The Gram-negative soil bacterium Myxococcus xanthus has a complex life cycle that includes vegetative growth and development (FIG. 1). During its life cycle, a myxo-bacterium exhibits social interactions of the type that are usually associated with more complex eukaryotic cells1, which has led to the use of M. xanthus as a model system for the study of social interactions and their regu-lation. M. xanthus grow by scavenging nutrients from decomposing soil and detritus, or by predation of other microorganisms2–5. M. xanthus cells lack flagella and are non-motile in liquid growth media, but can move on solid growth substrates at speeds of 2–4 µm per minute. This is extremely slow compared with other species such as Escherichia coli, which swims at a rate of ~50 µm per second6, and Flavobacterium johnsoniae, which glides at 5–10 µm per second7.
M. xanthus cells are usually present in biofilms that consist of cells that are organized into a series of layers. The clustering of cells into organized groups known as swarms (FIG. 2a) facilitates predation and food gathering, because numerous bacteria cooperate to produce antibi-otics and digestive enzymes8. These antibiotics and lytic enzymes kill and digest prokaryotic and eukaryotic micro-organisms. Indeed, an estimated 8% of the M. xanthus genome is dedicated to the production of secondary metabolites, and at least 18 gene clusters specify the pro-duction of polyketide antibiotics — almost twice as many as the number of genes that specify antibiotic production in Streptomyces coelicolor9, which is a model for antibi-otic production. Furthermore, the M. xanthus genome encodes numerous proteases, nucleases and lipases that function in the digestion of macromolecules.
Predation usually requires direct contact with the prey cells10, and this contact triggers the myxobacterial rippling response5,11 (FIG. 2b). Rippling is the coordinated
rhythmic movement of cells that is observed when myxobacteria feed on macromolecules or during the autolysis of cells that is associated with starvation and development11–13. Rippling creates ‘accordion waves’ (ReF. 14) (FIG. 2c) in which cells seem to form a travelling wave. However, when the cells of two waves contact each other, cell reversals are induced that result in the two waves reflecting off each other. Berleman et al.11 proposed that the function of travelling waves is to move cells back and forth, which might provide an efficient mechanism to ‘mop-up’ macromolecules from the surface and ensure efficient usage of growth substrates. Cells do not ripple when grown on rich media that lack macromolecules or lysed cells11.
When M. xanthus swarms cannot find sufficient nutrients or prey, they enter a developmental pathway that results in the formation of multicellular mounds, which develop into fruiting bodies (FIG. 1,2d). During this process, gene expression and the pattern of cell move-ments are highly regulated. Cells aggregate into streams that merge to form fruiting bodies that are 0.1–0.2 mm in height, and each contain 105–106 cells. In the fruit-ing bodies, most of the cells differentiate into spores, although some cells (about 10%), named peripheral rods, remain undifferentiated and are present as a monolayer of rod-shaped cells around and between fruiting bodies (FIG. 2e). Unfortunately, nothing is known about the ‘sig-nals’ that maintain this subpopulation of undifferenti-ated cells. Peripheral cells most likely function as ‘scout cells’ that identify new food sources, because they move about between fruiting bodies and maintain their veg-etative behaviour under starvation conditions14,15. The soluble nutrients that are released through digestion of food or prey by the peripheral rods might trigger spore germination in the fruiting body.
*Department of Molecular and Cell Biology, University of California, Berkeley, California 94720‑3204, USA. ‡Department of Biological Sciences, Virginia Polytechnic Institute and State University, Blacksburg, Virginia, USA. §Department of Microbiology, University of Iowa, Iowa City, Iowa 52242, USA. Correspondence to D.R.Z. and J.R.K. e‑mails: [email protected]; john‑[email protected]:10.1038/nrmicro1770Published online 8 October 2007
DevelopmentA programmed change in gene expression and morphology. In Myxococcus xanthus, this process is triggered by starvation and results in cellular aggregation, fruiting-body formation and sporulation.
RipplingCoordinated rhythmic movement of cells.
Cell reversalWhen a cell changes its direction along its long axis so that the leading cell pole becomes the lagging cell pole.
MoundAn early stage of development during which cells aggregate before sporulation.
Chemosensory pathways, motility and development in Myxococcus xanthusDavid R. Zusman*, Ansley E. Scott*, Zhaomin Yang‡ & John R. Kirby§
Abstract | The complex life cycle of Myxococcus xanthus includes predation, swarming, fruiting-body formation and sporulation. The genome of M. xanthus is large and comprises an estimated 7,400 open reading frames, of which approximately 605 code for regulatory genes. These include eight clusters of chemotaxis-like genes that define eight chemosensory pathways, most of which have dedicated functions. Although many of these chemosensory pathways have a role in controlling motility, at least two of these pathways control gene expression during development.
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Fruiting bodyThe final stage of Myxococcus xanthus development during which cells form a large aggregate and develop into environmentally resistant spores.
Peripheral rodA cell that maintains its rod shape and does not form a spore during development.
DZ2A wild-type laboratory strain of Myxococcus xanthus. It was obtained from the Berkeley microbiology laboratory culture collection in 1973. It is thought to be the parent strain of FB, DK101, DZF1 and DK1622.
FBA strain of Myxococcus xanthus that has been selected for its ability to form single colonies.
The final stages of development in M. xanthus are sporulation and, when nutrients become available, germination. Spores (FIG. 2e,f) are viable for long peri-ods of time and confer a strong survival benefit on starving cells. It has been proposed that sporulation in M. xanthus requires programmed cell death; cannibal-ism, it was postulated, could provide essential nutrients for the conversion of rod-shaped cells to mature spores in a starving population of cells16. However, develop-ment can also be triggered by a step-down in nutrition, rather than starvation, which might be a more normal transition, given what occurs in nature (sudden starva-tion is commonly used to trigger development in the laboratory, but is probably uncommon in nature)17. Thus, ‘developmental lysis’ might have more to do with severe starvation stress than it does as a requirement for sporulation, per se. notably, the amount of cell lysis during development varies between strains. For example, 20–25% of cells from the wild-type strain DZ2 exhibit developmental lysis11, whereas 80–90% of cells from strains derived from strain FB, such as DK1622, exhibit developmental lysis16.
The predatory and fruiting behaviours of M. xanthus depend on directed cell movements, so this Review focuses on the control of cell movements in myxobacte-ria and, in particular, on the eight chemosensory systems
that have been identified in the M. xanthus genome. Although some of these chemosensory pathways have a role in controlling motility, at least two of the pathways seem to be unrelated to motility and instead control gene expression during development.
Master regulators of cell movementour understanding of cell movement in M. xanthus began with the pioneering studies of Hodgkin and Kaiser18,19, who isolated hundreds of motility mutants that they analysed phenotypically and genetically. Their experiments revealed that motility in M. xanthus is controlled by two parallel and almost independent mul-tigene systems: system A (for adventurous) controls the movement of single cells, whereas system S (for social) controls the movement of groups of cells and generally depends on cell–cell contact (FIG. 3). These two motility systems are complex, require many genes and proteins, and are powered by different engines with different properties (BOX 1). Hodgkin and Kaiser did identify one gene, mglA (mutual gliding A), that is required for all myxobacterial cell movement18. Indeed, mglA mutants are phenotypically indistinguishable from double mutants that are defective in both A- and S-motility genes20. mglA is a 22-kDa protein that is a member of the Ras superfamily of small monomeric GTPases21. Surprisingly, Sar1, a yeast GTPase, can partially comple-ment the phenotype of a M. xanthus ∆mglA mutant. As Sar1 binds to microtubules22, mglA might interact with cytoplasmic microfilaments to directly affect motility. Recent work has shown that mglA interacts with AglZ, an A-motility protein23, and FrzS, an S-motility protein24. Furthermore, AglZ and FrzS are mislocalized in mglA mutants (T. mignot and D.R.Z., unpublished observa-tions). Although the specific role of mglA in motility is not understood, these different lines of evidence might indicate that it has a central role in controlling cell move-ments. Recently, nla24, a gene that encodes a putative ntrC-like σ54 transcription activator, was also found to be required for both motility systems25. This suggests that a transcriptional activator is common to both motility systems and might even be part of the mglA pathway.
