blair and Berg, 1988; Lapidus et al., 1988), whereas filaments in bundles on swimming cells have been observed to rotate at speeds up to almost 300 Hz (Lowe el al., 1987), indicating that the viscous drag on the tethered cells imparts a significant load to the motor of the tethering flagellum. Under these high-load conditions the motor runs at constant torque, the value for which is proportional to the proton-motive force, while its speed is inversely proportional to the viscosity of the medium (Berg and Turner, 1979; Manson et al., 1980; Khan el al., 1985). The efficiency of energy conversion in tethered motors is between 0.5 and 1.0 (Meister er al., 1987). The torque is independent of temperature and deuterium-isotope effects, implying that chemical bonding is not the rate-limiting step in motor rotation (Khan and Berg, 1983). The motors of free-swimming cells, operating under low-load conditions, run at constant speed (Berg et al., 1982). The speed and motor rotation rate of swimming cells increase linearly with temperature (Lowe et al., 1987). Torque decreases with increasing rate of rotation for these cells, suggesting that the motor does not run at constant power (Lowe et al., 1987). The


155

energy-conversion efficiency of the motors in free-swimming cells is very low, less than 5% (Meister et al., 1987). Under low-load conditions, the motor rotation rate is lowered by about 20% in the presence of deuterated water; the reason for the difference in isotope effect observed in motors operating with heavy and light loads is not known, but may point to differences in the rate-limiting step in these respective regimes (Meister et al., 1987).

3. Current Flow When the load on a flagellar motor increases, its rotation speed slows down proportionately, implying that the torque generated is constant at high load (Berg and Turner, 1979). If the load is further increased, rotation will eventually stop when the motor torque and the external load are balanced. This stall torque is the maximum torque that the motor can generate. Meister and Berg (1987) measured the torque required to halt the rotation of tethered cells and compared it with the torque of the motor operating over a wide range of proton-motive force and p H values. Although the absolute value for the stall torque was not determined, it was found that the stall torque and the running torque were correlated. This indicates that the flagellar motor operates by direct current. 4. Periodicities in Motor Function

Given the small size of the flagellar motor, it is possible that such parameters as rotation and torque generation are quantized. So far, attempts to detect stepping in the flagellar rotation of wild-type cells have failed (Berg, 1976; Berg et al., 1982). Calculations suggest that, if there are discrete stepping events in the rotation of the motor, the number of such steps must be of the order of 400 (Berg et al., 1982). One difficulty with these studies is that the elastic properties of the filament damp any periodicities which might be observed (Berg, 1976; Berg et al., 1982). De-energized, tethered cells of Streptococcus sp. stop at discrete positions, with a periodicity of five or six (Khan et al. , 1985). This locking of the motor is a result of energy barriers which prevent rotational diffusion of the cell; the energy of these barriers must be significantly greater than kT, where k is the Boltzmann constant and T is the temperature (Berg, 1976). Because the energy derived from the transport of a single proton is less than 2kTat the point where rotation becomes smooth (about -35 mV), several protons must act in concert to drive the motor (Khan, 1988,1990). It should be noted that the observation of motor locking in de-energized Streptococcus sp. is not consistent with an earlier report by Ishihara et al.

156


(1981), who found that the flagella of de-energized S. typhimun'um could freely rotate in reponse to a flow of viscous fluid past the cell body. Periodicity in torque generation has also been demonstrated. Paralysed motors of motA and motB mutants of E. coli were repaired by inducing synthesis of the missing protein from plasmids (Block and Berg 1984; Blair and Berg, 1988). Shortly after induction, the tethered mutant cells began to rotate, increasing their speed over the course of several minutes. This increase occurred in about eight discrete steps (Blair and Berg, 1988), and was ascribed to the stepwise incorporation of force generators consisting of MotA and MotB. Eisenbach et al. (1990) observed that both the rotation rate and angular velocity of tethered E. coli fluctuate significantly, which they suggest may reflect the gain and loss of these force generators.

