Mutations of the Act Promoter in Myxococcus xanthus

Mutations within the –12 and –24 elements provideevidence that the act promoter is recognized by sigma-54 RNApolymerase. Deletion of the –20 base pair, which liesbetween the two conserved elements of sigma-54 promoters, decreasedexpression by 90%. In addition, mutation of a potential enhancersequence, around –120, led to an 80% reduction in actgene expression. actB, the second gene in the act operon, encodesa sigma-54 activator protein that is proposed to be an enhancer-bindingprotein for the act operon. All act genes, actA to actE, areexpressed together and constitute an operon, because an in-framedeletion of actB decreased expression of actA and actE to thesame extent. After an initially slow phase of act operon expression,which depends on FruA, there is a rapid phase. The rapid phaseis shown to be due to the activation of the operon expressionby ActB, which completes a positive feedback loop. That loopappears to be nested within a larger positive loop in whichActB is activated by the C signal via ActA, and the act operonactivates transcription of the csgA gene. We propose that, ascells engage in more C signaling, positive feedback raises thenumber of C-signal molecules per cell and drives the processof fruiting body development forward.

INTRODUCTION

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Introduction

Sensing starvation, myxobacteria stop growing and start buildingfruiting bodies that fill with many thousands of spores (7).The cell-surface-associated, nondiffusible C signal is essentialfor both fruiting body formation and sporulation (39, 56). Csignal is a morphogen, encoded by the csgA gene (38), translatedas a 25-kDa protein, transported to the cell surface, and finallycleaved to the active 17-kDa signal molecule (40). The signalis transmitted from one cell to another by direct end-to-endcontact between them (32-34, 52), which links signaling to thearrangement of cells. Mathematical simulation of C signalingin a population of moving cells has shown that C signal controlsthe shape of the Myxococcus xanthus fruiting body (57, 58).The particular shape of a myxobacterial fruiting body is a speciescharacteristic and is inherited (49, 60).

Cell behavior changes as fruiting bodies form due to changesin the level of C signal, and successive stages of fruitingbody development arise from increasing levels of C signal (31,39). The low initial level of C signal induces traveling wavebehavior (20, 37, 55, 70), a higher level produces streamingbehavior, and the highest level induces spores to form. Cellsin a stream are arranged end to end, all following the sametrajectory, which leads to frequent end-to-end contacts (22-24).Streaming cells enlarge tiny initial aggregates by forming anonion-like succession of spherical shells around the aggregate.The cells within each shell stream in roughly circular orbits.This leads to a spherical outer domain of densely packed cellssurrounding an inner domain of threefold-lower density, whichis observed microscopically (28, 53). The level of C signalalso regulates the expression of many genes involved in fruitingbody development (35), in a timely way. Sporulation genes havea higher threshold of C-signal intensity than genes for aggregation(31, 39), and the highest level of C signal induces the cellsto differentiate into spores within the fruiting body (37).This property confines sporulation to the spherical mass ofa Myxococcus fruiting body, while cells around the outside withlower C-signal levels do not sporulate (25, 46).

Thus, cell movement and gene expression are coordinated forM. xanthus fruiting body development by the rising number ofC-signal molecules per cell (27). Transcription of csgA is initiatedby starvation and the stringent response (5). Later csgA expressionis enhanced by the act operon (17). Mutations in act were foundto affect the expression of developmental gene reporters thatare expressed after 6 h of development and are related to aggregationand sporulation (16). Moreover, deletion of the actC gene ledto the premature formation of many small fruiting bodies (17)—aneffect similar to a 10-fold overexpression of CsgA (37). Bycontrast, deletion of the actD gene led to delayed formationof large fruiting bodies. The kinetics of C-signal increasethus appears to control fruiting body size. After initiationof C-signal expression by starvation, how do the Act proteinsregulate the rate and magnitude of the coordinating increasein the signal level?

MATERIALS AND METHODS

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Materials and Methods

Cultures. Bacterial strains and plasmids are listed in Table 1. The conditionsfor growth; development, including sporulation; and electroporationhave previously been described (17). Transduction was carriedout as described previously (16).