Finding the right direction: the Frz systemThe frz (frizzy) genes (a cluster of seven genes) were discovered during a search for mutants that are defec-tive in cellular aggregation. Upon starvation, frz mutants cannot aggregate into fruiting bodies and instead form ‘frizzy’ filaments. frz mutants are also defective in vegeta-tive swarming26. Analyses of the individual movements of frz-mutant cells revealed that the frz genes control the frequency of cell reversals27, a function that is required for directed motility in gliding bacteria (BOX 2). Wild-type gliding M. xanthus cells reverse their direction of move-ment approximately every 7–8 min; net movement occurs because the interval between reversals can vary widely. most frz mutants rarely reverse. By contrast, some con-stitutively signalling frzCDc mutants reverse more often than wild-type cells27; these frzCDc mutants form tiny non-spreading colonies because individual cells exhibit no net movement.
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Vegetative growthcycle and swarming
Myxospore
Germination
Aggregation
Fruiting body
Mound
Starvation
Figure 1 | Life cycle of Myxococcus xanthus. Myxococcus xanthus cells are usually found on solid substrates. When nutrients are present, groups of cells (swarms) grow and divide and move outward in search of additional macromolecules or prey. Upon starvation, cells aggregate at discrete foci to form mounds and then macroscopic fruiting bodies. The rod-shaped cells in the fruiting bodies undergo morphogenesis and form spherical spores that are metabolically inactive and partly resistant to desiccation and temperature. Peripheral rods remain outside fruiting bodies and move as accordion waves in their search for food. When nutrients become available, the spores germinate and complete the life cycle.
DK1622A ‘wild-type’ laboratory strain of Myxococcus xanthus. It was derived by restoring full social motility to strain FB, which is thought to have originated by mutagenesis from strain DZ2. Strain DK1622 contains a ~250-kb deletion of unknown function.
S-motilityMovement of cells in groups that involves extension and retraction of the type IV pili.
CheWAn adaptor protein that links CheA to methyl-accepting chemotaxis protein receptors in bacterial chemotaxis.
Genetic and DnA-sequence analysis revealed that the seven frz genes are cotranscribed from divergent promoters and that they encode homologues to bac-terial chemosensory proteins found in E. coli and Salmonella enterica serovar Typhimurium (S. typh-imurium)28,29 (FIG. 4a; TABLe 1). The Frz chemosensory system comprises FrzCD (a cytoplasmic chemorecep-tor, originally thought to be encoded by two genes on the basis of phenotype but later shown to be encoded by one gene), FrzA and FrzB (two CheW homologues), Frze (a histidine-kinase response regulator fusion pro-tein), FrzF (a methyltransferase) and FrzG (a methyl-esterase) (FIG. 4a; TABLe 1). FrzZ consists of two response regulator (RR) domains that are connected by a linker region.
FrzCD, FrzA and the CheA domain of Frze consti-tute the core components of the Frz pathway, as they are essential for vegetative swarming, responses to repellents and directed movement during development. FrzB, FrzF, FrzG, FrzZ and the RR domain of Frze are required for vegetative swarming and development. on the basis of the E. coli model for bacterial chemotaxis, activated Frze kinase would stimulate cellular reversals, whereas
an inactive kinase would inhibit cellular reversals. Furthermore, adaptation to stimuli would be associated with the methylation and/or demethylation of the methyl-accepting chemotaxis protein (mCP) receptor FrzCD. This model is illustrated in FIG. 4b.
one major question that remains unresolved is the nature of the output of the Frz pathway. Recent work using a genetic screen and in vitro phosphorylation assays showed that FrzZ, a dual RR protein, is the primary out-put component of the pathway, as it is phosphorylated by Frze, a CheA–RR fusion protein30. Interestingly, the RR domain of Frze functions as a negative regulator of the CheA domain of Frze and controls its autophos-phorylation. However, it is not known which protein(s) regulates this regulator30. Furthermore, it is not known how reversal signals are transmitted to the two motility engines, the S-system engine and the A-system engine. We speculate that FrzS and AglZ might play roles in these output functions, although both of these proteins contain pseudo-receiver domains that are not phosphorylated31,32. A working model for the regulation of the two motility systems of M. xanthus through the Frz chemosensory system is presented in FIG. 5.
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a
d fe
cb
Figure 2 | Myxococcus cells: movement and developmental stages. a | Myxococcus xanthus cells move as groups by social motility (black arrow) and as single cells by adventurous motility (white arrow). Cells preferentially follow the paths of other cells. b | M. xanthus cells moving in ripples to consume Escherichia coli as prey11. Ripples have a wavelength of ~100 µm. Panels a and b courtesy of J.R.K. c | Peripheral rods form a monolayer of cells between fruiting bodies that can move as reversing accordion waves14. Panel c courtesy of O. Sliusarenko, University of California, Berkeley, USA. d | A M. xanthus fruiting body on animal dung95. Fruiting bodies are 0.1–0.2 µm in diameter. Panel d was first published in ReF. 96 (2006) Macmillan Magazines Ltd and was provided courtesy of Michiel Vos, University of Oxford, UK. e | An optical slice of a fruiting body. Note that the spores are contained in the fruiting body and peripheral rods appear as a monolayer around the fruiting body. Image courtesy of O. Sliusarenko, University of California, Berkeley, USA. f | A fruiting body that was cut to reveal the spores within. All scale bars are 15 µm. Panel f is reproduced with permission from ReF. 97 Elsevier.
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CheAA histidine kinase that is involved in bacterial chemotaxis and that phosphorylates a cognate response regulator such as CheY. Chemosensory pathways can usually be identified by a specific CheA.
ChemotaxisDirected movement towards attractants or away from repellents.
AdaptationA process whereby cells can adjust to the levels of an attractant or repellent. In bacteria this involves the methylation and/or demethylation of receptors or the dephosphorylation of response regulators. Adaptation allows bacteria to sense small changes in stimuli.
Methyl-accepting chemotaxis proteinA protein that is involved in bacterial chemotaxis and that senses attractants and repellents.
SlimePolysaccharide that contains material secreted by Myxococcus xanthus.
Does M. xanthus chemotax? Although the frz genes are homologous to the E. coli che genes, it was not clear that they performed similar functions. For example, the frz genes of M. xanthus cannot regulate flagellar rotation as this bacterium lacks flagella. Furthermore, M. xanthus moves at a speed that is slower than the rate
of diffusion of small molecules. This led some inves-tigators to suggest that chemotaxis in M. xanthus was impossible33. However, Shi et al.34 devised a chemotaxis assay that was based on chambered Petri dishes with steep chemical gradients. They found that M. xanthus cells show directed movement towards complex sub-strates in yeast extract and peptides (but not amino acids) and away from dimethyl sulphoxide (DmSo) and isoamyl alcohol. These movements are completely dependent on an intact Frz system. Indeed, FrzCD was shown to be highly methylated in the presence of attractants (peptides in the rich media) and demethyl-ated in the presence of repellents (DmSo and isoamyl alcohol). This is consistent with the E. coli model, in which methylation/demethylation of chemoreceptors is required for adaptation to stimuli.
Kearns and Shimkets tested the hypothesis that M. xanthus might show chemotaxis to slowly diffusing macromolecules35. For example, they demonstrated that M. xanthus showed chemotactic behaviour in response to phosphatidylethanolamine (Pe) gradients. When M. xanthus cells were placed on a Pe gradient, they moved up the gradient by suppressing cell reversals. After an hour, the cells showed adaptation by returning to the pre-stimulus cell-reversal frequency. Unexpectedly, stimulation was dependent on the Dif (defective in fruit-ing) chemosensory system (see below) rather than on the
Nature Reviews | Microbiology
Time 1
Time 2
Time 3
Box 1 | Two engines power gliding motility in Myxococcus xanthus
The social behaviours of Myxococcus xanthus depend on gliding motility (FIG. 2a), which is movement in the direction of the long axis of the cell at a solid–liquid, or a solid–air, interface without the aid of flagella7. Hodgkin and Kaiser19 used genetic analysis to show that M. xanthus uses two independent systems for gliding; social (S-) motility involves the movement of cells in groups and generally requires cell–cell contact; adventurous (A-) motility involves the movement of individual cells, with or without cell–cell contact.
