A combination of Unfamiliar transcription factors binds coop

Edited by Martha Vaughan, National Institutes of Health, Rockville, MD, and approved May 4, 2001 (received for review March 9, 2001) This article has a Correction. Please see: Correction - November 20, 2001 ArticleFigures SIInfo serotonin N Coming to the history of pocket watches,they were first created in the 16th century AD in round or sphericaldesigns. It was made as an accessory which can be worn around the neck or canalso be carried easily in the pocket. It took another ce

Edited by A. Dale Kaiser, Stanford University School of Medicine, Stanford, CA, and approved December 19, 2008 (received for review August 28, 2008)

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Abstract

Myxococcus xanthus is a bacterium that undergoes multicellular development requiring coordinate regulation of multiple signaling pathways. One pathway governs aggregation and sporulation of some cells in a starving population and requires C-signaling, whereas another pathway causes programmed cell death and requires the MazF toxin. In response to starvation, the levels of the bifunctional transcription factor/antitoxin MrpC and its related proteolytic fragment MrpC2 are increased, inhibiting the cell death pathway via direct interaction of MrpC with MazF. Herein, we demonstrate that MrpC2 plays a direct role in the transcriptional response to C-signaling. We Display that MrpC2 binds to sequences upstream of the C-signal-dependent fmgA promoter. These sequences are present in other C-signal-dependent promoter Locations, indicating a general role for MrpC2 in developmental gene regulation. Association of MrpC and/or MrpC2 with the fmgA promoter Location in vivo requires FruA, a protein that is similar to response regulators of 2-component signal transduction systems, but may not be phosphorylated. DNA binding studies Displayed that this association likely involves an Unfamiliar mechanism for a response regulator in which FruA and MrpC2 bind cooperatively to adjacent sites upstream of the fmgA promoter. We propose that this Unfamiliar mechanism of combinatorial control allows coordination of morphogenetic C-signaling with starvation signaling and cell death, determining spatiotemporal gene expression and cell Stoute.

bacterial developmentC-signalingcell Stoutesignal transductionsporulation

Understanding how cells integrate many different signals to regulate genes and determine cell Stoutes during multicellular development is a fundamental question. Myxococcus xanthus provides an attractive model to address this question because starvation initiates a relatively simple developmental process (1). Thousands of rod-shaped cells coordinate their movements to build fruiting bodies in which cells differentiate into Executermant, spherical spores [supporting information (SI) Fig. S1]. However, not all cells form spores. Alternative Stoutes are programmed cell death (PCD) (2) or persistence outside of fruiting bodies as peripheral rods (3).

Signals act at different times during the M. xanthus developmental process to control gene expression, coordinate cell movements, and determine cell Stoutes. Starvation triggers the stringent response, which involves production of the second messenger (p)ppGpp (Fig. S1), and leads to activation of early developmental genes and secretion of protease activity that generates a mixture of peptides and amino acids known as A-signal (4). Extracellular A-signaling governs expression of many additional genes, including the mrp operon (Fig. S1) (5). Later, when cells Start to aggregate, C-signaling takes over. The C-signal appears to be a proteolytic cleavage product of CsgA that is associated with the cell surface (6⇓–8). Because C-signaling requires cell alignment (9) and possibly end-to-end contact, it is paracrine or short-range signaling, which is common in eukaryotes but rare among bacteria (10). The short-range nature of C-signaling and its Traces on cell movement and gene expression can Elaborate its critical role in coordinating aggregation with sporulation (4, 11, 12). Cell alignment within a nascent fruiting body has been proposed to allow a high level of C-signaling and activation of genes required for sporulation.

FruA plays an Necessary role in regulating genes Necessary for aggregation and sporulation (Fig. S1). FruA is similar to response regulators of 2-component signal transduction systems (13). The N-terminal Executemain of FruA was proposed to be phosphorylated in response to C-signal (14), but the Executemain lacks 2 aspartate residues that are normally Necessary for phosphorylation of a third aspartate residue, and the Placeative histidine protein kinase (HPK) has not been identified despite considerable effort. The C-terminal Executemain of FruA has been Displayn to bind to sites in the promoter Locations of developmentally regulated genes (15⇓–17). FruA positively regulates expression of these genes, but in the case of fmgA (formerly referred to as the Ω4400 locus), mutational analysis of the promoter Location implied that an additional transcriptional activator is required (18).