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TABLE 1. Strains and plasmids

Preparation of RNA from developing cells. A culture of 1 x 10¹⁰ to 2 x 10¹⁰ cells was induced to developat 32°C on plates consisting of TPM buffer (10 mM Tris-HCl,pH 7.6, 1 mM KPO₄, pH 7.6, 8 mM MgSO₄) solidified with 1.5%Bacto Agar. Bacteria, or spores, were harvested at the specifiedtime from the surface by scraping, suspension, and washing inTPM buffer. The solid material from harvest was resuspendedin lysozyme-containing buffer (10 mM Tris-HCl, pH 8.0, 1 mMEDTA, 2 mg/ml lysozyme) and then disrupted with a Branson 450Tip sonifier, to ensure that both cells and prespores wouldbe lysed and homogenized. RNA was isolated from the homogenateusing RNeasy Midi Spin columns (QIAGEN).

Quantifying mRNA. To prepare slot blots, 25 µl of each developmental RNAsample dissolved in water was diluted by adding 25 µl2 mM EDTA, 30 µl 2x SSC (1x SSC is 0.15 M NaCl plus 0.015M sodium citrate) (54), and 20 µl 37% formaldehyde. Theentire 100-µl volume of the mixture was then heated at60°C for 15 min and transferred to a Hybond-N nylon membrane(Amersham) with 10x SSC, according to the protocol for Bio-DotSF (Bio-Rad). Finally the membranes were irradiated with UVlight to cross-link the RNA to them. DNA probes for actA, actB,and actE were labeled with ³²P (17). Probes were annealed ontothe RNA-loaded membranes, as described for Southern blotting(54).

Site-directed mutagenesis. A two-step method of overlap extension PCR was employed, whoseprinciples have been described previously (19). Two universalprimers designated upstream XhoI and downstream BamHI were designed.Each universal primer had the indicated synthetic restrictionsite, following a terminal AAAG spacer. The upstream XhoI primersequence is 5' AAAGCTCGAGGCGGCTCGTCGGACACCTTC 3'. The downstreamBamHI primer is 5' AAAGGGATCCCACCGCCTCTTCGTCATCCAC 3', withthe act operon sequences underlined in both sequences. Fromthese two primers, the act promoter region starting at nucleotide(nt) 331 in the upstream region of the GenBank sequence AF350253and running to nucleotide 2501 of that sequence near the beginningof actB was amplified by PCR. The XhoI and BamHI restrictionsites oriented the PCR products for cloning into pBluescriptor pREG1727.

To introduce particular deletions or point mutations at predeterminedsites between nucleotides 331 and 2501, a set of mutagenic primers,shown in Table S2 in the supplemental material, was designedfor both template strands. Each primer, located at the centerof the cloned promoter region, contained 10 nucleotides of wild-typeact sequence, followed by the desired mutant sequence, followedby another 10 nucleotides of the wild-type sequence. The firststep of mutation induction involved two sets of 20 to 25 cyclesof PCRs with one universal primer (either the upstream XhoIprimer or the downstream BamHI primer) and one of each of themutagenic primers. The mutagenic primer design ensures thatpairs of first-step PCR products overlapped by at least 20 bpwith the other member of the pair. The PCR products were purifiedby agarose gel electrophoresis. The second step involved theupstream XhoI primer, the downstream BamHI primer, and the twoPCR products from the first step overlapping at the same mutationfrom the first step, and the complementary strands that formeda duplex in the overlapping mutant region were completed. Thissecond-step PCR product, which contained a duplex version ofthe mutant sequence at the appropriate internal site, was digestedwith XhoI and BamHI and then cloned into the XhoI-BamHI siteof pBluescript, for structure verification by sequencing. Verifiedclones were transferred from pBluescript into the XhoI-BamHIsite of pREG1727. Plasmid pREG1727, constructed by Ron Gillof the University of Colorado and described in reference 10,forms transcriptional fusions between the cloned fragment anda promoterless lacZ gene. Cloning into the XhoI-BamHI site ensuredthat the promoter would be oriented properly for transcriptionof lacZ. The cloned mutant plasmids are described in Table 1.pREG1727 contains the prophage attachment site attP of myxophageMx8 which promotes site-specific integration of this plasmidinto the attB locus of the M. xanthus chromosome (12). pREG1727also carries the NPTII gene, encoding kanamycin resistance,which can be used for selection of a plasmid integrant intoDK1622, since pREG1727 cannot replicate as a plasmid in M. xanthus(12). That each new plasmid had integrated into the attB locusof DK1622 was confirmed by Southern blot hybridization of theproducts of restriction endonuclease digestion. The mutatedsequence was again verified. Finally, each integrant strainwas induced to develop, and its ß-galactosidase activitywas measured relative to that of wild type in the same experiment,as described previously (16). Each expression test was performedthree to six times, to obtain a reliable average. Specific activitieswere measured as nanomoles of o-nitrophenol per minute per milligramof protein.