S-motility S-motility is similar to twitching motility in Pseudomonas aeruginosa and is powered by the extension and retraction of type IV pili46,73. Pili are extended from one end of the cell (the leading cell pole), where they bind to polysaccharides that are present on another cell or on the substrate surface. Retraction of the pili pulls the cell forwards (FIG. 3). When cells reverse, the pili are extended from the opposite cell pole, which now becomes the new leading pole. S-motility requires type IV pili74, extracellular polysaccharide (EPS)75 and lipopolysaccharide O-antigen76.
A-motilityThe engine for A-motility has been difficult to characterize owing to the absence of A-motility-specific appendages on Myxococcus cells. In 1924, Jahn77 proposed that the A-motor was powered by extrusion and hydration of slime. More recently, Wolgemuth et al.78 calculated that a slime-extrusion engine could theoretically produce enough force to move myxobacteria at the observed speed. This model was consistent with the observation that the rates of slime secretion and cell movement were similar and that putative nozzles for slime secretion were observed clustered at the cell poles. However, this model predicts that the A-motility engines should be located primarily at the posterior cell pole, and that slime secretion from this pole would push cells forward. Sliusarenko et al.79 treated A-motile cells with non-lethal concentrations of the antibiotic cephalexin, which caused the cells to become filamentous, as described by Sun and colleagues80. They observed that in these cells the anterior portions of cells moved forward using their A-motors whereas the posterior portions lagged behind, or did not move. When cells reversed, the same pattern was observed for the new leading and lagging poles, so the A-engine must be distributed (but not necessarily uniformly) along the cell body rather than localized at the posterior pole. The observed motility pattern indicated that it is unlikely that slime extrusion mediates propulsion.
An alternative ‘focal adhesions’ model proposed by Mignot et al.81,82 (FIG. 3) is based on the observation that AglZ, an A-motility protein, is localized in clusters that remain fixed relative to the substratum as cells move forward. According to this model (FIG. 3), uncharacterized protein motors attach to bacterial ‘focal adhesion complexes’. The motors are hypothesized to move on helical cytoskeletal filaments, possibly the actin homologue MreB83. Movement would occur if the adhesion complexes pushed against the surface, gaining traction with the aid of extracellular polysaccharide slime. This model is consistent with the observation that A-motility functions efficiently on firm, dry surfaces, such as 1.5% agar84 (which can provide surface adherence and support). Cellular movement in eukaryotic cells involves transient surface adhesions, as demonstrated by adhesion-coupled traction in apicomplexan gliding motility and eukaryotic focal adhesion85,86. So, some eukaryotic motility systems might have their roots in bacterial gliding motility.
Figure 3 | Myxococcus xanthus motility systems. Social motility is mediated by the extension and retraction of type IV pili (black tendrils) at the leading pole of a cell. Adventurous motility involves multiple transient adhesion complexes (coloured ovals on the bottom of cells) that are located throughout the length of a cell82. This model shows cells rotating as the unidentified motors move on a hypothetical cytoskeletal filament. If cells move on a linear filament, they need not rotate.
Extracellular polysaccharides extracellular polysaccharides that stimulate the retraction of the type IV pili of Myxococcus xanthus.
Dif pathwayA Myxococcus xanthus chemosensory pathway that controls the production of extracellular polysaccharide and lipid chemotaxis. This pathway is essential for social motility.
CheYA receiver-domain protein that is phosphorylated by CheA on an Asp residue. Phosphorylated CheY interacts with the output functions of the chemosensory system, which signals stimulation. In Escherichia coli, this results in reversal of the flagellar motor rotation.
CheRA methyltransferase that methylates methyl-accepting chemotaxis protein receptors in response to stimuli. It is involved in adaptation in bacterial chemotaxis.
CheBA methylesterase that removes methyl groups from receptors and is involved in adaptation. In Escherichia coli, CheB is phosphorylated, and thereby activated, by phosphorylated CheA.
Frz system, although adaptation was dependent on the Frz system. movements towards Pe were bona fide chemotaxis because: biased movements were correlated with the sup-pression of reversals to achieve longer runs when exposed to Pe; the responses were to specific Pe molecules; and cells showed adaptation to the lipid attractants.
FrzCD signalling during aggregation. Significant increases in the methylation of FrzCD during fruiting-body formation indicated that FrzCD senses and adapts to attractants that are produced by other cells and that this methylation pattern contributes to aggregation36. Geng et al.37 found that developmental mutants could be divided into two groups on the basis of the amount of FrzCD methylation. mutants that were blocked in late development, or sporulation, had wild-type FrzCD methylation, whereas non-aggregating, or abnormally aggregating, mutants showed aberrant FrzCD meth-ylation. Further research revealed that the abnormally aggregating mutants could be divided into three groups on the basis of their phenotypes. The first group, which included asg, bsg, esg, actA and mrpABC mutants could not aggregate and showed no increase in FrzCD methylation37–40; the second group, which included csgA, fruA and devT mutants, showed a delay in FrzCD methylation and either did not aggregate or showed delayed aggregation41–43; the third group, which only contained a rodK mutant, sporulated without forming fruiting bodies and had increased FrzCD methylation during early development44. Thus, the methylation of FrzCD defines a discrete step in the developmental programme that is controlled by many developmental regulatory genes.
The Dif chemosensory systemThe Dif chemosensory system was discovered in a genetic screen for mutants that could not form fruiting bodies45. The dif mutants that were initially identified could not aggregate during starvation, were defective in S-motility and did not clump together in liquid media. Surprisingly, some dif mutants produced more pili than wild-type cells, although these mutants lacked extracellu-lar polysaccharides (ePS). The hyperpiliated difA-mutant phenotype was complemented by the addition of ePS or chitin to the growth medium46. These studies suggest that ePS triggers pilus retraction, that ePS (and possibly other complex polysaccharides) constitute ‘receptors’ for pilus-mediated motility, and that ePS biosynthesis is regulated by the Dif chemosensory system.
The Dif pathway contains five chemotaxis homo-logues: mCP (DifA), CheW (DifC), CheY (DifD), CheA (Dife) and CheC (DifG), but not CheR and CheB45 (FIG. 5; TABLe 1). DifC interacts with both DifA and Dife, which suggests that these proteins form a ternary signalling complex in M. xanthus47. mutants of difA, difC and difE are defective in S-motility and in development45,46. These defects are most likely due to the lack of ePS production, which requires DifA, DifC and Dife45,48,49. Deletion of either difD or difG results in the overproduction of ePS, which might indicate that these loci negatively regulate the activity of the dif pathway 50,51. The DifG protein is a CheC-like phosphatase and so is predicted to be the main component that mediates adaptation in the Dif system, in lieu of a methylation system.
Pili might function as part of a sensory apparatus for the perception of signals for the Dif pathway, because the presence of pili is required for ePS production52.
Box 2 | Changing direction
Controlling reversal frequencies enables the directed movement of cells towards attractants or away from repellents. Bacteria are often considered too small to detect a concentration gradient that spans the length of a cell. To overcome this problem, bacteria move using a biased ‘random walk’, and methylation or demethylation of receptors is used to ‘remember’ past concentrations of attractants or repellents. Bacteria that are propelled by flagella adjust directional bias of the flagella by controlling their rotation. The enteric bacteria do this by modulating the reversal frequency of peritrichous flagella to produce a ‘run’ when rotating anti-clockwise (because the flagella form a bundle) or a ‘tumble’ when rotating clockwise (because the flagella are unbundled)6. Cells reorient themselves at random during a tumble. In a gradient of attractant, the tumble frequency is decreased, which increases the length of the run. By contrast, in a gradient of repellent, the frequency of tumbling is increased, which reduces movement in that direction.
