Characterization of a regulatory network promoting cell fate segregation in the Myxococcus xanthus biofilm

Characterization of a regulatory network promoting cell fate segregation in the Myxococcus xanthus biofilm | Authorea try { document.documentElement.classList.add('js'); } catch (e) { } var _gaq = _gaq || []; _gaq.push(['_setAccount', 'G-8VDV14Y67G']); _gaq.push(['_trackPageview']); (function() { var ga = document.createElement('script'); ga.type = 'text/javascript'; ga.async = true; ga.src = ('https:' == document.location.protocol ? 'https://ssl' : 'http://www') + '.google-analytics.com/ga.js'; var s = document.getElementsByTagName('script')[0]; s.parentNode.insertBefore(ga, s); })(); Skip to main content Preprints Collections Wiley Open Research IET Open Research Ecological Society of Japan All Collections About About Authorea FAQs Contact Us Quick Search anywhere Search for preprint articles, keywords, etc. Search Search ADVANCED SEARCH SCROLL Molecular Microbiology This is a preprint and has not been peer reviewed. Data may be preliminary. 5 September 2025 V1 Latest version Share on Characterization of a regulatory network promoting cell fate segregation in the Myxococcus xanthus biofilm Authors : Shelby Kasto and Penelope Higgs 0000-0003-4726-3329 [email protected] Authors Info & Affiliations https://doi.org/10.22541/au.175710079.93897849/v1 Published Molecular Microbiology Version of record Peer review timeline 300 views 176 downloads Contents Abstract Information & Authors Metrics & Citations View Options References Figures Tables Media Share Abstract Most bacterial populations exhibit phenotypic heterogeneity to increase fitness in rapidly changing environmental conditions. Myxococcus xanthus is an environmental bacterium that displays pronounced phenotypic heterogeneity in its complex lifecycle. Under nutrient-limitation, M. xanthus produces a specialized biofilm in which cells segregate into two spatially distinct fates: fruiting bodies filled with spores, and a persister-like peripheral rod population. Little is known about the regulatory mechanisms controlling peripheral rods. To begin to investigate this cell fate segregation mechanism, we focused on the EspAC signaling system which controls accumulation of MrpC, a central transcription factor necessary to induce fruiting body formation. Single-cell reporters and in situ confocal microscopy demonstrated that expression of the esp genes is enriched in the peripheral rods. We identified three transcription factors necessary for espAC transcriptional control: MrpC; FruA, a transcription factor that coordinates sporulation within fruiting bodies; and the xenobiotic response element, Xre0228. We demonstrate that MrpC directly activates espA and espC ; FruA represses espC, but not espA ; and Xre0228 activates espA but represses espC . These genetic interactions fit common network motifs that promote or stabilize phenotypic heterogeneity. We propose a model by which cell fate segregation is directed, stabilized and tuned to environmental conditions. Introduction To persist in changing environmental conditions, many microbial species exhibit phenotypic heterogeneity, in which clonally-derived (i.e. genetically-uniform) bacterial populations segregate into two or more phenotypically distinct subpopulations (Ackermann, 2015) . P henotypic heterogeneity can confer a fitness advantage by mediating efficient resource management during production of energetically-expensive shared metabolites (division of labor), or by producing subpopulations that can be pre-adapted to survive rapid adverse environmental changes (bet-hedging) (De Jong et al. , 2011; Gasperotti et al. , 2020; Reyes Ruiz et al. , 2020). Multiple mechanisms have been described that can lead to phenotypic heterogeneity, including genetic rearrangements, epigenetic modification, or genetic regulatory networks (GRNs) (Hallet, 2001; Low et al. , 2001; Smits et al. , 2006; Raj and Van Oudenaarden, 2008; Fraser et al. , 2024) . GRNs are a particularly versatile mechanism to produce distinct cell states that can be rapidly tuned to environmental conditions (Shen-Orr et al. , 2002) . These signal responsive GRNs are built from interconnected network motifs (recurring patterns of regulatory connections) associated with specific attributes (Milo et al. , 2002; Alon, 2007) . For example, feedback loops can transform stochastic gene expression to produce distinct gene expression states (bistability) (Alon, 2007; Park et al. , 2012) , and interconnected feedforward loop motifs can produce ordered gene expression (Mangan and Alon, 2003; Eichenberger et al. , 2004; Alon, 2007; Chepyala et al. , 2016) . Thus, the precise topology of these network motifs can promote developmental programs in which cells segregate into distinct fates (Lopez et al. , 2009; Balázsi et al. , 2011) . Myxococcus xanthus , a Gram-negative soil-dwelling bacterium, has a complex lifecycle with multiple phenotypic states (Velicer and Vos, 2009; Muñoz-Dorado et al. , 2016). In the growth phase of the lifecycle, M. xanthus obtains nutrients by predation on other microbial species. At least two types of cells are produced, “yellow” and “tan”, that exhibit division of labor to produce distinct secondary metabolites necessary for effective predation (Furusawa et al. , 2011; Xiao et al. , 2011; Dziewanowska et al. , 2014; Dong et al. , 2022; Contreras-Moreno et al. , 2024). In the developmental phase of the M. xanthus lifecycle, cells segregate into distinct fates. In the wild-type DZ2 strain (Campos and Zusman, 1975; Dey et al. , 2016), ~80% of the cells undergo lysis, likely via programmed cell death, which is thought to release essential nutrients and/or molecules that support the development of the remaining population (Wireman and Dworkin, 1977; Janssen and Dworkin, 1985; Lee et al. , 2012). Two additional spatially segregated cell fates arise from different developmental trajectories: spore filled fruiting bodies and peripheral rods, which account for ~15% and ~5% of the starting population, respectively (O’Connor and Zusman, 1991a; Julien et al. , 2000; Lee et al. , 2012; Higgs et al. , 2014). Fruiting bodies, a specialized biofilm, are produced after approximately 100,000 cells migrate into haystack-shaped mounds and subsequently differentiate into stress-resistant spores. In contrast, peripheral rods neither aggregate nor sporulate; they instead remain arrayed around fruiting bodies (FBs) (O’Connor and Zusman, 1991a; O’Connor and Zusman, 1991b). Peripheral rods (PRs) are thought to have a low metabolic activity, akin to persister cells (O’Connor and Zusman, 1991b; Bhat et al. , 2014; Whitfield et al. , 2020). These two developmental cell fates can be thought of as a form of bet-hedging, because fruiting bodies allow a proportion of the population to survive harsh environmental conditions (Reichenbach, 1993; Lall et al. , 2024), while peripheral rods have been hypothesized to take advantage of low levels of nutrients (O’Connor and Zusman, 1991b). The developmental program is highly tunable to environmental conditions. For instance, the proportion of cells that are segregated into PRs can be altered by local nutrient levels (O’Connor and Zusman, 1991b). Several signaling systems seem to influence the timing of fruiting body formation (Cho and Zusman, 1999; Rasmussen and Søgaard-Andersen, 2003; Jagadeesan et al. , 2009; Glaser and Higgs, 2019), and bypass of fruiting bodies to form rapid spores can be induced in the presence of distinct environmental insults (O’Connor and Zusman, 1997; Marcos-Torres et al. , 2020). A signal-responsive GRN has been well defined for the production fruiting bodies (Kroos, 2017). Two important developmental transcription factors, MrpC and FruA, are central to this GRN, and both are necessary for aggregation and sporulation (Ogawa et al. , 1996; Sun and Shi, 2001). MrpC, a member of the Crp/Fnr transcriptional regulator family, is a global regulator, controlling expression of hundreds of genes throughout the developmental program (Sun and Shi, 2001; Robinson et al. , 2014). Importantly, MrpC activates expression of FruA (Ueki and Inouye, 2003). FruA is an atypical response regulator of the two-component signal transduction family: it lacks key conserved residues required for activation by phosphorylation, no cognate histidine kinase has been identified, and its DNA-binding activity is unaffected by exogenous phosphodonors (Ogawa et al. , 1996; Mittal and Kroos, 2009a). Instead, FruA is thought to be post-translationally activated in response to C-signaling, a short-range cell-cell contact-dependent signaling mechanism (Ellehauge et al. , 1998; Lobedanz and Søgaard-Andersen, 2003; Saha et al. , 2019). The C-signal, a cell surface-associated fragment of the CsgA protein, helps coordinate gene expression in aggregates by relaying information about local cell density (Lobedanz and Søgaard-Andersen, 2003; Gómez-Santos et al. , 2019; Hoang et al. , 2021), but the exact mechanism of FruA activation by the C-signal remains unclear. MrpC and FruA can modulate target gene expression independently, or together via cooperative- or