Nucleotide sequence accession number. The sequence of the act operon DNA is published in GenBank underaccession number AF350253.

RESULTS

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Results

Dynamics of act operon expression. A translational fusion between ActA, the first protein of theoperon, and LacZ was constructed for initial measurements ofact operon expression. A 1,696-bp SmaI restriction fragmentthat starts 906 bp upstream of the actA (MXAN3213) translationstart, which is located at position 1402 on the fragment shownin Fig. 1A, was inserted into the SmaI site of plasmid pJBZ281.The SmaI site at position 496 lies within MXAN3211; consequentlythe cloned fragment includes MXAN3211, the 333-bp intergenicregion upstream of actA, and the N-terminal 263-amino-acid polypeptideof ActA translationally fused to LacZ. This fusion plasmid integratedby homologous recombination into the act region of the chromosomeof DK1622, the strain which provided a standard genetic backgroundfor this work. Insertion generated a tandem duplication strain,DK10602; its chromosomal structure, which was confirmed by Southernanalysis, is shown in Fig. 1B. The fruiting body sporulationefficiency of DK10602 was equal to that of DK1622: at 3 days,there were 6.4 x 10⁵ viable spores for DK1622 and 6.0 x 10⁵spores for DK10602. Therefore, the 906-bp upstream segment whichincludes MXAN3211 and the 333-bp intergenic region in DK10602contains all the necessary positive elements for act expression(Fig. 1B).

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FIG. 1. Map of the act operon. (A) DNA fragment containing the region upstream of the operon and the N-terminal part of the ActA protein. (B) The fragment shown in panel A was cloned into pJBZ281, and the plasmid was inserted by homologous recombination into the act region of DK10602, a translational fusion. (C) The transcriptional fusion plasmid pTG027 was inserted in strain DK1622, creating DK10642. Position 331 is on the universal PCR primer, described in Materials and Methods.

Developmental expression of the act operon, as measured in DK10602,appears to rise in two distinct phases (Fig. 2). It increasesgradually during the first 18 h of development and then risesrather steeply to a maximum at 21 to 24 h, when heat-resistantspores begin to appear. A single, continuous rise in expressionbeginning at 6 h seems incompatible with the experimental data,given the size of measurement errors shown by the bars in Fig.2. To probe the existence of two distinct phases of act expression,the same 906-bp upstream sequence of strain DK10602 was clonedinto pREG1727, to generate a transcriptional fusion of actAto lacZ, and to build a reporter plasmid, pTG027. This fusionplasmid has translational stops in all reading frames aheadof lacZ, ensuring that the fusion of actA to lacZ is transcriptional,not translational. A transcriptional fusion eliminates any effectsthat translational fusion might have that limit protein synthesis.Plasmid pREG1727 is designed to integrate by site-specific recombinationinto the Mx8 attB site of M. xanthus (12), which is distantfrom the chromosomal act region. Figure 1C shows pTG027 insertedinto the chromosome of DK1622 at the Mx8 prophage attB site,creating strain DK10642. The chromosomal act region was foundto be undisturbed in this strain; its expression should be normal,and pTG027 should be an accurate reporter. Southern blot assaysperformed on DK10642 supported the idea that insertion had occurredwithin the attB site and not within the act operon. The sporetiter of DK10642 (6.1 x 10⁵) was the same as that of its DK1622parent (6.4 x 10⁵), confirming that the chromosomal act regionwas normal. The solid line with filled points in Fig. 3A showsmeasurement of DK10642 lacZ expression. Again, two phases oflacZ expression are evident. A gradual increase during the first18 h is followed by a rapid increase to 24 h. Because the expressionprofiles of Fig. 2 and Fig. 3 are similar, it is likely thatthe two transcriptional phases seen with DK10642 are responsiblefor the two phases of translation seen with the fusion DK10602.Thus, Act protein levels are primarily controlled at the levelof transcription in these strains, and they can be used as sensitivereporters of transcription.

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FIG. 2. Expression of ß-galactosidase in the translational fusion strain DK10602. Specific ß-galactosidase activity is shown during the first 30 h of development with error bars showing the standard deviations. Where error bars are not evident, they lie inside the square symbols centered on the average.