The best understood paradigm for chemosensory signal transduction is the chemotaxis pathway of Escherichia coli6,87,88. In E. coli, most signals are sensed by receptors ( methyl-accepting chemotaxis proteins (MCPs)). MCPs regulate the activity of the histidine protein kinase CheA through the linker protein CheW. In the absence of an attractant, or in the presence of a repellent, CheA is autophosphorylated and then a phosphoryl group is transferred to the response regulator, CheY; phosphorylated CheY increases the likelihood that cells will change direction by tumbling by stimulating flagellar rotation to change from anti-clockwise to clockwise89. Phosphorylated CheY is dephosphorylated by the phosphatase CheZ. Adaptation to the continued presence of a chemical stimulus is facilitated by the methylation and demethylation of the receptors by the CheR methyltransferase and the CheA-activated CheB methylesterase, respectively6,87,90–92. Rhodobacter sphaeroides controls its directional movements in a similar manner, except that it contains a single flagellum that rotates clockwise to generate runs; the flagellum stops rotating to allow Brownian motion to randomly reorient the cells93.
M. xanthus cells lack flagella and only move on surfaces in two dimensions. To control directional movements, cells periodically reverse their movements: the leading cell pole becomes the new lagging cell pole. In the presence of attractants, reversal frequencies are reduced, whereas in the presence of repellents, reversal frequencies are increased. Interestingly, several proteins, such as FrzS and RomR (L. Søgaard-Anderson, personal communication), change their localization during reversals and are transported from pole to pole as cells change their polarity31,94. Periodic reversals are particularly important for M. xanthus as the cells are remarkably flexible and bend, and can even make U-turns.
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Che3 pathway A Myxococcus xanthus chemosensory pathway that controls the expression of many developmental genes.
Che4 pathwayA Myxococcus xanthus chemosensory pathway that regulates social motility.
As S-motility requires functional pili, ePS and cell prox-imity, a positive regulatory loop could exist, in which ePS triggers pilus retraction, thereby transducing a signal to the Dif pathway to upregulate ePS production52. This model accounts for the observation that cells require the Dif system to maintain ePS-dependent coordinated association and motility.
To probe the function of the DifA chemoreceptor, Xu et al.53 created a chimaera between the n terminus of the E. coli nitrate sensor (narX) and the C terminus of DifA to generate nafA. They found that a difA mutant that expresses the nafA chimaera only produces ePS in the presence of nitrate, which indicates that DifA transduces a signal through its highly conserved C-terminal signal-ling domain to the downstream Dife kinase to regulate ePS production.
Studies by Kearns, Bonner and Shimkets indicated that the Dif system is required for a tactic response to Pe lipids54–56. The most potent chemoattractant Pe spe-
cies were 16:1ω5c (a lipid that is unusual in bacteria, but common in M. xanthus) and 18:1ω9c (which is com-mon in E. coli). Interestingly, not all Dif components are required for sensing both lipids. DifA is required for a response to 16:1ω5c but not to 18:1ω9c, which suggests that DifA is responsible for self-recognition through particular lipids. Furthermore, the Frz pathway, DifB and DifG (the CheC-like phosphatase) are important for adaptation to 16:1ω5c, whereas neither DifB nor DifG is required for adaptation to 18:1ω9c. notably, only Dife (the CheA kinase) and DifD are involved in responses to both lipids. The mechanism by which the Dif system affects ePS and subsequent taxis to Pe remains unknown.
Multiple M. xanthus chemosensory systemsAnalysis of a cosmid library for mCP and CheA homo-logues followed by analysis of the M. xanthus genome upon its completion revealed the presence of six more
Nature Reviews | Microbiology
cheCdif
che3
frz
che5
che6
che4
Che4Che3
che7
che8
cheW mcp cheA cheB cheRRR
//
a
b
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H3C H3C H3C H3CCH3
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CrdBGeneric chemotaxis Dif Frz
G
EEA A4C D
F R3 R4
Z
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Y4
MCP CheWCheA CheY/RRCheB CheRCheC Non Che proteins
CrdC
A3B
Figure 4 | multiple chemosensory gene clusters in Myxococcus xanthus. a | Chemotaxis pathways in Myxococcus xanthus were identified by computer searches (BLAST) for genes that encode CheA homologues and other chemotaxis proteins. The genome also contains 13 orphan chemoreceptors of unknown function that are not shown. b | A schematic diagram of the Dif, Frz, Che3 and Che4 pathways of M. xanthus. It has been proposed that the chemosensory proteins form multiple complexes. For simplicity, we only show a single representative of each protein in each pathway. Che3 contains two unique methyl-accepting chemotaxis proteins. The colours of the proteins match the respective genes in panel a. It should be noted that CheY-like proteins contain a REC domain but might not function as CheY homologues, as only one domain is similar to CheY. Crd, chemosensory regulator of development. REC, CheY-homologous receiver domain; RR, response regulator.
Type IV piliLong flexible appendages that are found at the poles of cells and can power motility.
chemosensory systems9 (FIG. 4; TABLe 1). each system contained one CheA-kinase homologue and several auxiliary chemotaxis homologues, including mCP chemoreceptors, CheW coupling proteins, CheR and CheB adaptation enzymes, and CheY-like (receiver-domain) output proteins. no other bacterial genome sequence completed so far has as many chemosensory systems as M. xanthus (TABLe 2). This discovery raised several interesting questions. Which functions are regu-lated by these multiple chemosensory systems? How are these chemosensory systems functionally coupled or insulated to prevent crosstalk? What specific advantages do chemosensory systems provide over other signal-transduction systems? The answers to some of these questions can be obtained by the analysis of mutants; others remain the subject of speculation.
Che3 controls developmental genes. The Che3 system was the third chemosensory system to be identified in M. xanthus57. Surprisingly, mutants in che3 genes are not defective for either A- or S-motility under labora-tory conditions; however, they are defective for colony morphology and development. For example, on rich media che3 mutants form aggregates, or early fruiting bodies, that contain spores. Upon starvation, aggrega-tion is rapid and premature. Additionally, expression of developmentally regulated genes, such as spi, tps and mbhA, is elevated during vegetative growth and during early stages of development. These results indicated that the Che3 system senses an unknown signal that inhibits developmental gene expression when nutrients
are present; in the absence of this chemosensory system, developmental gene expression is turned on inappropri-ately and at high levels. Interestingly, the output for the Che3 system is a σ54 activator, CrdA (chemosensory regulator of development A), that is divergently tran-scribed from the che3 operon57. CrdA was proposed to regulate developmentally controlled genes and to autoregulate the che3 operon. CrdB (the first open reading frame that is encoded in the che3 operon) is an outer membrane lipoprotein. CrdB has the follow-ing domain organization: an n-terminal lipoprotein anchor, followed by a low similarity domain, followed by a peptidoglycan-binding (ompA-like) domain. It is proposed to function as an input to the pathway, as the crdB mutant has a ‘premature development’ phenotype that is similar to che3 mutants (J.R.K., unpublished observations). CrdB might sense outer membrane integrity or periplasmic signals that trigger the devel-opmental programme. If so, CrdB would be the first lipoprotein to be shown to function as an input to a chemosensory pathway.
Other chemosensory systems. Analysis of the Che4, Che5, Che6, Che7 and Che8 chemosensory systems revealed that these chemosensory modules are versa-tile and complex. For example, in a strain that only has S-motility (A–S+), the deletion of the entire che4 operon (FIG. 4; TABLe 1) causes enhanced vegetative swarming, but prevents aggregation and sporulation58. The che6 operon is needed for type IV pilus assembly. mutants in the Che6 pathway have greatly reduced S-motility but
Table 1 | The chemosensory domains of Myxococcus xanthus chemotaxis proteins and chemotaxis domains of other bacteria
*Proteins that function in chemotaxis that were first described in Escherichia coli, Salmonella typhimurium or Bacillus subtilis . ‡The M. xanthus proteins from each of the eight chemosensory pathways contain domains similar to those found in other bacterial chemotaxis proteins. §The locus identifying each gene in the National Center for Biotechnology Information (NCBI) database is given as the MXAN number in parentheses. MXAN number designations and annotations are based on the M. xanthus sequence completed by The Institute for Genomic Research and the Monsanto Company. ¶CheY-like proteins contain a REC domain (CheY-homologous receiver domain), but might not function as CheY homologues as only one domain is similar to CheY.