5. Rotational States The flagellar motor rotates both clockwise and counterclockwise (Silverman and Simon, 1974~).Swimming motility corresponds to counterclockwise rotation, and tumbling corresponds to clockwise rotation (Larsen et al., 1974a). Although the cell normally swims for a period of the order of one second, after which it then tumbles for a period of the order of 0.1 second (Berg and Brown, 1972; Berg and Tedesco, 1975; Macnab and Ornston, 1977), the period of time each motor spends in clockwise and counterclockwise rotation under normal conditions is not known. Attempts to estimate the number of flagella which must be in simultaneous clockwise rotation in order to cause the cell to tumble have not been successful (Khan and Macnab, 1980a; Ishihara et al., 1983). The duration of the clockwise and counterclockwisestates of the motor are affected by tactic stimuli (see, for example, Tsang et al., 1973; Larsen et a f . , 1974a; Berg and Tedesco, 1975), mutations in components of the chemotaxis system (see, for example, Berg and Brown, 1972; Larsen et al., 1974a; Szupica and Adler, 1985) and changes in the proton-motive force (Miller and Koshland, 1977; Taylor et al., 1979). Motors in wild-type cells and cellenvelope preparations rotate exclusively counterclockwise in the absence of signals from the chemotaxis system, but this inherent bias can be shifted to various extents by mutations in the switch complex (Ravid and Eisenbach, 1984a; Szupica and Adler, 1985; Ravid et al., 1986; Wolfe et al., 1987). The rate of clockwise rotation is somewhat faster than that of counterclockwise rotation (Eisenbach et al., 1990). The counterclockwise rate increases in the presence of attractants, apparently by suppression of pauses (see below), but the clockwise rate is not affected by repellent addition (Eisenbach et al., 1990). The motor switches abruptly between the clockwise and counterclockwise

0

0.5

1.0

1.5

2 .o

Time (sec)

FIG. 11. Analysis of the rotation of a tethered bacterial cell. (a) Negative image of a radially asymmetric slit (the openings are shown in black) and a tethered cell body (crosshatched). The slit was centred over the point of rotation of the cell body and the photons passing through the slit were recorded over time. As the cell body rotates counterclockwise, the amount of light passing through the slit will gradually diminish, then increase abruptly as the cell body passes the "6 o'clock'' position on the slit. The pattern of light detected will be reversed during clockwise rotation. (b) Sample data. Light intensity, measured as photons passing through the slit, is indicated along the vertical axis. The cell rotated clockwise during the period indicated by the arrows. Note the constant rotation rate and the abrupt change in the direction of rotation. The gate time for photon counting was ten milliseconds. The figure is reproduced courtesy of Y. Magariyama.

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C. J. JONES AND

S.4. AIZAWA

rotational states (Fig. 11; Berg, 1974, 1976); switching occurs in less than one millisecond (Kudo et al., 1990). Switching to the clockwise state is caused by binding of the CheY protein to the switch complex; CheZ enhances switching to the counterclockwise state by interfering with CheY binding (Ravid et al., 1986; Kuo and Koshland, 1987; Wolfe et al., 1987). The switching events of individual motors on a single cell are not coordinated (Ishihara et al., 1983; Macnab and Han, 1983). The switching probabilities of the motor are inversely related; that is, as the probability of switching from clockwise to counterclockwise rotation decreases, the probability of switching from counterclockwise to clockwise rotation increases proportionately (Khan and Macnab, 1980a). In addition to the transient tactic effects of small changes in proton-motive force already referred to, large decreases in the magnitude of this force result in permanent changes in the switching probability of the motor, shifting in favour of counterclockwise rotation (Khan and Macnab, 1980a; Miller and Koshland, 1980). Whether this reflects a direct effect of the proton-motive force on the conformation of one or more switch proteins (cf. Richard and Miller, 1990) or an indirect result of effects on the chemotaxis pathway is not known. A third state of the normal motor, namely pausing, consists of very brief, intermittent episodes in which the motor does not seem to move, although the time period involved is so short that detailed behaviour cannot be resolved. The flagellar motors of other species stop as part of their normal function (Alam and Oesterholt, 1984; Alam et al., 1984; Armitage and Macnab, 1987; Gotz and Schmitt, 1987; McCain et al., 1987) but, until recently, pausing in the rotation of flagella on E. coli and S. typhimurium was generally considered to be an artefact of the cell-tethering technique, ascribed to the cell body sticking to the surface of the slide. This has been shown not to be correct; the motors of free-swimming cells also pause (Eisenbach et al., 1990). Eisenbach and his coworkers have analysed motor pausing quantitatively (Lapidus et al., 1988; Eisenbach et al., 1990). Although the flagella of cells whose energy levels have been depleted slow down and eventually stop, energy levels do not appear to be responsible for pausing (Lapidus el al., 1988). Pausing is independent of the sense of rotation of the motor and requires the operation of at least part of the chemotaxis system. Repellents induce longer and more frequent pauses, in addition to clockwise rotation; attractants abolish pausing and clockwise rotation. Examination of the behaviour of a wide variety of Che- mutants (which have altered switching probabilities) showed that reversal frequency and pausing frequency were closely correlated. The data indicate that pausing events are unsuccessful