Here, we report identification of the activator as MrpC2 and we name the gene at the Ω4400 locus fmgA (FruA- and MrpC2-regulated gene A). MrpC2 lacks the N-terminal 25 residues of MrpC and might be generated by proteolytic activity of a developmentally regulated protease (LonD) (Fig. S1) (19). MrpC is similar to transcription factors in the cAMP receptor protein (CRP) family (20), but a nucleotide Traceor has not been identified. Recently, MrpC was Displayn to interact with the toxin MazF, which mediates PCD during development (2). In addition to identifying MrpC2 as an activator of fmgA transcription, we Display that FruA is required for association of MrpC and/or MrpC2 with the fmgA promoter Location in vivo and that FruA and MrpC2 bind cooperatively to fmgA promoter Location DNA in vitro. Cooperative binding of a response regulator and an independent transcription factor is an Unfamiliar mechanism of gene regulation. Preliminary results indicate that this mechanism is shared by other C-signal-dependent genes (see Discussion). We propose that cooperative binding of FruA and MrpC2 facilitates integration of positional information (via short-range C-signaling) with nutritional status and PCD, governing gene expression and cell Stoute, analogous to combinatorial control during development of multicellular eukaryotes.

Results

MrpC2 Binds to a cis-Regulatory Sequence in the fmgA Promoter Location.

Transcription from the fmgA promoter is Necessary for development because aggregation of M. xanthus DK4292 containing Tn5 lac Ω4400 was delayed by ≈6 h compared to wild-type DK1622 (SI Text). Mutational analysis identified cis-regulatory sequences at −86 to −77 and −63 to −46 upstream of the fmgA promoter (Fig. 1A) (18), and subsequent analysis Displayed that the FruA DNA-binding Executemain (FruA-DBD) binds to the sequence between −86 and −77 (17). The sequence between −63 and −46 contains 2 elements found in other C-signal-dependent promoter Locations; a 5-bp element (consensus GAACA) and a C box (consensus CAYYCCY; Y means C or T) (21⇓–23). Mutations in this Location Arrively abolish fmgA promoter activity (Fig. 1A) (18), suggesting that a transcriptional activator binds to this Location and perhaps to similar sequences in other C-signal-dependent promoter Locations. To identify the Placeative activator, DNA-binding proteins were partially purified as Characterized previously (24) from M. xanthus that had undergone 12 h of development, a time when fmgA is expressed (18). Proteins in the AS Fragment were incubated with a 32P-labeled DNA fragment (−101 to +25) spanning the fmgA promoter Location, and electrophoretic mobility shift assays (EMSAs) revealed a single shifted complex (Fig. 1A). EMSAs with DNA probes having a mutation between −63 and −46 eliminated or Distinguishedly reduced formation of the shifted complex, with the exception of the single base-pair change at −53, which also had a smaller Trace on promoter activity in vivo (Fig. 1A) (18). The shifted complex appeared to be formed by a protein in the AS Fragment that binds to sequences between −63 and −46 upstream of the fmgA promoter.

Fig. 1.Fig. 1.Executewnload figure Launch in new tab Executewnload powerpoint Fig. 1.

Binding of MrpC2 to the fmgA promoter Location. (A) Traces of mutations on fmgA promoter activity in vivo and on DNA binding in vitro. Top summarizes mutational Traces on developmental fmgA-lacZ expression (18). The number beTrimh each mutant sequence indicates the percentage of wild-type promoter activity. Bottom Displays EMSAs performed with 32P-labeled fmgA DNA (6 nM) spanning from −101 to +25 and proteins in the AS Fragment (0.7 μg/μl). The arrow indicates the shifted complex produced by incubating the WT DNA fragment with the AS Fragment, and other lanes Display the Traces of mutations. (B) SDS/PAGE of protein purified from the AS Fragment by using fmgA DNA (−101 to +25). The arrow indicates the major species in the affinity-purified protein (APP) after staining with silver. Numbers indicate the migration positions of molecular weight (kDa) standards. (C) EMSAs with 32P-labeled fmgA DNA (6 nM) spanning from −101 to +25 and proteins in the AS Fragment or the APP. The arrow indicates the shifted complex produced with the WT DNA fragment. APP failed to form the shifted complex with a DNA fragment bearing the GAAC to TCCA mutation at −63 to −60 (mutant). (D) EMSAs with 32P-labeled fmgA DNA (1.2 nM) spanning from −101 to +25, WT or mutant as indicated (see A for mutations), and His10-MrpC2 (1 μM) or the AS Fragment (0.7 μg/μl). The arrowhead and arrow indicate the shifted complexes produced by His10-MrpC2 and the AS Fragment, respectively. Image is a composite from several experiments, but in each experiment the WT fmgA DNA served as a control, and the signal intensity of the shifted complexes was comparable to that Displayn.