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FIG. 3. A. Transcription of ß-galactosidase in DK10642 and mutants defective in asgA, fruA, and sdeK. Closed circles, DK10642; open circles, sdeK mutant; open squares, fruA plasmid insertion mutant; closed squares, fruA Tn5 insertion mutant; open diamonds, asgA mutant. B. Transcription of ß-galactosidase in DK10627 (closed circles) and mutants defective in csgA (open diamonds), actA (closed squares), actB (open circles), and actC (open squares). Specific activity is shown as nanomoles of o-nitrophenol per minute per milligram of protein.

To find transcriptional regulators that are needed for act operonexpression, pTG027 with the act insert was transduced or transfectedinto a series of candidate regulatory mutant strains derivedfrom DK1622. Figure 3A shows evidence for two similar phasesof act expression in an sdeK mutant background. The very similartime course of the ${Delta}$ sdeK strain compared with DK10627, whichhas the wild-type allele of sdeK, indicates that act transcriptiondoes not depend on sdeK. The sdeK gene encodes a histidine proteinkinase that has been shown to be essential for expression ofmany developmental genes (11, 36, 48). The substrate(s) forthe putative sdeK kinase has not yet been identified, and thedata in Fig. 3 imply that neither FruA, for which there is evidenceof phosphorylation correlated with activity (8), nor any ofthe act products—among which ActB is likely to be phosphorylated—isa substrate for the sdeK kinase. However, it is evident fromFig. 3A that transcription of act depends on asgA, since expressionis greatly depressed in that mutant. Expression of act alsodepends on the response regulator protein FruA, which has beenverified with two different fruA insertion mutations (8, 47).The Ogawa fruA plasmid insertion mutant happens to be in a DZF1background that contains a pilQ1 motility mutation (47), whichmay account for the slightly lower level of act expression thanin the Ellehauge Tn5 insertion mutant, which is pilQ⁺. Expressionof fruA depends on the A signal (8). Together, the asgA andfruA defects in act operon expression imply a FruA requirementfor the transcription of act.

The second phase of act expression that is evident in Fig. 2and Fig. 3A rises steeply at 20 h when CsgA protein is reachingits highest specific activity under the same conditions (17).The data of Fig. 3B indicate that transcriptional expressionof act directly depends on C signal for several reasons. First,there is very little, if any, increase in act expression ina csgA mutant. Second, the ${Delta}$ actC mutant, which gives precociouscsgA expression (16), shows precocious expression of the actAtranscriptional reporter. This argues that the level of C signalnormally limits actA expression. The steepness of the earlyrise in the ${Delta}$ actC expression profile in Fig. 3B recalls the secondphase of act expression in DK10642 cells. For that reason, itappears that in the ${Delta}$ actC mutant phase two is precocious whilephase one is truncated. Together, the effect of the null csgAand of the ${Delta}$ actC mutants implies that the steep phase of actexpression depends on C signaling. The early phase is depressedin both the asgA mutant and in two fruA mutants (Fig. 3A).

A promoter for act. Figure 3B shows that act reporter expression is substantiallyand equally depressed in the ${Delta}$ actA and ${Delta}$ actB mutants. Inasmuchas actB encodes a sigma-54 activator protein (15), and actAencodes a response regulator that works with actB (17), thedependence of act expression on actB strongly suggests thatthe act operon is expressed from a sigma-54 promoter, and thatinference has been tested by site-directed mutagenesis.

When the actA, actB, actC, and actD genes are expressed as anoperon, they are transcribed from a single adenosine residue57 nt ahead of the presumed actA translational start (17). Thesequence ahead of the transcription start site, taken from theGenBank sequence with accession number AF350253, is shown inFig. 4. The entire sequence was verified in the complete sequenceof the M. xanthus genome, GenBank accession number CP000113.A proposed ribosome binding site (TGAGGAAGT) and the ATG translationstart codon of actA (nucleotide +1) that lie to the right ofthe sequence shown in Fig. 4 are evident in the published sequence.Relative to the transcription start, the upstream region from–167 to –12 is spelled out in Fig. 4. It includes–15 to –29 with a TGGCAC-N₅-TTGC sequence resemblingthe sigma-54 consensus from a compilation of sigma-54 promoters(2). That compilation shows that transcription initiates from8 to 21 nt downstream from the highly conserved GC element inthe proximal (–12) element of a sigma-54 promoter (2).In the act operon, transcription starts 15 nt from that GC element.This resembles the extensively studied M. xanthus sigma-54 pilApromoter (71). It also resembles the M. xanthus ${Omega}$ 4521 promoter,for which sequence mutations have indicated sigma-54 specificity(29). The sigma-54 gene (rpoN), which is in single copy, isessential for M. xanthus development. Uniquely among bacteriachecked, rpoN of M. xanthus is also essential for growth (30).