THE BACTERIAL FLAGELLUM

AND FLAGELLAR

MOTOR

159

attempts to switch the sense of motor rotation (Eisenbach et af., 1990). Analysis of the rotation of single flagella on various switch mutants has also implicated the switch complex in pausing (Kudo et al., 1990). This three-state system may still be insufficient to describe the flagellar motor. High-resolution video analysis of the rotation of tethered wild-type cells indicates that both clockwise and counterclockwise rotational states consist of at least two components (Kuo and Koshland, 1989). B. MODELS

Many models for the rotation mechanism of the flagellar motor have been developed (see, for example, Glagolev and Skulachev, 1978; Macnab, 1979b; Berg and Khan, 1983; Oosawa and Hayashi, 1983; Mitchell, 1984; Wagenknecht, 1986; Kobayasi, 1988; Lauger, 1988; Fuhr and Hagedorn, 1989; Murata et al., 1989). Most of these models assume a direct coupling of proton transfer to torque generation, but this has not been demonstrated. The nature of this coupling, i.e. whether a fixed number of protons serve to drive the motor through each rotation, is also not known, but most of the available evidence favours a tight-coupling mechanism (Meister et al., 1987, and references therein). On the other hand, recent studies on the effects of mutant MotA proteins on motor function suggest that coupling of proton transfer and motor rotation may not be obligatory, at least for tethered cells (Blair and Berg, 1990). For a discussion of some of these models, see Khan (1990). C. RECENT TECHNICAL ADVANCES

As mentioned above, the flagellar motor is too small for direct measurements to be carried out. However, techniques such as cell tethering (Fig. 12(a)) have provided some understanding of this device, and several recent innovations promise to further this understanding. These techniques are all specifically aimed at analysing the flagellar motor. Improvements in the capabilities of more generally applicable devices, such as computers and video recording equipment (see, for example, Kuo and Koshland, 1989), will also contribute to future investigations.

a

C

FIG. 12. Schematic diagram illustrating the methods used to analyse the flagellar motor. (a) Tethered cells. If a cell is attached to a glass slide by a single flagellum using antiflagellin antibodies, rotation of the flagellar motor drives rotation of the cell body. Observing the rotating cell and measuring some characteristics of the motor are relatively straightforward, but the motor is here operating under extremely high load. (b) Free-swimmingcell analysis. By analysing the vibration and counter-rotation of freeswimming cells under different load conditions, Lowe et al. (1987) have been able to infer motor speed and torque. (c) Laser dark-field microscopy. If an individualflagellum on a cell stuck to a glass slide is illuminated from one direction, the rotation of that flagellum can be observed and measured with a sufficiently sensitive detector. In this method the functioning of a single motor under approximately normal load conditions can be observed (Kudo el al., 1990).


161

1. Free-Swimming Cell Analysis

The swimming speed of a motile bacterium will increase with the rotation speed of filaments in the flagellar bundle. Flagellar rotation speed can be deduced by analysing the vibration frequencies of swimming cells caused by asymmetries in flagellar orientation (Fig. 12(b); Lowe et al., 1987). In this method, images of swimming cells are focused on a photomultiplier tube and the vibration frequencies of the cells determined by Fourier analysis of the output signal. Rotation rates as high as 268 Hz (at 32°C) have been determined for the flagellar bundles of swimming E. coli (Lowe et al., 1987). 2. Laser Dark-Field Microscopy

High-intensity dark-field microscopy has been employed to visualize single filaments on cells. If freely rotating flagella on cells stuck to a glass slide (Fig. 12(c)) are illuminated at right angles to the filament axis, the filament appears as a line of dots moving away from the cell. By using a laser to illuminate the flagella and focusing the resulting image through an optical slit onto a photomultiplier tube, it is possible to measure the rotation rates of single flagella on cells (Kudo et al., 1990). This system has a temporal resolution of the order of 0.1 millisecond. The maximum rotation speed of flagella observed by this method was about 210 Hz at 37°C.

3. Optical Tweezers Even with the thinnest glass needle, bacteria cells are too small to handle. Block et al. (1989) used a laser-based optical particle trap (Ashkin et al., 1987) to spin tethered cells under a variety of conditions in order to investigate the torsional properties of the flagellum. Their results are consistent with the filament being rigid and attached to a more flexible hook, although they do not rule out the possibility that this may reflect nonlinear properties of the filament itself. Using this system it is possible to rotate a tethered cell at speeds greater than the zero-load speed or backwards. Meister et al. (1989) pointed out that results from such experiments would help discriminate between current models of motor mechanism.