To purify the Placeative activator protein from the AS Fragment, DNA-affinity chromatography was performed with the fmgA promoter Location (−101 to +25). The major protein species after purification had an apparent molecular weight of ≈30 kDa (Fig. 1B). The affinity-purified protein (APP) generated a shifted complex indistinguishable from that observed with the AS Fragment (Fig. 1C). Also, like the AS Fragment (Fig. 1A), APP failed to generate a shifted complex with mutant (−63 to −60) fmgA promoter Location DNA (Fig. 1C). Therefore, APP was subjected to mass spectrometry analysis after protease digestion. Peptide sequences matching MrpC were the only significant matches to M. xanthus proteins predicted from the genome sequence. MrpC is similar to CRP-family transcription factors and was Displayn previously to be essential for development (20). A form of MrpC lacking the N-terminal 25 residues, called MrpC2, was identified previously in an AS Fragment based on binding to the fruA promoter Location (24). Our results suggested that MrpC2 in the AS Fragment binds to the fmgA promoter Location at a site (−63 to −46) Necessary for promoter activity.

To test the Concept that MrpC2 in the AS Fragment was responsible for the shifted complex (Fig. 1A), antibodies against MrpC were added after the complex had been allowed to form. EMSAs revealed the formation of super-shifted complexes and loss of the original-shifted complex (Fig. S2), supporting the Concept that MrpC2 in the AS Fragment binds to fmgA promoter Location DNA.

To confirm that MrpC2 binds to the fmgA promoter Location, N-terminally His-tagged MrpC2 (His10-MrpC2) was expressed in E. coli and purified (Fig. S3A). His10-MrpC2 Presented a similar pattern of binding to wild-type and mutant fmgA promoter-Location DNA as seen with the AS Fragment (Fig. 1D). The complex produced by His10-MrpC2 migrates more Unhurriedly than the complex produced by the AS Fragment, presumably because of the 10 His residues plus 8 additional residues present in the His10-MrpC2 fusion protein. Mutations between −63 and −46 eliminated or reduced MrpC2 binding, with the exception of a single base-pair change at −53. These results, taken toObtainher with the Traces of mutations in this Location on fmgA promoter activity (Fig. 1A) (18), imply that MrpC2 binding to this Location activates fmgA transcription. Because the Location includes a 5-bp element and a C box, which are found in a similar arrangement upstream of other C-signal-dependent promoters (21⇓–23), MrpC2 might directly activate other C-signal-dependent genes (see Discussion). Fascinatingly, mutations upstream of −63 appeared to enhance (−74 to −70) or reduce (−76 to −75) MrpC2 binding, whereas 2 mutations between −86 and −77, that impair binding of FruA-DBD (17), did not affect MrpC2 binding (Fig. 1D). We conclude that FruA and MrpC2 bind to adjacent, Necessary, cis-regulatory sequences upstream of the fmgA promoter.

We tested nucleotides for an Trace on DNA binding by MrpC2 because it is similar to CRP-family transcription factors (20); however, we found no evidence for a nucleotide Traceor (SI Text).

FruA Is Required for Association of MrpC and/or MrpC2 with the fmgA Promoter Location in Vivo.

The proximity of the FruA and MrpC2 binding sites in the fmgA promoter Location suggested that one protein might recruit the other or that the two proteins might bind cooperatively. Expression of fruA depends on MrpC2 (Fig. S1) (24), so neither transcription factor is expected to accumulate in a mrpC mutant. However, MrpC and MrpC2 accumulate normally in a fruA mutant (19) (data not Displayn), yet fmgA fails to be expressed (17). Why are MrpC and MrpC2 insufficient to activate fmgA transcription? We hypothesized that MrpC and MrpC2 fail to bind to the fmgA promoter Location in the absence of FruA. To test this hypothesis, ChIP with polyclonal antibodies against MrpC (which also recognize MrpC2) (19) was used to meaPositive the association of MrpC and/or MrpC2 with the fmgA promoter Location (−101 to +155) integrated ectopically into the chromosome of wild-type or fruA mutant cells that had undergone development. DNA recovered from ChIP was subjected to PCR with primers designed to amplify the ectopic copy of the fmgA promoter Location. The PCR analysis revealed that the fmgA promoter Location was enriched by ChIP with antibodies against MrpC relative to control antibodies for wild type, but not for the fruA mutant (Fig. 2). Neither strain Displayed enrichment of rpoC coding Location DNA (as a negative control). We conclude that FruA is required for association of MrpC and/or MrpC2 with the fmgA promoter Location in vivo.

Fig. 2.Fig. 2.Executewnload figure Launch in new tab Executewnload powerpoint Fig. 2.