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FIG. 4. DNA sequence of the act promoter region from residues –167 to –12. The transcription start, AGC, is residue +1, and it is located to the right of the sequence shown. The –12 and –24 regions of a sigma-54 promoter, two potential enhancer sequences, and two potential C boxes are included. Boxed elements in the line were replaced by the sequences shown below them at the arrow tips, and the numbers next to the new sequence give the resulting level of actA::lacZ fusion reporter expression, expressed as a percentage of the unmutated sequence. Specific ß-galactosidase activities relative to the wild-type promoter were measured as described in Materials and Methods. Site-specific mutations were introduced with the PCR primers presented in Table S2 in the supplemental material.

Individual nucleotides in the promoter-like sequence were alteredand then tested for level of act expression. Mutant promoterswere generated by sequence overlap extension PCR (Materialsand Methods), using the oligonucleotides shown in Table S2 inthe supplemental material. The mutated segments were then individuallycloned into pREG1727, replacing the normal sequence with itsmutant versions in pTG027. This cloning incorporated each ofthem into the same transcriptional fusion segment so that theactivity of each mutant promoter could be quantified by measuringthe ß-galactosidase activity relative to that of thesequence without mutation. ß-Galactosidase specificactivity was measured 24 h after initiation of development,the time at which maximum act expression was observed from thenormal promoter, as indicated in Fig. 2 and Fig. 3. To compensatefor the biological variation in the level of gene expression,each mutant-to-wild-type comparison was made within the sameexperiment. Figure 4 presents the relative ß-galactosidaseactivities observed in each mutant as relative promoter activityin percent. Changes in the –14 to –20 region, whichis the downstream or proximal oligonucleotide element of thepromoter (the –12 element), show that those bases areimportant for expression. By the change of residue –14from T to C, activity fell to 34% of the original level; changingresidue –16 from G to a nonconsensus T reduced activityto 7% of that of the wild-type promoter. Changing residue –18from T to C reduced activity to 20% of the original level, andat –20 changing C to T decreased activity to 50% of theoriginal level. Changes in the upstream promoter element (the–24 element) at residue –24 or –27 decreasedactivity to less than 10% of that of the wild-type promoter.By contrast, changing the –30 G residue to T had no effect,suggesting that –30 does not have a significant interactionwith sigma-54 holoenzyme (Fig. 4). If the act promoter wererecognized by a member of the sigma-70 family of factors, assumingthat –30 is part of the –35 region, an effect wouldhave been expected (18). M. xanthus is known to employ severalmembers of the sigma-70 family of factors (67).

One of the largest mutant defects (93%) was obtained by deletingthe residue at position –21, which lies between the upstreamand downstream elements of a sigma-54 promoter. This defectexemplifies a requirement for proper spacing between the –12and –24 elements, as has been observed elsewhere (3, 29,43-45). Strict spacing between promoter elements helps to distinguishsigma-54 promoters from sigma-70 promoters (2, 18). Consideringthe conserved residues, the interelement spacing, and the distancefrom the act transcription start, the data strongly supportrecognition by sigma-54.

Upstream regulatory sequences. Sigma-54 promoters often have upstream regulatory elements.To identify regulatory elements, five to seven base pairs werereplaced in each of the potential regulatory sites, indicatedat the top of Fig. 4. Kroos and coworkers have identified aset of septanucleotide sequences, named C boxes, near the transcriptionstart of a number of established C-signal-dependent promoters(9, 41, 61, 66, 73-75). Expression of the act operon is C signaldependent (17), but C boxes have yet to be reported with sigma-54promoters. Two sequences that resemble the C-box consensus CAYYCCY(where Y = C or T) (41) are evident –167 and –120nt upstream of actA (Fig. 4). Both sequences were mutated, replacingall seven or four of seven bases with nonconsensus residues.Replacing all five residues of potential box 2 lowered activityto 17% of the wild-type level, a substantial decrease (Fig.4). Changing all seven residues of potential box 1 significantlydecreased the activity to 45% of the wild-type level (Fig. 4).These changes may be contrasted with mutation of a hexanucleotidesequence, "control," in the same general area, centered at –130.Replacement of all six bases of this sequence, which lies betweenthe two C boxes and between the two enhancer-like sequencesdescribed below, was found to have no detrimental effect (Fig.4).