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MyxosporeA spherical, environmentally resistant and metabolically inactive Myxococcus xanthus cell.
form premature (early) fruiting bodies (J.R.K., unpub-lished observations). In addition to che homologues, the che6 operon encodes SocD (a suppressor of a mutant that fails to produce the extracellular C-signal that controls aggregation and sporulation59) and KefC, a potassium efflux pump homologue. The Che7 system represents another significant deviation from the E. coli paradigm in terms of function. Although many details are not yet known, it is clear that this system is required for proper regulation of fatty acid composition in both vegetative cells and spores. The che7 gene cluster encodes Che homologues, a homologue of cyanobacterial phyco-cyanobilin lyase (Nostoc sp. PCC 7120) and a fatty acid desaturase. mutant analyses indicate that the Che7 sys-tem is important for resistance to temperature stress and for the production of viable spores (J.R.K., unpublished observations). Functions have not yet been identified for the Che5 and Che8 systems.
How did M. xanthus acquire so many chemosen-sory genes? Goldman et al.9 found that the myxobac-teria contain many more genes than other members of the sequenced deltabacteria and that this most likely arose by gene duplications and divergence. We propose that whereas some che genes arose as a result of duplication events, other clusters could have been acquired horizontally, as suggested by the gene order and the similarity with clusters that are present in other organisms.
Multiple chemosensory systems in other bacteriamultiple chemosensory systems that have been identi-fied in several bacterial species regulate various cellular functions as well as chemotaxis (TABLe 2). Two examples are the Che3 system in Rhodospirillum centenum and the Wsp (wrinkly spreader) system in Pseudomonas aeruginosa. like M. xanthus, R. centenum has a complex life cycle that includes sensing surfaces and forming cysts in response to starvation that are meta-bolically dormant and similar to myxospores60. These behaviours use several chemosensory systems. First, the Che1 pathway controls chemotactic responses to pyruvate and phototactic responses to light. Second, the Che2 system is necessary for assembly of both the polar flagellum and lateral flagella, but not for the expres-sion of the genes that encode the protein components. In this regard the R. centenum Che2 system is simi-lar to the M. xanthus Che6 system, which controls type Iv pilus assembly without affecting gene expres-sion. Third, the Che3 system regulates cyst develop-ment in R. centenum. mutations in che3 genes lead to premature formation of cysts, even in the presence of nutrients60.
The Wsp chemosensory system in P. aeruginosa is another interesting example of a non-traditional chemo-sensory system: it regulates cyclic-di-GmP production and, subsequently, biofilm formation61. Cyclic-di-GmP is produced by the GGDeF domain of the WspR response
GTP GTP
Nature Reviews | Microbiology
P
P
P
Signal
Signal
FrzF FrzCD
CH3
H3C
FrzA
FrzE
FrzZ
GTP
GDPMgIA
FrzS
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AgIZ
MgIB
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a Input mode b Output mode
Figure 5 | model for the regulation of directional control in Myxococcus xanthus. a | The Frz chemosensory system is shown as the input module for receiving and processing signalling information. The Frz system is a two-component signal-transduction pathway that consists of a cytoplasmic methyl-accepting chemotaxis protein, FrzCD, a methyltransferase, FrzF, a CheW adaptor protein, FrzA, a CheA–response regulator (RR) hybrid protein, FrzE, and dual RR receiver protein, FrzZ. In the model, reversals are inhibited by one or more unknown signals (attractants) that interact with FrzCD and/or FrzF and cause site-specific methylation of FrzCD. FrzG is not shown because its regulatory role remains unclear. In the absence of attractants, or in the presence of repellents, FrzCD induces autophosphorylation of FrzE, which is negatively regulated by the C-terminal domain of FrzE. FrzZ accepts the phosphoryl group from FrzE and propagates signalling to the output module30. b | The output module consists of complexes of protein that trigger reversals of both the adventurous (A-) and social (S-) motility systems. Phosphorylated FrzZ is proposed to mediate an unknown interaction with components of the MglA complex, which independently and coordinately signals the S-motility system and the A-motility system to trigger a reversal and define a new leading pole. MglA interacts with FrzS and AglZ, components of the S- and A-motility systems, respectively.
regulator protein. Thus, the Wsp chemosensory sys-tem directly regulates this alternative output. The Wsp system is one of four chemosensory systems present in P. aeruginosa: genes in cluster 1 regulate flagellar-based motility and genes in the pil–chp cluster regulate type-Iv-pilus-based motility62. The mechanism by which the other Che systems affect pilus assembly remains unknown. Interestingly, these genes are also implicated in Pseudomonas pathogenesis in a Drosophila model sys-tem63, which suggests a role for the pil–chp chemosensory system in other complex regulatory networks.
Why have bacteria evolved multiple chemosensory systems? The key advantage provided by chemosen-sory systems is temporal regulation of a given output in response to a persistent stimulus. Because Che systems display adaptation via modification of the chemore-ceptor proteins, a stimulus can elicit a response that is subsequently attenuated even though the stimulus remains. For M. xanthus, high cell density is required for successful fruiting-body development. High cell density leads to expression of some genes that are only required at early stages of development (for example, spi 64), and these are subsequently downregulated during spore maturation even though high cell density persists. Thus, crucial functions that require temporal regulation might be regulated best by chemosensory systems.
not all bacterial models for development use chem-osensory two-component systems to regulate their life cycles. Bacillus subtilis, like E. coli, has only one CheA that interacts with multiple receptors. However, there is no evidence to implicate these Che proteins in sporulation. Furthermore, S. coelicolor, another spore-producing bacterium that is used as a model for development, has no chemotaxis homologues or mCPs. Caulobacter crescentus has two che gene clusters and four mCPs that regulate motility during planktonic growth65. Thus, the expansion of chemosensory systems by bacteria
to regulate development might be limited to just a few microorganisms.
Receptor signalling arrays in MyxococcusE. coli has five mCP homologues, but only one CheW coupling protein and one CheA kinase. The mCPs are produced in varying amounts and are known to form functional arrays at one cell pole that are composed of approximately 10,000 total chemoreceptors66–68. As a result, any information sensed by the mCPs is transferred to CheW and CheA proteins, which together with the mCPs form a ‘signalling patch’ in the cytoplasm. According to the latest model, when one receptor molecule binds to the appropriate ligand, the ligand-bound receptor complex can influence the conformation of neighbouring receptors, which affects the associated CheA kinases, generating ‘gain’ in the system. The current model accounts for the observa-tion that only one ligand-bound receptor is needed to elicit a detectable behavioural response by the cell69.
The receptor-signalling-array model poses a major problem in organisms that use multiple CheW and CheA homologues, such as M. xanthus. Do arrays follow the model for the E. coli paradigm? If arrays form, how is crosstalk between the different chemo-sensory pathways prevented? Do these arrays have molecular insulation (to prevent crosstalk) or some level of cross-regulation? on the basis of the analyses of several genomes that contain multiple chemosensory systems, it seems likely that the integrity of the multiple chemosensory systems is maintained by encoding spe-cific mCP–CheW–CheA sets of homologues, typically present in an operon (FIG. 4a).
In M. xanthus, many of the Che systems function independently to control specific outputs, as described above. no cytological evidence exists for a single large signalling array such as that observed in E. coli. (D.R.Z.,
Table 2 | Comparison of chemosensory systems from selected bacteria and archaea
*Known bacterial and archaeal chemosensory pathways contain multiple Che proteins, one of which must be CheA. MCPs; methyl-accepting chemotaxis proteins; TFP, type IV pili.
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unpublished observations). The M. xanthus genome contains 13 orphan chemoreceptor genes (not present in che clusters) that are predicted to interact with CheW homologues encoded in che gene clusters (J.R.K., unpub-lished observations). Phenotypic analysis supports this argument and indicates that multiple receptors funnel information to each CheA protein to form a defined signalling pathway (J.R.K., unpublished observations).