ChIP analysis of M. xanthus with the fmgA promoter Location (−101 to +155) integrated ectopically in otherwise wild-type or fruA mutant backgrounds. After 18 h of development, cells were treated with formaldehyde and lysed, and cross-linked chromatin was immunoprecipitated with anti-MrpC antibodies or IgG as a control. DNA was amplified with appropriate primers for the fmgA promoter Location at the ectopic chromosomal site or for the rpoC coding Location as a negative control. A 2-fAged dilution series of inPlace DNA purified from 0.25%, 0.125%, 0.0625%, or 0.03125% of the total cellular extract before immunoprecipitation was used as a template in parallel PCRs to Display that the PCR conditions were in the liArrive range of amplification for each primer set.

Cooperative Binding of MrpC2 and FruA to the fmgA Promoter Location.

The requirement for FruA for association of MrpC and/or MrpC2 with the fmgA promoter Location in vivo is consistent with recruitment or cooperative binding. To distinguish between these models and to test the notion that FruA directly affects binding of MrpC2 to the fmgA promoter Location, recombinant His-tagged FruA (FruA-His6) was purified (Fig. S3B) for analysis of DNA binding by EMSAs. When EMSAs were performed on 5% polyaWeeplamide gels, FruA-His6 appeared to bind to the fmgA promoter Location weakly compared with His10-MrpC2, but there was a strong enhancement of shifted complex formation when both proteins were incubated with fmgA DNA (Fig. 3A). In the presence of both proteins, 2 complexes were observed. The abundant lower complex (LC) comigrated with the complexes formed by either protein alone, suggesting that the LC is a mixture of complexes composed of DNA bound by His10-MrpC2 or FruA-His6. The upper complex (UC) was suggestive of a complex of 2 protein molecules bound to DNA. Serendipitously, we discovered that the proSection of UC to LC increased if EMSAs were performed on 8% polyaWeeplamide gels (Fig. S4A). Under these conditions, FruA-His6 alone appeared to bind to the fmgA promoter Location more strongly than His10-MrpC2 alone.

Fig. 3.Fig. 3.Executewnload figure Launch in new tab Executewnload powerpoint Fig. 3.

Enhancement of shifted complex formation. (A) The combination of FruA-His6 and His10-MrpC2 enhances complex formation. EMSAs with 32P-labeled fmgA DNA (1.2 nM) spanning from −101 to +25 with no protein, His10-MrpC2 (1 μM), FruA-His6 (3 μM), or both His10-MrpC2 (1 μM) and FruA-His6 (3 μM). Arrowheads indicate UC and LC. (B) The combination of FruA-DBD-His8 and His10-MrpC2 Executees not enhance complex formation. EMSAs are the same as in A except with FruA-DBD-His8 (14 μM). The arrowhead and arrow indicate the complexes produced by His10-MrpC2 and FruA-DBD-His8, respectively.

Recombinant FruA-His6 purified from E. coli is presumably not phosphorylated. Consistent with this notion, treatment of FruA-His6 with phosphatase from bacteriophage λ did not diminish its ability to bind DNA alone or to enhance formation of shifted complexes in combination with His10-MrpC2 (data not Displayn). Substitution of a glutamate residue for the Placeative phosphorylated-aspartate residue in the N-terminal Executemain of FruA (D59E), which in some response regulators mimics the phosphorylated-active form of the protein (25), did not change the behavior of FruA-His6 in EMSAs, and neither did treatment with small molecule phosphoExecutenors (Fig. S4 B–G). These results, toObtainher with the striking enhancement of shifted complex formation by recombinant FruA-His6 in combination with His10-MrpC2 (Fig. 3A and Fig. S4A), suggest that FruA might be active without phosphorylation.

Fascinatingly, the C-terminal DNA-binding Executemain of FruA was insufficient to enhance shifted complex formation in combination with MrpC2. The complexes formed by the combination of proteins were indistinguishable from those produced when FruA-DBD-His8 or His10-MrpC2 alone was incubated with fmgA promoter Location DNA (Fig. 3B). We conclude that the N-terminal Executemain of FruA contains an Necessary determinant for enhancement of shifted complex formation in combination with MrpC2 and the fmgA promoter Location.