Holoenzyme E ${sigma}$ ⁵⁴ forms physically detectable closed-promoter complexesat known ${sigma}$ ⁵⁴ promoters, but the enzyme is unable to initiatetranscription spontaneously. To separate the polynucleotidestrands of an E ${sigma}$ ⁵⁴-DNA complex requires an enhancer sequencein the DNA (42). A promoter-specific activator protein bindsthe enhancer to interact, by DNA looping, with holoenzyme boundat the promoter (51, 62, 68). Two candidate sequences, whichare boxed in Fig. 4, were identified by comparing the sequenceupstream of the act promoter with several sigma-54 enhancersthat have been studied—the enhancers for glnAp₂ and pspAin Escherichia coli (50, 69) and for nodD3 in Sinorhizobiummeliloti (6). Two sequences upstream of the act promoter areindicated, one located around –150 (candidate enh1) andanother sequence located around –110 (candidate enh2).Candidate enhancer 2 includes C box 2 (Fig. 4). Because theglnAp₂ enhancers have dyad symmetry, the two half-sequenceswere separately mutated. Localized multibase mutations wereobtained by substituting either A₆, C₆, _or G₅ for the fiveor six resident bases (see Table S2 in the supplemental material).Candidate half-sequence 2-1 proved important for expressionof the act operon, since expression in the substitution mutantfell to 17% of the wild-type level (Fig. 4). Mutation of candidateelements 1-1, 1-2, and 2-2 showed expression reduced to 60%,62%, and 60% of wild-type levels, respectively (Fig. 4). Thepossibility that A₆ bends DNA should be noted.

actE. Examination of the act DNA sequence revealed an open readingframe that starts just downstream of actD and that could encodea polypeptide of 753 amino acids. Figure 5 presents a fine-scalemap of the whole region. The newly recognized gene, MXAN3217,named actE, is cooriented with the other act genes and is likelyto be cotranscribed with actA and actB, according to the timepattern of hybridization of developmental RNA to open readingframe-specific probes (Fig. 6). Deletion of actB decreases theamount of actE transcript to the same extent as it decreasesactA and actB transcript levels, as if all were transcribedtogether. The actE stop codon is at nucleotide residue 8591in the GenBank sequence with the accession number AF350253.A 7.3-kb act mRNA has been reported elsewhere (17)—justlong enough to extend from the beginning of actA through theend of actE (Fig. 5). Mutations in actE have not yet been sought,and no sequence homologs have been found to suggest function.

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FIG. 5. The act operon, showing the location of all act genes, including actE. Horizontal arrows indicate the promoter and direction of transcription of each gene.

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FIG. 6. Relative levels of act operon mRNA at 0, 8, and 24 h of development are shown expressed as percentages. The 24-h level was set as 100%. DNA probes specific for actA and actB were described in reference 17, and an ApaI fragment was employed for actE. The mRNA levels were quantified by measuring ³²P on slot blots as described in Materials and Methods.

DISCUSSION

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Discussion

The effects of sequence changes in the act operon promoter ofM. xanthus are strong evidence that the promoter is recognizedby sigma-54. Considering the large number of sigma-54 activatorproteins encoded by the genome (13, 21), and with this additionto the list of established sigma-54 promoters, it is evidentthat sigma-54 regulates a substantial part of fruiting bodydevelopment. The mutations reported here lie in the right halfof the 333-bp segment that separates the actA (MXAN3213) transcriptionstart from the start of MXAN3211, an open reading frame which,if expressed, is expected to be transcribed in the oppositedirection from act (Fig. 5). Two candidate sigma-54 enhancerelements were found, one around –150 and one around –110.Site-specific mutagenesis of those two elements led to an 80%activity reduction from the wild type when the –110 elementwas mutated and a smaller, but significant, reduction when theone at –150 was mutated. Direct evidence for enhancerfunction is lacking.