counter- regulation (Nariya and Inouye, 2006; Viswanathan et al. , 2007; Mittal and Kroos, 2009a; Mittal and Kroos, 2009b; Lee et al. , 2011; A. Farrugia et al. , 2025). The regulatory mechanisms governing peripheral rod differentiation are poorly understood. However, it is known that accumulation of several key regulatory proteins necessary to induce fruiting bodies, including MrpC, FruA, and CsgA, are underrepresented in cells in the peripheral rod population (Julien et al. , 2000; Lee et al. , 2012; McLaughlin and Higgs, 2023). Since FruA and CsgA are induced by MrpC (Ueki and Inouye, 2003; Higgs and Mataczynski, unpublished), it is likely that it is the reduced MrpC accumulation that is key to production of peripheral rods. MrpC is subject to multiple levels of transcriptional (Sun and Shi, 2001; McLaughlin et al. , 2018; Marcos-Torres et al. , 2020) and post-translational regulation (Nariya and Inouye, 2006; Rajagopalan and Kroos, 2014; Feeley et al. , 2019), including regulated proteolysis induced by the EspAC signaling system (Schramm et al. , 2012). The EspAC signaling system consists of two hybrid histidine kinases, EspA and EspC, that participate in a cross-phosphorylation signaling mechanism (Cho and Zusman, 1999; Lee et al. , 2005; Higgs et al. , 2008; Schramm et al. , 2012). No cognate response regulator(s) is known, but EspA-dependent phosphorylation of both EspA and EspC receiver domains is necessary to trigger an unknown protease to degrade MrpC (Schramm et al. , 2012). In espA and/or espC deletion strains, MrpC accumulates more rapidly than in wildtype, resulting in premature fruiting body production. Furthermore, these mutants produce spores within the peripheral rod population (Cho and Zusman, 1999; Schramm et al. , 2012). The later observation suggests that appropriate accumulation of MrpC is important for proper cell fate segregation and hints that EspAC may control MrpC accumulation differently within FB and PR developmental trajectories. To examine the hypothesis that EspAC are necessary to promote distinct levels of MrpC accumulation in peripheral rods vs fruiting bodies, we first examined whether the genes were expressed equivalently in the two developmental subpopulations. Using single-cell in situ fluorescent reporters, we demonstrated that expression of espA and espC is detected in both aggregating and peripheral rod populations but is enriched in the peripheral rod population. As little is known about the trans factors controlling esp gene expression, DNA affinity chromatography was used to capture potential regulators from developmental cell lysates. Several regulatory proteins were identified, including MrpC; FruA; the xenobiotic response regulator, Xre0228; and SigA (RpoD). Characterization of the roles of MrpC, FruA and Xre0228 on esp gene expression reveals a surprisingly complex regulatory network that may induce or reinforce subpopulation-specific MrpC accumulation and likely tunes phenotypic heterogeneity within the population to environmental conditions. espA and espC expression is enriched in the peripheral rod population We have previously demonstrated that EspAC signaling system controls accumulation of MrpC, that espAC mutants aggregate prematurely, and some cells within the peripheral rod population sporulate inappropriately (Cho and Zusman, 1999; Higgs et al. , 2008; Schramm et al. , 2012) ( Fig. 1A and B ). Since MrpC is less abundant in the peripheral rod population relative to the aggregating population in our wild type strain DZ2 (Lee et al. , 2012; McLaughlin and Higgs, 2023), we set out here to test the hypothesis that EspAC may control accumulation of MrpC differently within the two developmental subpopulations. We first set out to determine whether expression of espA and espC were differentially regulated within the two populations by examining in situ single-cell espA and espC expression patterns in peripheral rod versus fruiting body populations using fluorescent reporters. An espA fluorescent reporter was constructed by fusing the upstream region (-500 to +21 bp relative to the start codon) to the second codon of mCherry , hereafter P espA WT mCh (Fig. 1C ). Attempts to generate a similar reporter for espC failed likely due to difficulty with homologous recombination (data not shown). However, we were able to construct an espC transcriptional reporter by inserting the tdTomato gene downstream of the espC stop codon. To ensure efficient translation while preserving transcriptional regulation of espC , the pilA ribosome binding site (rbs pilA ) (Wu et al. , 1997) was