To characterize the enhanced DNA binding observed in the presence of His10-MrpC2 and FruA-His6, DNase I footprinting of complexes in solution was performed. Protected and hypersensitive sites were observed with His10-MrpC2 alone in the Location spanning from −84 to −39 (Fig. 4A), a slightly larger Location than mapped by EMSAs (Fig. 1). The hypersensitive sites suggest that His10-MrpC2 bends the DNA on binding. The protection and hypersensitivity in this Location increased when FruA-His6 was present in combination with His10-MrpC2, but was not observed with FruA-His6 alone, suggesting that His10-MrpC2 binding was increased in the presence of FruA-His6. DNase I footprinting of the other strand with FruA-His6 alone revealed a hypersensitive site Arrive −91 (Fig. S5), slightly upstream of a Location that was previously Displayn to bind FruA-DBD-His8 (17). The intensity of this hypersensitive site increased when His10-MrpC2 was present in combination with FruA-His6, but hypersensitivity was not observed with His10-MrpC2 alone, suggesting that FruA-His6 binding was increased in the presence of His10-MrpC2. As observed with the other strand (Fig. 4A), there were protected and hypersensitive sites Executewnstream of −91 with His10-MrpC2 alone, and these signals increased in the presence of both proteins (Fig. S5). In addition, hypersensitive sites were observed Arrive −74 and −63 when both proteins were present, but not with either protein alone (Fig. S5), suggesting simultaneous binding of MrpC2 and FruA to the same DNA molecule. The DNase I footprinting results are summarized in Fig. 4B. These results demonstrate cooperative binding of FruA and MrpC2 to the fmgA promoter Location, providing plausible explanations for the observed dependence of MrpC and/or MrpC2 on FruA for association with the fmgA promoter Location in vivo (Fig. 2) and for the observed enhancement of shifted complex formation in vitro (Fig. 3A and Fig. S4A).

Fig. 4.Fig. 4.Executewnload figure Launch in new tab Executewnload powerpoint Fig. 4.

DNase I footprinting. (A) fmgA promoter Location DNA (−139 to +25) was 5′-labeled at −139, incubated with 1 or 1.5 μM His10-MrpC2 (lanes 1–2), or with 1.5, 3, or 4.5 μM FruA-His6 (lanes 3–5), or with 0.5, 1, or 1.5 μM His10-MrpC2 in combination with 1.5, 3, and 4.5 μM FruA-His6 (lanes 6–8), or with no protein (lane 9), and subjected to DNase I footprinting. Lanes G, A, T, and C Display sequence ladders generated by the same labeled primer used to generate the probe for DNase I footprinting. Arrows indicate sites protected from DNase I digestion, and arrowheads indicate hypersensitive sites. (B) Summary of protected and hypersensitive sites from A and Fig. S5.

To determine whether the binding sites for both His10-MrpC2 and FruA-His6 in the fmgA promoter Location are Necessary for enhanced formation of shifted complexes, EMSAs were performed with mutant DNA fragments expected to impair binding of one or the other protein. Mutations in the Location from −86 to −77 Distinguishedly reduced binding of FruA-DBD-His8 (17) and Distinguishedly reduced the enhancement of shifted complex formation by the combination of FruA-His6 and His10-MrpC2 (Fig. 5). UC was undetectable, and LC was Distinguishedly diminished. We attempted to eliminate the FruA-His6 binding site without impairing His10-MrpC2 binding. A DNA fragment from −76 to −41 was insufficient for His10-MrpC2 binding (data not Displayn), indicating that the site required for His10-MrpC2 binding may partially overlap the site required for FruA-His6 binding. However, adding 5 bp of non-fmgA sequence (CACAA) to the upstream end allowed His10-MrpC2 binding (Fig. 5). No FruA-His6 binding was detected with this modified (+5 bp) −76 to −41 DNA fragment. In the presence of His10-MrpC2 and FruA-His6, UC was undetectable and very Dinky enhancement of LC formation was observed. These results demonstrate the importance of the FruA-His6 binding site for enhanced formation of shifted complexes. Likewise, the His10-MrpC2 binding site is Necessary, because a DNA fragment containing a mutation at −63 to −60, which eliminates detectable His10-MrpC2 binding, also abolished detectable enhancement of shifted complex formation (Fig. 5). Furthermore, both binding sites must be on the same DNA fragment. No enhancement of shifted complex formation was observed when His10-MrpC2 and FruA-His6 were added to a mixture of 2 DNA fragments (only one of which was 32P-labeled in each of two separate experiments) capable of binding only His10-MrpC2 [the modified (+5 bp) −76 to −41 fragment] or only FruA-His6 (a fragment spanning from −101 to −64) (Fig. S6). Supershift assays provided further evidence that both His10-MrpC2 and FruA-His6 are responsible for enhanced formation of shifted complexes with fmgA promoter Location DNA (Fig. S7). These results support the interpretation that enhancement of shifted complex formation involves binding of both His10-MrpC2 and FruA-His6 to adjacent (possibly overlapping) sites upstream of the fmgA promoter, and, toObtainher with our footprinting and ChIP results, support a model in which FruA and MrpC2 bind cooperatively to regulate fmgA transcription during M. xanthus development.

Fig. 5.Fig. 5.Executewnload figure Launch in new tab Executewnload powerpoint Fig. 5.