A chemosensory pathway present in Rhodobacter sphaeroides suggests one mechanism that might be used to prevent crosstalk between chemoreceptors. Genetic and biochemical analysis of the che gene clus-ters and mCP homologues in R. sphaeroides indicates that specific mCP genes are regulated and the resulting mCP receptors are differentially localized in the cell70. Temporal regulation of che gene expression during development has been demonstrated in M. xanthus71. The che3 gene cluster is upregulated approximately 6–10 fold during development72. Thus, the putative signalling array must change substantially, at least with respect to the number of Che3 homologues that it contains, as the cells transition from vegeta-tive growth into development. Furthermore, evidence
indicates that the cytoplasmic mCP, FrzCD, is localized in multiple discrete clusters throughout the cell and that this might affect the type of signals that are perceived and the subsequent signal transduction (e. mauriello, D. Astling and D.R.Z., unpublished observations). Thus, the sequestering of the different Che systems in the cell might prevent unwanted crosstalk between chemosensory systems with different functions.
OutlookCan we predict the roles (with regards to behaviour, tran-scription and physiology) of uncharacterized chemosen-sory systems? Genes that are transcribed together in an operon are likely to encode proteins that function together. Because genes that have no homology to che genes are sometimes present in che gene clusters, these genes might provide clues to the function of the Che system. However, whereas predictions are easily made, the biochemical con-nections and overall mechanism of regulation is not dis-cernable without significant effort. Adopting M. xanthus as a model provides a unique opportunity for the analy-sis of the role of chemosensory signal transduction in bacterial development and motility.
1. Kaiser, D. Signaling in myxobacteria. Annu. Rev. Microbiol. 58, 75–98 (2004).
2. Kaiser, D. A microbial genetic journey. Annu. Rev. Microbiol. 60, 1–25 (2006).
3. Kaiser, D. Coupling cell movement to multicellular development in myxobacteria. Nature Rev. Microbiol. 1, 45–54 (2003).
4. Shimkets, L. J. Intercellular signaling during fruiting-body development of Myxococcus xanthus. Annu. Rev. Microbiol. 53, 525–549 (1999).
5. Reichenbach, H. The ecology of the myxobacteria. Environmen. Microbiol. 1, 15–21 (1999).
6. Baker, M. D., Wolanin, P. M. & Stock, J. B. Signal transduction in bacterial chemotaxis. Bioessays 28, 9–22 (2006).
7. McBride, M. J. Bacterial gliding motility: multiple mechanisms for cell movement over surfaces. Annu. Rev. Microbiol. 55, 49–75 (2001).
8. Rosenberg, E., Keller, K. H. & Dworkin, M. Cell density-dependent growth of Myxococcus xanthus on casein. J. Bacteriol. 129, 770–777 (1977).
9. Goldman, B. S. et al. Evolution of sensory complexity recorded in a myxobacterial genome. Proc. Natl Acad. Sci. USA 103, 15200–15205 (2006).
10. McBride, M. J. & Zusman, D. R. Behavioral analysis of single cells of Myxococcus xanthus in response to prey cells of Escherichia coli. FEMS Microbiol. Lett. 137, 227–231 (1996).
11. Berleman, J. E., Chumley, T., Cheung, P. & Kirby, J. R. Rippling is a predatory behavior in Myxococcus xanthus. J. Bacteriol. 188, 5888–5895 (2006).
12. Igoshin, O. A., Mogilner, A., Welch, R. D., Kaiser, D. & Oster, G. Pattern formation and traveling waves in myxobacteria: theory and modeling. Proc. Natl Acad. Sci. USA 98, 14913–14918 (2001).
13. Sager, B. & Kaiser, D. Intercellular C-signaling and the traveling waves of Myxococcus. Genes Dev. 8, 2793–2804 (1994).
14. Sliusarenko, O., Neu, J., Zusman, D. R. & Oster, G. Accordion waves in Myxococcus xanthus. Proc. Natl Acad. Sci. USA 103, 1534–1539 (2006).
15. O’Connor, K. A. & Zusman, D. R. Behavior of peripheral rods and their role in the life cycle of Myxococcus xanthus. J. Bacteriol. 173, 3342–3355 (1991).
16. Wireman, J. W. & Dworkin, M. Developmentally induced autolysis during fruiting body formation by Myxococcus xanthus. J. Bacteriol. 129, 798–802 (1977).
17. Berleman, J. E. & Kirby, J. R. multicellular development in Myxococcus xanthus is stimulated by predator–prey interactions. J. Bacteriol. 189, 5675–5682 (2007).
18. Hodgkin, J. & Kaiser, D. Cell-to-cell stimulation of movement in nonmotile mutants of Myxococcus. Proc. Natl Acad. Sci. USA 74, 2938–2942 (1977).
19. Hodgkin, J. & Kaiser, D. Genetics of gliding motility in Myxococcus xanthus (myxobacteriales) — two gene systems control movement. Mol. Gen. Genet. 171, 177–191 (1979). This is a seminal paper that showed that there are two motility systems in M. xanthus.
20. Stephens, K., Hartzell, P. & Kaiser, D. Gliding motility in Myxococcus xanthus: mgl locus, RNA, and predicted protein products. J. Bacteriol. 171, 819–830 (1989).
21. Hartzell, P. L. Complementation of sporulation and motility defects in a prokaryote by a eukaryotic GTPase. Proc. Natl Acad. Sci. USA 94, 9881–9886 (1997).
22. Barnes, G., Louie, K. A. & Botstein, D. Yeast proteins associated with microtubules in vitro and in vivo. Mol. Biol. Cell 3, 29–47 (1992).
23. Yang, R. et al. AglZ is a filament-forming coiled-coil protein required for adventurous gliding motility of Myxococcus xanthus. J. Bacteriol. 186, 6168–6178 (2004).
24. Ward, M. J., Lew, H. & Zusman, D. R. Social motility in Myxococcus xanthus requires FrzS, a protein with an extensive coiled-coil domain. Mol. Microbiol. 37, 1357–1371 (2000).
25. Lancero, H. et al. Characterization of a Myxococcus xanthus mutant that is defective for adventurous motility and social motility. Microbiology 150, 4085–4093 (2004).
26. Zusman, D. R. “Frizzy” mutants: a new class of aggregation-defective developmental mutants of Myxococcus xanthus. J. Bacteriol. 150, 1430–1437 (1982).
27. Blackhart, B. D. & Zusman, D. R. “Frizzy” genes of Myxococcus xanthus are involved in control of frequency of reversal of gliding motility. Proc. Natl Acad. Sci. USA 82, 8767–8770 (1985).This paper showed that the Frz system controls the frequency of cell reversals.
28. McBride, M. J., Weinberg, R. A. & Zusman, D. R. “Frizzy” aggregation genes of the gliding bacterium Myxococcus xanthus show sequence similarities to the chemotaxis genes of enteric bacteria. Proc. Natl Acad. Sci. USA 86, 424–428 (1989). This paper showed that the frz genes encode chemosensory homologues.
29. Ward, M. J. & Zusman, D. R. Motility in Myxococcus xanthus and its role in developmental aggregation. Curr. Opin. Microbiol. 2, 624–629 (1999).
30. Inclan, Y. F., Vlamakis, H. C. & Zusman, D. R. FrzZ, a dual CheY-like response regulator, functions as an output for the Frz chemosensory pathway of Myxococcus xanthus. Mol. Microbiol. 65, 90–102 (2007).
31. Fraser, J. S. et al. An atypical receiver domain controls the dynamic polar localization of the Myxococcus xanthus social motility protein FrzS. Mol. Microbiol. 65, 319–332 (2007).
32. Mignot, T., Merlie, J. P. Jr & Zusman, D. R. Two localization motifs mediate polar residence of FrzS during cell movement and reversals of Myxococcus xanthus. Mol. Microbiol. 65, 363–372 (2007).
33. Dworkin, M. & Eide, D. Myxococcus xanthus does not respond chemotactically to moderate concentration gradients. J. Bacteriol. 154, 437–442 (1983).
34. Shi, W., Kohler, T. & Zusman, D. R. Chemotaxis plays a role in the social behaviour of Myxococcus xanthus. Mol. Microbiol. 9, 601–611 (1993).