VI. Summary The bacterial flagellum is a complex multicomponent structure which serves as the propulsive organelle for many species of bacteria. Rotation of the

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helical flagellar filament, driven by a proton-powered motor embedded in the cell wall, enables the flagellum to function as a screw propeller. It seems likely that almost all of the genes required for flagellar formation and function have been identified. Continuing analysis of the portions of the genome containing these genes may reveal the existence of a few more. Transcription of the flagellar genes is under the control of the products of a single operon, and so these genes constitute a regulon. Other controls, both transcriptional and post-transcriptional, have been identified. Many of these genes have been sequenced, and the information obtained will aid in the design of experiments to clarify the various regulatory mechanisms of the flagellar regulon. The flagellum is composed of several substructures. The long helical filament is connected via the flexible hook to the complex basal body which is located in the cell wall. The filament is composed of many copies of a single protein, and can adopt a number of distinct helical forms. Structural analyses of the filament are adding to our understanding of this dynamic polymer. The component proteins of the hook and filament have all been identified. Continuing studies on the structure of the basal body have revealed the presence of several hitherto unknown basal-body proteins, whose identities and functions have yet to be elucidated. The proteins essential for energizing the motor, the Mot and switch proteins, are thought to exist as multisubunit complexes peripheral to the basal body. These complexes have yet to be identified biochemically or morphologically. Not surprisingly, flagellar assembly is a complex process, occurring in several stages. Assembly occurs in a proximal-to-distal fashion; the basal body is assembled before the hook, and the hook before the filament. This pattern is also maintained within the filament, with monomers added at the distal end of the polymer; the same is presumably true of the other axial components. An exception to this general pattern is assembly of the Mot proteins into the motor, which appears to be possible at any time during flagellar assembly. With the identification of the genes encoding many of the flagellar proteins, the roles of these proteins in assembly is understood, but the function of a number of gene products in flagellar formation remains unknown. To reach their sites of assembly, the axial components of the flagellum utilize a unique pathway, travelling through the hollow core of the flagellum. Many aspects of this transport pathway, including signals marking the proteins which use it, how proteins transported at different stages in assembly are differentiated, and how they are propelled through the flagellum, are not understood and are under investigation. The motor which drives flagellar rotation is powered by the flow of protons across the cytoplasmic membrane. It can rotate both clockwise and


163

counterclockwise, and switches between these states rapidly. The motor also pauses intermittently for brief intervals; these pauses are believed to result from failed attempts to switch the direction of rotation. Many models attempting to explain the mechanism of motor rotation have been presented, but we do not yet have enough data to decide which is correct. Although most previous studies of motor function were limited to analysing its behaviour under abnormally high loads, recent technical developments promise to rapidly expand our understanding of how the world’s smallest rotary engine has been designed to work. VII. Acknowledgements

We thank I. Kawagishi, K.Kutsukake, R. M. Macnab and P. Matsumura for communicating results prior to publication. We also thank T. Akiba, K. Namba, K. Oosawa and F. Vonderviszt for comments on an earlier version of the manuscript. REFERENCES

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Structure and function of the bi-directional bacterial flagellar motor.

Loose coupling in the bacterial flagellar motor.

The bacterial flagellar motor and its structural diversity.

Crystal structure of the flagellar accessory protein FlaH of Methanocaldococcus jannaschii suggests a regulatory role in archaeal flagellum assembly.

Effect of the MotB(D33N) mutation on stator assembly and rotation of the proton-driven bacterial flagellar motor.

[Structure and function of the bacterial flagellar type III protein export system in Salmonella ].

The cytoplasmic component of the bacterial flagellar motor.

Switching dynamics of the bacterial flagellar motor near zero load.

Impact of fluorescent protein fusions on the bacterial flagellar motor.

Molecular architecture of the bacterial flagellar motor in cells.

Form, Fabric, and Function of a Flagellum-Associated Cytoskeletal Structure.

Bacterial Flagellar Motor Switch in Response to CheY-P Regulation and Motor Structural Alterations.

Structure and assembly of filamentous bacterial viruses.

Assembly dynamics and the roles of FliI ATPase of the bacterial flagellar export apparatus.

Rotation and switching of the flagellar motor assembly in Halobacterium halobium.

Load-sensitive coupling of proton translocation and torque generation in the bacterial flagellar motor.

Internal and external components of the bacterial flagellar motor rotate as a unit.

Energy transduction in the bacterial flagellar motor. Effects of load and pH.

Building a flagellum outside the bacterial cell.

The role of the bacterial flagellum in adhesion and virulence.

Type III secretion systems: the bacterial flagellum and the injectisome.

Genetic analysis of the bacterial flagellum.

Coordinated switching of bacterial flagellar motors: evidence for direct motor-motor coupling?

Coevolved Mutations Reveal Distinct Architectures for Two Core Proteins in the Bacterial Flagellar Motor.