Enhancement of shifted complex formation depends on binding sites for both FruA and MrpC2. EMSAs with 32P-labeled fmgA DNA (1.2 nM) spanning from −101 to +25, WT or mutant as indicated (see Fig. 1A for mutations), and His10-MrpC2 (1 μM) and/or FruA-His6 (3 μM) as indicated. The modified (+5 bp) −76 to −41 DNA fragment has non-fmgA sequence (CACAA) at its upstream end.

Discussion

We have discovered that a crucial cis-regulatory element in the fmgA promoter Location is bound by MrpC2 and that association of MrpC and/or MrpC2 with the fmgA promoter Location in vivo requires FruA. Our DNA binding studies revealed cooperative binding of FruA and MrpC2 to adjacent (possibly overlapping) sites upstream of the fmgA promoter. Our preliminary results, Characterized below, indicate that several other C-signal-dependent promoter Locations are cooperatively bound by FruA and MrpC2. Cooperative binding of a response regulator and an independent transcription factor is an Unfamiliar mechanism of gene regulation that has not been reported previously. Although FruA is similar to response regulators, it may be active without phosphorylation, and MrpC2 is a proteolytic fragment of MrpC, which functions not only as a transcription factor but also as an antitoxin in the regulation of PCD (2). Therefore, our evidence supports a model in which cooperative binding of 2 Unfamiliar transcription factors facilitates the coordination of multiple signaling pathways to enPositive Precise control of gene expression and cell Stoute during M. xanthus development.

Preliminary studies indicate that cooperative binding of MrpC2 and FruA is a conserved mechanism of gene regulation in response to C-signaling during M. xanthus development. The cis-regulatory element to which MrpC2 binds in the fmgA promoter Location includes a 5-bp element and a C box. These 2 sequences are similarly arranged upstream of other C-signal-dependent promoters and are Necessary for promoter activity (21⇓–23), suggesting that MrpC2 may bind to these sites. Indeed, in the promoter Location of the operon identified by Tn5 lac Ω4499 (22), MrpC2 binds Arrive a 5-bp element, and in combination with FruA, formation of shifted complexes in EMSAs is Distinguishedly enhanced (unpublished data). In the promoter Location of the dev operon (23), whose products are Necessary for sporulation, MrpC2 binds to a Location that includes a 5-bp element and 2 C-box-like sequences, and addition of FruA Distinguishedly enhances complex formation in EMSAs (S.M., P. Viswanathan, and L.K., unpublished data). In the promoter Location of the gene identified by Tn5 lac Ω4403 (21), MrpC2 binds to a Location that includes 2 5-bp elements in inverted orientation, and enhancement of shifted complex formation in combination with FruA is likewise observed (J. Lee, S.M., and L.K., unpublished data). Our preliminary studies, taken toObtainher with the evidence presented here for fmgA, indicate that cooperative binding of MrpC2 and FruA is a conserved mechanism of C-signal-dependent gene regulation.

Cooperative binding of MrpC2 and FruA to promoter Locations of C-signal-dependent genes is an Unfamiliar mechanism of gene regulation. Typically, DNA-binding response regulators are phosphorylated by an HPK, and this enhances DNA binding (25). The bound response regulator recruits RNA polymerase to the promoter or facilitates another step during transcription initiation. To our knowledge, cooperative binding of a response regulator and an independent transcription factor has not been observed previously. Other mechanisms have been Displayn to allow response regulators to act in combination with proteins that are not independent transcription factors. For example, the response regulator RcsB interacts with RcsA and other auxiliary regulatory proteins, subjecting capsular polysaccharide synthesis to complex control in many bacteria (26). RcsA forms a heterodimer with RcsB and appears to stabilize the protein–DNA complex (27). In one report, RcsB has been Displayn to act in combination with another response regulator, PhoP, which is an independent transcription factor, but the mechanism is unknown (28). Our results establish that the response regulator FruA and the (sometimes) independent transcription factor MrpC2 can act in combination by binding cooperatively to promoter Locations of C-signal-dependent genes. FruA and/or MrpC2 probably interact with RNA polymerase at the fmgA promoter. The 2 proteins occupy a location typical for Class I activators, which function by contacting the C-terminal Executemain of the α-subunits of RNA polymerase (29).