included upstream of tdTomato to generate espC -rbs pilA - tdTomato (hereafter espC-tdTom ) ( Fig. 1C ). Additionally, each strain contained a P vanillate - mNeonGreen construct (McLaughlin and Higgs, 2023) integrated at an exgenous site, such that every cell could be induced to produce green fluorescence. No phenotypic differences were observed in the reporter-bearing strains relative to the wild type parent cells (data not shown). Confocal laser scanning microscopy (CLSM) was used to detect single-cell fluorescence in the peripheral rod versus fruiting body populations for each strain at 30, 36, and 42 hours of development in submerged culture conditions ( Fig. 1D ). To control for non-specific variation in reporter signal, fluorescence detected from P espA WT mCh or espC-tdTom was first normalized to mNeonGreen fluorescence to generate a red-to-green (RG) ratio (McLaughlin and Higgs, 2023). Average single-cell RG ratios were calculated from at least three distinct fields of peripheral rods (PR) or fruiting bodies (FB) in two independent biological replicates. Enrichment in one population was calculated by dividing the average peripheral rod RG ratio (PR RG ) by the average fruiting body RG ratio (FB RG ) (PR RG /FB RG ratio). Values approximately equal to 1 indicate equivalent expression in both populations, whereas > 1 or < 1 indicate enrichment in the peripheral rod or fruiting body populations, respectively. In the case of P espA WT mCh, we observed average PR RG /FB RG ratios of 1.7, 1.6, and 1.4 at 30, 36, and 42 hours of development, respectively, suggesting espA was consistently enriched in the peripheral rod population ( Fig. 1E ). Likewise, espC-tdTom PR RG /FB RG ratios were 1.2, 1.5, and 1.2 at 30, 36, and 42 hours of development, respectively (Fig. 1E ), suggesting espC was also enriched in the PR population but to a lesser extent. Thus, PR enrichment of espA and espC expression suggests a mechanism by which EspAC may control accumulation of MrpC differently within the two subpopulations. Specifically, PR-enriched EspAC likely contributes to more efficient turnover of MrpC, leading to reduced MrpC levels (Lee et al. , 2012; McLaughlin et al. , 2018; McLaughlin and Higgs, 2023) and perhaps stabilizing the distinct developmental trajectories of peripheral rods versus aggregating cells. RpoD, MrpC, and Xre0228 bind to both espA and espC promoter regions in vitro To begin to understand the basis of differential regulation in espA and espC between the PR and FB subpopulations, we explored the regulatory elements controlling their respective expression. espA and espC genes are separated by >7.0 x 10 6 bp and are therefore expressed from independent promoters (Fig. 2) . The espA start codon is consistent with original annotation from the sequenced M. xanthus strain DK1622 (Goldman et al. , 2006), while the espC start codon was adjusted to the GTG 36 bp downstream of the original annotation based on the transcriptional start site (TSS) mapped by Cap-Seq data (Kuzmich et al. , 2022). Accordingly, all reported positions of espC cis -elements are numbered relative to the updated start codon ( Fig. 2 ). To identify potential regulators acting at these promoters, we performed affinity chromatography using probes encompassing either the espA or espC putative promoter regions (-500 bp to +100 bp and -539 bp to +61 bp, respectively ( Fig. 2, red boxes ). A probe containing an internal region of mrpC (+178 to 688 bp) was used as a control for non-specific DNA binding . These probes were each incubated with wild type strain protein lysates generated from cells developing for 19 hours under submerged culture conditions. Captured proteins were identified by mass spectrometry, and proteins with an average enrichment of ≥ 2-fold relative to the control from two independent biological replicates were considered putative regulators of interest. These analyses identified the sigma factor SigA (RpoD; σ 70 ) (Biran and Kroos, 1997), and transcription factors MrpC, and Xre0228 (Furusawa et al. , 2011) as binding to both espA and espC probes, while FruA was only recovered from the espC probe ( Table 1 ). SigA was the only sigma factor significantly enriched in any of our replicates, however analysis of the putative espA and espC promoter regions showed sequences with poor fit to the canonical -10 or -35 consensus sequences ( Fig. S1 ). Assuming that the SigA interaction is valid, this observation suggested that its binding is likely weak and the additional regulatory proteins are needed to facilitate transcriptional initiation (Lawson et al. , 2004). Thus, we focused on characterization of the transcriptional factors identified. Expression of espA and espC is induced by MrpC Interestingly, MrpC was identified with an enrichment of approximately 50-fold ( espA ) and 1500-fold ( espC ) relative to the negative control ( Table 1) . To confirm MrpC binding, we performed an electrophoretic mobility shift assay (EMSA) with Cy3-labeled versions of either the espA or the espC probes that were used for DNA affinity chromatography. The 600 bp probes were each incubated with 5-20 pmol of purified affinity tagged MrpC (His 6 -MrpC) (McLaughlin et al. , 2018), and the reactions were resolved on a 5% non-denaturing gel (Fig. 3). One ( espA ) and two ( espC ) distinct shifts were observed, suggesting that MrpC directly binds to one site within the espA probe and two sites within the espC probe. MrpC has previously been shown to function as either an activator or repressor of target genes in vivo (Ueki and Inouye, 2003; McLaughlin et al. , 2018; A. Farrugia et al. , 2025). To examine how MrpC affects esp gene regulation, we first examined the EspA and EspC protein levels in wild type and ∆ mrpC strains. Developmental cell lystates were harvested from each strain and subject to immunoblot analysis with anti-EspA or anti-EspC antibodies. Consistent with previous observations (Schramm et al. , 2012), in the wild type strain EspA and EspC accumulated by 12 hours of development and continued to accumulate over the course of 36 hours. Both proteins were highly reduced in the ∆ mrpC strain: in 24 hour developmental lysates, EspA and EspC were approximately 60- and 6- fold reduced, respectively (Fig. 4A and 4B) . Consistently, qRT-PCR analyses of cDNA generated from cells at 24 hours of development indicated espA and espC transcripts were significantly reduced 6-fold (p = 0.02) and 3-fold (p = 0.008), respectively, in the ∆ mrpC strain relative to wild type (Fig. 4C). Together, these data suggest that MrpC directly activates espA and espC expression during development. MrpC binds to one site in the espA promoter region and two sites in the espC promoter region MrpC ChIP-Seq analysis performed at the aggregation stage of development (18 hours post-starvation in the DK1622 M. xanthus wild type strain), indicates an occupancy peak corresponding to -93 bp relative to the espA start codon (Robinson et al. , 2014). We used the previously defined MrpC consensus sequence (Nariya and Inouye, 2006; Robinson et al. , 2014) to identify two possible binding sites located -103 bp and -144 bp from the start codon, with 93% and 87% sequence identity to the MrpC consensus sequence, respectively ( Fig. 5A ). To determine if MrpC binds these sites, EMSAs were performed using 30 bp Cy5-labeled probes encompassing either the -103 bp site ( espA -M1) or the -144 bp site ( espA -M2) (Fig. 5A and 5B top). An MrpC-dependent shift of espA -M1 could be observed using 10-40 pmol MrpC. This binding was specific because it was not observed in the presence of 109-fold excess unlabeled espA -M1 probe. In contrast, no obvious shift was observed for espA -M2, suggesting MrpC specifically binds to only espA -M1, consistent with the single shift observed for the extended probe (Fig. 3). Within the espC probe, we could identify two possible MrpC consensus sequences at -99 bp ( espC -M1) and -453 bp ( espC -M2) with 93% and 100% sequence identity to the MrpC consensus sequence, respectively (Fig. 5A). EMSA analysis using a 30 bp espC -M1 probe indicated a specific shift was detected with 20 and 40 pmol of purified MrpC ( Fig. 5B, bottom) . For the 30 bp espC -M2 probe, a clear shift could be detected with at least 10 pmol MrpC, suggesting an at least 2-fold higher affinity for espC -M2 relative to espC -M1 (Fig. 5B, bottom) . The higher affinity binding is consistent with a perfect match between espC -M2 and the MrpC consensus sequence. Addition of 109-fold excess unlabelled probe to the 40 pmol MrpC reaction resulted a large reduction (~75%) of the shifted band; this result suggests MrpC binding is specific but has higher affinity for the espC -M2 probe relative to espC -M1. Thus, MrpC directly activates expression of espC through binding of at least one of the two identified MrpC binding sites. MrpC acitvates espA from a single CRP-Class I-like binding site To examine the in vivo role of the esp MrpC binding sites, we first examined whether espA -M1 is the sole site necessary for MrpC-dependent activation of espA expression. For this purpose, a reporter was generated where the first 3 bases of the