The product of actB is shown here to be an activator of actoperon expression, since deletion of actB decreases expressionof actA (Fig. 3B) and of actE (Fig. 6). ActB might interactwith one or both of the potential enhancers, but no bindingstudies have been carried out. ActA and ActB appear to be partof a signal transduction pathway that receives input from Csignal on the transmitting cell. The output of this signal transductionpathway would be a transcriptionally active ActB (perhaps phosphorylatedActB [ActB $~$ P]), whose absence would account for the reduced actoperon expression in a C-signal-deficient or an ActA-deficientmutant (Fig. 3B). Thus, the rapidly rising late phase of ActAand ActB expression evident in Fig. 2 and Fig. 3 could be explainedby ActB binding to the enhancer of its own promoter. Becausea positive feedback loop would thereby be created, an acceleratedrise in expression would be expected.

Expression of the act operon starts by 6 h of development, afterthe traffic jams that nucleate aggregates have formed (28).Ellehauge et al. have shown that A signaling, but not C signaling,is required for fruA expression (8). FruA protein is a DNA-bindingtranscriptional activator that is necessary for expression ofa number of developmentally regulated genes, devRS, ${Omega}$ 4400, ${Omega}$ 4469, ${Omega}$ 4273, ${Omega}$ 4500, and fdgA (8, 47, 64, 75), as well as for act. FruAbelongs to the FixJ family, and it begins to be expressed between3 and 6 h (8, 47), in preparation for aggregation gene expression,including the act genes. ActB belongs to the NtrC family oftranscriptional activators, which have an ATPase domain locatedbetween their receiver and DNA-binding domains that FruA lacks.It is attractive to think that FruA is a positive transcriptionalregulator of the act operon for the early phase of act operonexpression that occurs at a lower rate than the late phase becauseActB has an ATPase activity to drive promoter opening whileFruA does not.

Early-phase expression of the act operon would produce someActB protein. ActB could then bind one or more specific enhancers.Using its ATPase to open the sigma-54 promoter, ActB would initiatethe second, rapidly rising phase of act transcription. In additionto actB, csgA and actA are also needed for the second, morerapid phase of act expression (Fig. 3B). p17 CsgA on the transmittingcell, via a (currently unknown) receptor on the receiving cell,would signal ActA, which, in turn, would activate ActB, presumablyto ActB $~$ P. With active ActB protein, expression of the entireact operon would accelerate, rapidly producing all the Act proteins,including ActC and ActD for timing.

Recently it has been suggested that the actD gene, MXAN3216,is a cysteine protease homolog that belongs to clan CD, whichincludes family C14 of caspase-like enzymes EC 3.4.22.36 andwhich is directed toward particular proteins (1, 4, 63, 65).Members of this family have a His/Cys catalytic dyad aroundwhich sequence is conserved. In actD the His/Cys residues arefound in the consecutive sequences LVYYSGHS and LDSCASG (withputative catalytic residues underlined). Because deletion ofactD delays C-signal-dependent gene expression, the normal timingof csgA expression is likely to involve proteolysis. Since deletionof actC leads to precocious C-signal-dependent gene expression,actC protein appears to inhibit the rise of csgA expression.One scheme for the temporal control of csgA expression wouldstart with production of comparable amounts of ActC and ActDproteins from their coexpression, and ActC (MXAN3215) wouldinhibit csgA expression. ActD caspase could degrade ActC, releasingthe inhibition and allowing the expression of csgA to rise atthe appropriate time. The final level of csgA expression isset by actA and actB (17). Since C signaling stimulates csgAexpression, ActB $~$ P might be a transcription factor or an activatorof a transcription factor for the csgA promoter.

ACKNOWLEDGMENTS

Lisa Gorski was the first to identify actB, and we thank herfor help.

This investigation was supported by U.S. Public Health Servicegrant GM 23441 to D.K. from the National Institute of GeneralMedical Sciences.

FOOTNOTES

* Corresponding author. Mailing address: Department of Developmental Biology, Beckman Center, B300, Stanford University School of Medicine, 279 Campus Drive, Stanford, CA 94305-5329. Phone: (650) 723-6165. Fax: (650) 725-7739. E-mail: kaiser{at}cmgm.stanford.edu.

Published ahead of print on 22 December 2006.

Supplemental material for this article may be found at http://jb.asm.org/.

Present address: Center of Advanced European Studies and Research,Ludwig-Erhard-Allee 2, 53175 Bonn, Germany.

REFERENCES

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