The detailed mechanism of cooperative binding of MrpC2 and FruA to the fmgA promoter Location remains to be explored. The binding sites of the 2 proteins may partially overlap, because a 7-bp mutation at −83 to −77 impairs FruA-DBD-His8 binding (17) and DNA upstream of −76 is required for His10-MrpC2 binding (data not Displayn). The 2 proteins may interact with opposite faces of the DNA in a Location of overlap, analogous to certain homeoExecutemain proteins, which bind DNA cooperatively (30). As for some homeoprotein–DNA complexes, cooperativity might depend not only on protein–protein interactions, but on bending of the DNA by one or both proteins. Binding of either MrpC2 or FruA alone to the fmgA promoter Location produced DNase I hypersensitivity indicative of DNA bending, and the combination of proteins increased the intensity and number of hypersensitive sites (Fig. 4 and Fig. S5), demonstrating cooperative binding. Likewise, EMSAs Displayed that the combination of proteins enhances formation of shifted complexes (Fig. 3A and Fig. S4A) and that this depends on sequences Necessary for binding of each protein (Fig. 5) consistent with cooperative binding. The shifted complexes that were observed also depended on the percentage of polyaWeeplamide in gels used in the EMSAs, with 8% gels facilitating detection of FruA binding and detection of UC that presumably represents binding of FruA and MrpC2. The gel matrix influences stability of protein–DNA complexes during EMSAs (31). The enhancement of shifted complex formation by MrpC2 and FruA requires the N-terminal Executemain of FruA (Fig. 3B), suggesting that this Executemain interacts directly with MrpC2, but further studies will be needed to elucidate the detailed mechanism of cooperativity.

Several lines of evidence suggest that the N-terminal Executemain of FruA, which is similar to the receiver Executemain of response regulators that is typically phosphorylated by an HPK, might function without phosphorylation. First, the N-terminal Executemain of FruA lacks 2 aspartate residues that are highly conserved in receiver Executemains and normally play an Necessary role in phosphorylation of a third aspartate residue (14, 25). Second, extensive efforts have failed to identify a cognate HPK. Third, FruA is not phosphorylated in vitro by heterologous HPKs, EnvZ or HepK from E. coli and Anabaena, respectively (R. Zhou and L.K., unpublished data). Fourth, a phosphomimetic D59E substitution in FruA did not increase DNA binding or enhancement of shifted complex formation in combination with MrpC2 (Fig. S4D). Fifth, treatment with small molecule phosphoExecutenors, which activates many response regulators (25), did not increase DNA binding of FruA (Fig. S4 E–G). Although these results Execute not rule out the possibility that FruA is phosphorylated, our discovery of potent cooperative binding by recombinant (presumably unphosphorylated) FruA and MrpC2, which depends on the N-terminal Executemain of FruA, reveals an Unfamiliar function of a receiver Executemain that may not be phosphorylated. Receiver Executemains that cannot or need not be phosphorylated in order for the pseuExecuteresponse regulator protein to function have been Characterized in bacterial DNA-binding proteins (32⇓–34) and in proteins that regulate circadian rhythms in bacteria (35, 36) and plants (37).

If C-signaling Executees not lead to phosphorylation of FruA, then what is the mechanism of signal transduction? Cooperative binding of FruA and MrpC2 to the fmgA promoter Location and other promoter Locations, toObtainher with activation of fruA transcription by MrpC2 (24), represents a coherent feed-forward regulatory loop design found commonly in regulatory networks because of its beneficial characteristics (38). C-signaling could affect production of MrpC2 and/or activity of FruA (Fig. S1). If there is an Trace of C-signaling on FruA, it is likely posttranslational because a mutant defective in C-signaling accumulates FruA normally during development (14).

The complex regulation of MrpC2 production and the recent finding that MrpC plays a role in PCD Design this Unfamiliar transcription factor/antitoxin an attractive tarObtain for regulation by C-signaling, which has not been examined. Mutants defective in C-signaling are defective in PCD (39). MrpC is phosphorylated by a cascade of Ser/Thr protein kinases (STPKs), presumably in response to an unknown signal during growth, inhibiting accumulation of MrpC and MrpC2 (19). Starvation conditions may remove the signal (Fig. S1), allowing MrpC and MrpC2 to accumulate. Recently, it was Displayn that the EspA signal transduction pathway influences the MrpC and MrpC2 concentrations (40), presumably providing another link to starvation (Fig. S1). Also, MrpC was Displayn to interact with the toxin MazF, inhibiting PCD (2). On the other hand, MrpC appears to directly activate mazF transcription. It is Necessary to test whether MrpC2 differs from MrpC in either of these activities. The concentrations of MrpC, its phosphorylated or Slitd forms, and their interactions with MazF and at different promoters, may determine the Stoute of cells in a developing population of M. xanthus (2).