35. Kearns, D. B. & Shimkets, L. J. Lipid chemotaxis and signal transduction in Myxococcus xanthus. Trends Microbiol. 9, 126–129 (2001). This paper describes lipid chemotaxis in M. xanthus.
36. McBride, M. J. & Zusman, D. R. FrzCD, a methyl-accepting taxis protein from Myxococcus xanthus, shows modulated methylation during fruiting body formation. J. Bacteriol. 175, 4936–4940 (1993).
37. Geng, Y., Yang, Z., Downard, J., Zusman, D. & Shi, W. Methylation of FrzCD defines a discrete step in the developmental program of Myxococcus xanthus. J. Bacteriol. 180, 5765–5768 (1998).
38. Gorski, L., Gronewold, T. & Kaiser, D. A σ54 activator protein necessary for spore differentiation within the fruiting body of Myxococcus xanthus. J. Bacteriol. 182, 2438–2444 (2000).
39. Sun, H. & Shi, W. Genetic studies of mrp, a locus essential for cellular aggregation and sporulation of Myxococcus xanthus. J. Bacteriol. 183, 4786–4795 (2001).
40. Sun, H. & Shi, W. Analyses of mrp genes during Myxococcus xanthus development. J. Bacteriol. 183, 6733–6739 (2001).
41. Boysen, A., Ellehauge, E., Julien, B. & Sogaard-Andersen, L. The DevT protein stimulates synthesis of FruA, a signal transduction protein required for fruiting body morphogenesis in Myxococcus xanthus. J. Bacteriol. 184, 1540–1546 (2002).
42. Ellehauge, E., Norregaard-Madsen, M. & Sogaard-Andersen, L. The FruA signal transduction protein provides a checkpoint for the temporal co-ordination of intercellular signals in Myxococcus xanthus development. Mol. Microbiol. 30, 807–817 (1998).
43. Sogaard-Andersen, L. et al. Coupling gene expression and multicellular morphogenesis during fruiting body formation in Myxococcus xanthus. Mol. Microbiol. 48, 1–8 (2003).
44. Rasmussen, A. A., Porter, S. L., Armitage, J. P. & Sogaard-Andersen, L. Coupling of multicellular morphogenesis and cellular differentiation by an unusual hybrid histidine protein kinase in Myxococcus xanthus. Mol. Microbiol. 56, 1358–1372 (2005).
45. Yang, Z., Geng, Y., Xu, D., Kaplan, H. B. & Shi, W. A new set of chemotaxis homologues is essential for Myxococcus xanthus social motility. Mol. Microbiol. 30, 1123–1130 (1998).This was the first paper to describe the Dif chemosensory system.
46. Li, Y. et al. Extracellular polysaccharides mediate pilus retraction during social motility of Myxococcus xanthus. Proc. Natl Acad. Sci. USA 100, 5443–5448 (2003).
47. Yang, Z. & Li, Z. Demonstration of interactions among Myxococcus xanthus Dif chemotaxis-like proteins by the yeast two-hybrid system. Arch. Microbiol. 183, 243–252 (2005).
48. Bellenger, K., Ma, X., Shi, W. & Yang, Z. A CheW homologue is required for Myxococcus xanthus fruiting body development, social gliding motility, and fibril biogenesis. J. Bacteriol. 184, 5654–5660 (2002).
49. Yang, Z. et al. Myxococcus xanthus dif genes are required for biogenesis of cell surface fibrils essential for social gliding motility. J. Bacteriol. 182, 5793–5798 (2000).
50. Black, W. P. & Yang, Z. Myxococcus xanthus chemotaxis homologs DifD and DifG negatively regulate fibril polysaccharide production. J. Bacteriol. 186, 1001–1008 (2004).
51. Bonner, P. J. et al. The Dif chemosensory pathway is directly involved in phosphatidylethanolamine sensory transduction in Myxococcus xanthus. Mol. Microbiol. 57, 1499–1508 (2005).
52. Black, W. P., Xu, Q. & Yang, Z. Type IV pili function upstream of the Dif chemotaxis pathway in Myxococcus xanthus EPS regulation. Mol. Microbiol. 61, 447–456 (2006).
53. Xu, Q., Black, W. P., Ward, S. M. & Yang, Z. Nitrate-dependent activation of the Dif signaling pathway of Myxococcus xanthus mediated by a NarX–DifA interspecies chimera. J. Bacteriol. 187, 6410–6418 (2005).
54. Bonner, P. J. & Shimkets, L. J. Phospholipid directed motility of surface-motile bacteria. Mol. Microbiol. 61, 1101–1109 (2006).
55. Kearns, D. B., Bonner, P. J., Smith, D. R. & Shimkets, L. J. An extracellular matrix-associated zinc metalloprotease is required for dilauroyl phosphatidylethanolamine chemotactic excitation in Myxococcus xanthus. J. Bacteriol. 184, 1678–1684 (2002). This paper showed that the Che3 system controls gene expression during development.
56. Kearns, D. B., Campbell, B. D. & Shimkets, L. J. Myxococcus xanthus fibril appendages are essential for excitation by a phospholipid attractant. Proc. Natl Acad. Sci. USA 97, 11505–11510 (2000).
57. Kirby, J. R. & Zusman, D. R. Chemosensory regulation of developmental gene expression in Myxococcus xanthus. Proc. Natl Acad. Sci. USA 100, 2008–2013 (2003).
58. Vlamakis, H. C., Kirby, J. R. & Zusman, D. R. The Che4 pathway of Myxococcus xanthus regulates type IV pilus-mediated motility. Mol. Microbiol. 52, 1799–1811 (2004).
59. Lee, K. & Shimkets, L. J. Suppression of a signaling defect during Myxococcus xanthus development. J. Bacteriol. 178, 977–984 (1996).
60. Berleman, J. E. & Bauer, C. E. A Che-like signal transduction cascade involved in controlling flagella biosynthesis in Rhodospirillum centenum. Mol. Microbiol. 55, 1390–1402 (2005).
61. Hickman, J. W., Tifrea, D. F. & Harwood, C. S. A chemosensory system that regulates biofilm formation through modulation of cyclic diguanylate levels. Proc. Natl Acad. Sci. USA 102, 14422–14427 (2005).
62. Whitchurch, C. B. et al. Characterization of a complex chemosensory signal transduction system which controls twitching motility in Pseudomonas aeruginosa. Mol. Microbiol. 52, 873–893 (2004).
63. D’Argenio, D. A., Gallagher, L. A., Berg, C. A. & Manoil, C. Drosophila as a model host for Pseudomonas aeruginosa infection. J. Bacteriol. 183, 1466–1471 (2001).
64. Guo D, W. Y., Kaplan HB. Identification and characterization of genes required for early Myxococcus xanthus developmental gene expression. J. Bacteriol. 182, 4564–4571 (2000).
65. Alley, M. R., Gomes, S. L., Alexander, W. & Shapiro, L. Genetic analysis of a temporally transcribed chemotaxis gene cluster in Caulobacter crescentus. Genetics 129, 333–341 (1991).
66. Maddock, J. R. & Shapiro, L. Polar location of the chemoreceptor complex in the Escherichia coli cell. Science 259, 1717–1723 (1993).
67. Bray, D., Levin, M. D. & Morton-Firth, C. J. Receptor clustering as a cellular mechanism to control sensitivity. Nature 393, 85–88 (1998).
68. Parkinson, J. S., Ames, P. & Studdert, C. A. Collaborative signaling by bacterial chemoreceptors. Curr. Opin. Microbiol. 8, 116–121 (2005).
69. Segall, J. E., Block, S. M. & Berg, H. C. Temporal comparisons in bacterial chemotaxis. Proc. Natl Acad. Sci. USA 83, 8987–8991 (1986).
70. Wadhams, G. H. et al. TlpC, a novel chemotaxis protein in Rhodobacter sphaeroides, localizes to a discrete region in the cytoplasm. Mol. Microbiol. 46, 1211–1221 (2002).
71. Weinberg, R. A. & Zusman, D. R. Evidence that the Myxococcus xanthus frz genes are developmentally regulated. J. Bacteriol. 171, 6174–6186 (1989).