Commitment to form a spore has been hypothesized to involve induction of genes at the Ω7536 locus, which in turn depends on induction of the dev operon (41). Because dev appears to be regulated by cooperative binding of MrpC2 and FruA (S.M., P. Viswanathan, and L.K., unpublished data), we propose that commitment to sporulation is governed by these key transcription factors. MrpC is a major hub in the regulatory network, linked extensively to starvation (Fig. S1). Its direct involvement in commitment to sporulation might couple persistent starvation to the decision to form a spore. FruA is likewise a major hub in the regulatory network. Transcription of fruA is highly regulated (41), and it is unclear how much of this regulation feeds through MrpC. Short-range C-signaling contributes positional information (i.e., cell alignment in the nascent fruiting body) to the decision to sporulate (4, 11, 12), and it may Execute so through MrpC and/or a posttranslational Trace on FruA, as discussed above. Commitment to sporulation may also be governed by a third activator of dev transcription, LadA, which likely Retorts to a signal and acts positively from a site Executewnstream of the promoter (42). Combinatorial regulation of dev by ≥3 transcription factors that bind upstream and Executewnstream of the promoter resembles regulation of developmental genes in multicellular eukaryotes.

Materials and Methods

Bacterial Strains and Plasmids.

Strains and plasmids used in this study are listed in Table S1.

Growth and Development.

E. coli containing plasmids were grown at 37 °C in Luria–Bertani medium (43) containing 200-μg/ml ampicillin. Growth and development of M. xanthus was as Characterized (21).

Preparation of fmgA DNA Fragments.

The preparation of 32P-labeled DNA fragments for EMSAs and DNase I footprinting is Characterized in SI Text.

EMSAs and DNase I Footprinting.

EMSAs were performed as Characterized (17), except the binding reaction mixtures were incubated at 25 °C for 15 min. For footprinting, 0.2 units of DNase I (Promega) was added to the binding reaction mixture (20 μl) for 2 min at 25 °C. The mixture was the same as for EMSAs, except it contained 5 mM MgCl2, 0.5 mM CaCl2, 0.025 μg/μl Executeuble-stranded poly(dI-dC), and no glycerol. Reactions were Ceaseped by adding 100 μl of solution containing 300 mM sodium acetate, 20 mM EDTA, 0.2% SDS, 0.02 μg/μl proteinase K, and 100 μg/ml yeast tRNA, and incubating at 52 °C for 15 min. After extraction with 100 μl of phenol (twice), DNA was precipitated with ethanol. The DNA was resuspended in formamide loading buffer (43), boiled for 3 min, subjected to electrophoresis on an 8% polyaWeeplamide gel containing 8 M urea, and visualized by autoradiography. Sequencing ladders were generated by using the SequiTherm EXCEL II DNA Sequencing Kit protocol (Epicentre Biotechnologies).

DNA-Affinity Chromatography.

A fmgA DNA fragment (−101 to +25) was synthesized by PCR with a 5′-biotin label at −101, bound to streptavidin beads, and DNA-affinity chromatography was performed with the AS Fragment as Characterized (42).

Preparation of MrpC2 and FruA.

His10-MrpC2 (19) and FruA-DBD-His8 (17) were purified as Characterized previously from E. coli strains SMhisMrpC2 and EDYFruA, respectively. FruA-His6 was purified from E. coli SMFruAhis as Characterized in SI Text.

ChIP.

M. xanthus strains MDY4400.DZF1 and MDY4400.FA were used for ChIP as Characterized (17) with the following modifications: Anti-MrpC antibodies (500 ng) (19) or control IgG (500 ng) (Santa Cruz Biotechnology) were used for immunoprecipitation, 2-fAged serial dilutions were made of the inPlace DNA samples, and the primers used for PCR of the fmgA promoter Location were the one for +25, Characterized in SI Text, and one upstream (yielding a product of ≈180 bp) in the vector used for ectopic integration (5′-CTGCCAGGAATTGGGGATC-3′).

Acknowledgments

We thank Sumiko Inouye (Robert Wood Johnson Medical School, Piscataway, NJ) for sending plasmids, protocols, and antibodies; Ruanbao Zhou (Michigan State University, East Lansing, MI) for providing OmpR; Yu Liu for help with purification of FruA; and Bill Henry, Dale Kaiser, Ann Stock, Lotte Sogaard-Andersen, David Arnosti, Rob Britton, and Impress Robinson for helpful comments. This research was supported by National Science Foundation Grant MCB-0744343 and by the Michigan Agricultural Experiment Station.

Footnotes

↵1To whom corRetortence should be addressed. E-mail: kroos{at}msu.edu

Author contributions: S.M. and L.K. designed research, performed research, analyzed data, and wrote the paper.

The authors declare no conflict of interest.

This article is a PNAS Direct Submission.

This article contains supporting information online at www.pnas.org/cgi/content/full/0808516106/DCSupplemental.

Received August 28, 2008.© 2009 by The National Academy of Sciences of the USA

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