72. Jakobsen, J. S. et al. σ54 enhancer binding proteins and Myxococcus xanthus fruiting body development. J. Bacteriol. 186, 4361–4368 (2004).
73. Sun, H., Zusman, D. R. & Shi, W. Type IV pilus of Myxococcus xanthus is a motility apparatus controlled by the frz chemosensory system. Curr. Biol. 10, 1143–1146 (2000).
74. Wall, D. & Kaiser, D. Type IV pili and cell motility. Mol. Microbiol. 32, 1–10 (1999).
75. Arnold, J. W. & Shimkets, L. J. Inhibition of cell–cell interactions in Myxococcus xanthus by congo red. J. Bacteriol. 170, 5765–5770 (1988).
76. Bowden, M. G. & Kaplan, H. B. The Myxococcus xanthus lipopolysaccharide O-antigen is required for social motility and multicellular development. Mol. Microbiol. 30, 275–284 (1998).
77. Jahn, E. Beitrage zur Botanischen Protistologie (Gebruder Borntraeger, Leipzig, 1924).
78. Wolgemuth, C., Hoiczyk, E., Kaiser, D. & Oster, G. How myxobacteria glide. Curr. Biol. 12, 369–377 (2002).
79. Sliusarenko, O., Zusman, D. R. & Oster, G. The motors powering A-motility in Myxococcus xanthus are distributed along the cell body. J. Bacteriol. 17 Aug 2007 (doi: 10.1128/JB.00923-07).
80. Sun, H., Yang, Z. & Shi, W. Effect of cellular filamentation on adventurous and social gliding motility of Myxococcus xanthus. Proc. Natl Acad. Sci. USA 96, 15178–15183 (1999).
81. Mignot, T. The elusive engine in Myxococcus xanthus gliding motility. Cell. Mol. Life Sci. 25 July 2007 (doi: 10.1007/s00018-007-7176-x).
82. Mignot, T., Shaevitz, J. W., Hartzell, P. L. & Zusman, D. R. Evidence that focal adhesion complexes power bacterial gliding motility. Science 315, 853–856 (2007).
83. Graumann, P. Cytoskeletal elements in bacteria. Annu. Rev. Microbiol. (in the press).
84. Shi, W. & Zusman, D. R. The two motility systems of Myxococcus xanthus show different selective advantages on various surfaces. Proc. Natl Acad. Sci. USA 90, 3378–3382 (1993).
85. Wozniak, M. A., Modzelewska, K., Kwong, L. & Keely, P. J. Focal adhesion regulation of cell behavior. Biochim. Biophys. Acta 1692, 103–119 (2004).
86. Baum, J., Papenfuss, A. T., Baum, B., Speed, T. P. & Cowman, A. F. Regulation of apicomplexan actin-based motility. Nature Rev. Microbiol. 4, 621–628 (2006).
87. Wadhams, G. H. & Armitage, J. P. Making sense of it all: bacterial chemotaxis. Nature Rev. Mol. Cell Biol. 5, 1024–1037 (2004).
88. Wolanin, P. M. et al. Self-assembly of receptor/signaling complexes in bacterial chemotaxis. Proc. Natl Acad. Sci. USA 103, 14313–14318 (2006).
89. Welch, M., Oosawa, K., Aizawa, S. & Eisenbach, M. Phosphorylation-dependent binding of a signal molecule to the flagellar switch of bacteria. Proc. Natl Acad. Sci. USA 90, 8787–8791 (1993).
90. Okumura, H., Nishiyama, S., Sasaki, A., Homma, M. & Kawagishi, I. Chemotactic adaptation is altered by changes in the carboxy-terminal sequence conserved among the major methyl-accepting chemoreceptors. J. Bacteriol. 180, 1862–1868 (1998).
91. Ninfa, E. G., Stock, A., Mowbray, S. & Stock, J. Reconstitution of the bacterial chemotaxis signal transduction system from purified components. J. Biol. Chem. 266, 9764–9770 (1991).
92. Levit, M. N. & Stock, J. B. Receptor methylation controls the magnitude of stimulus-response coupling in bacterial chemotaxis. J. Biol. Chem. 277, 36760–36765 (2002).
93. Armitage, J. P. & Schmitt, R. Bacterial chemotaxis: Rhodobacter sphaeroides and Sinorhizobium meliloti — variations on a theme? Microbiology 143, 3671–3682 (1997).
94. Mignot, T., Merlie, J. P. Jr & Zusman, D. R. Two localization motifs mediate polar residence of FrzS during cell movement and reversals of Myxococcus xanthus. Mol. Microbiol. 65, 363–372 (2007).
95. Gross, L. Antisocial behavior in cooperative bacteria. PLoS Biol.3, 1847–1848 (2005).
96. Foster, K. R. Sociobiology: the Phoenix effect. Nature 441, 291–292 (2006).
97. Zusman, D. R. & O’Connor, K. A. Development in Myxococcus xanthus involves aggregation and sporulation as well as non-aggregation and non-sporulation. Sem. Dev. Biol. 2, 37–45 (1991).
98. Zhulin, I. B. The superfamily of chemotaxis transducers: from physiology to genomics and back. Adv. Microb. Physiol. 45, 157–198 (2001).
99. Alam, M. & Oesterhelt, D. Morphology, function and isolation of halobacterial flagella. J. Mol. Biol. 176, 459–475 (1984).
100. Meier, V. M., Muschler, P. & Scharf, B. E. Functional analysis of nine putative chemoreceptor proteins in Sinorhizobium meliloti. J. Bacteriol. 189, 1816–1826 (2007).
101. Ng, W. V. et al. Genome sequence of Halobacterium species NRC-1. Proc. Natl Acad. Sci. USA 97, 12176–12181 (2000).
102. Pittman, M. S., Goodwin, M. & Kelly, D. J. Chemotaxis in the human gastric pathogen Helicobacter pylori: different roles for CheW and the three CheV paralogues, and evidence for CheV2 phosphorylation. Microbiology 147, 2493–2504 (2001).
103. Croxen, M. A., Sisson, G., Melano, R. & Hoffman, P. S. The Helicobacter pylori chemotaxis receptor TlpB (HP0103) is required for pH taxis and for colonization of the gastric mucosa. J. Bacteriol. 188, 2656–2665 (2006).
104. Foynes, S. et al. Helicobacter pylori possesses two CheY response regulators and a histidine kinase sensor, CheA, which are essential for chemotaxis and colonization of the gastric mucosa. Infect. Immun. 68, 2016–2023 (2000).
105. Kato, J., Nakamura, T., Kuroda, A. & Ohtake, H. Cloning and characterization of chemotaxis genes in Pseudomonas aeruginosa. Biosci. Biotechnol. Biochem. 63, 155–161 (1999).
106. Jiang, Z. Y. & Bauer, C. E. Analysis of a chemotaxis operon from Rhodospirillum centenum. J. Bacteriol. 179, 5712–5719 (1997).
107. Jiang, Z. Y., Gest, H. & Bauer, C. E. Chemosensory and photosensory perception in purple photosynthetic bacteria utilize common signal transduction components. J. Bacteriol. 179, 5720–5727 (1997).
108. Attmannspacher, U., Scharf, B. & Schmitt, R. Control of speed modulation (chemokinesis) in the unidirectional rotary motor of Sinorhizobium meliloti. Mol. Microbiol. 56, 708–718 (2005).
109. Bhaya, D. Light matters: phototaxis and signal transduction in unicellular cyanobacteria. Mol. Microbiol. 53, 745–754 (2004).
AcknowledgementsWe are grateful to members of the Zusman laboratory, past and present, for many helpful discussions. Our research is supported by grants from the National Institutes of Health GM20509 and GM64463 to D.R.Z., GM071601 to Z.Y. and AI59682 to J.R.K. A.E.S was supported by a predoctoral fel-lowship from the National Science Foundation.
FURTHER INFORMATIONDavid R. Zusman’s homepage: http://mcb.berkeley.edu/faculty/BMB/zusmand.htmlJohn R. Kirby’s homepage: http://www.medicine.uiowa.edu/microbiology/faculty/kirby.htmMyxopedia: http://xanthusbase.org
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