espA- M1 MrpC consensus (Fig. 5A) were mutated to GAA (P espA -M1* mCh ). EMSA analysis indicated this mutation abolished MrpC binding in vitro (Fig. S2A) . Integration of the reporter construct in the wild type strain had no effect the developmental phentotype (data not shown). We then compared total population fluorescence accumulation of P espA -M1* mCh to that of the wildtype promoter (P espA WT mCh). Both strains were induced to develop in 96 well plates, and a microplate reader was used to detect mCherry fluorescence every 60 min from 0 to 48 hrs. In the wild type backgrounds, P espA WT mCh fluorescence increased an average of 3-fold post-starvation (Fig. 6A, top) , whereas P espA -M1* mCh fluorescence was significantly reduced 2.4-fold (p = 0.0052) and 1.9- fold (p = 0.0053) relative to P espA WT mCh at 24 and 48 hours of development, respectively (Fig. 6A, top) . A similar pattern was observed if P espA WT mCh reporter was analyzed in the ∆ mrpC background (Fig. S3A). Because the total population expression patterns were not able to be normalized to cell number, we next confirmed these expression patterns by assaying single-cell fluorescence at 0, 24, and 48 hours of development. Average single-cell fluorescence values produced similar patterns to that observed in the total population analysis, except P espA -M1* mCh fluorescence was reduced 16-fold (p = 0.0123) at 24 hours and 7-fold (p = 0.0062) at 48 hours relative to P espA WT mCh (Fig. 6A, bottom) . Together, these results indicate that espA -M1 is both required and sufficient for direct activation of espA gene expression by MrpC. MrpC primarily regulates espC expression from a CRP-Class I-like site To examine the contribution of espC -M1 and espC -M2 to activation of espC expression in vivo , it was necessary to construct an additonal espC expression reporter which could be integrated at the ectopic 1.38 kb integration region (Iniesta et al. , 2012). We first demonstrated that a construct containing -536 bp and the espC coding region complemented a ∆ espC mutant with respect to developmental phenotype and EspC protein accumulation (data not shown). We therefore generated an espC expression reporter that contained -536 bp to +39 bp espC fused to the tdTomato fluorescent gene (Hoang et al. , 2021). Analysis of tdTomato fluorescence in an automated microplate reader over the course of 48 hours of development in submerged culture showed the expected post-starvation increase in expression (Schramm et al. , 2012) ( Fig. 6B, top ). To examine whether both M1 and M2 binding sites contribute to espC regulation, each site was mutated independently or in combination to generate P espC -M1* tdTom , P espC -M2* tdTom or P espC -M1*/M2* tdTom reporter constructs. To generate the mutations, the first four nucleotides of the MrpC site consensus sequences (Fig. 5A) were subsituted to GAAA. EMSA analyses indicated this mutation completely abolished detectable MrpC binding to espC -M1 and espC -M2 in vitro (Fig. S2B). Integration of the reporter constructs in the wild type strain had no effect on the developmental phentotype (data not shown). Analysis of total population fluorescence indicated that in the wild type background, P espC WT tdTom fluorescence increased during development, but did not significantly increase in the ∆ mrpC mutant background (Fig. S3B). At 24 hours of development, fluorescence from P espC -M1* tdTom and P espC - M1*/M2* tdTom were significantly reduced by 1.4-fold (p = 0.035) and 1.5- fold (p = 0.018), respectively. In contrast, fluorescence from P espC -M2* tdTom was only slightly (1.3-fold) reduced compared to the parent, which was not considered statistically signifcant (p = 0.22) (Fig. 6B, top) . By 48 hours, all three mutant reporters were reduced ≥1.5-fold relative to the wildtype reporter, but were not considered statistically significant (P espC -M1* p = 0.12, P espC -M2* p = 0.12, P espC - M1*/M2* p = 0.07) (Fig. 6B, top) . In contrast to reporter accumulation in the total population where tdTom appeared to increase throughout development, single-cell analysis revealed a sharp decrease in tdTom accumulation between 24 and 48 hours (Fig. 6B, bottom) . This discrepancy is likely due to the release of tdTomato into the well as a result of developmental cell lysis (Wireman and Dworkin, 1977; Janssen and Dworkin, 1985; Lee et al. , 2012). In single cells at 24 hours of development, tdTom from both P espC -M1* and P espC - M1*/M2* displayed the largest reduction in expression relative to the wild type reporter, with levels decreased by 2.1-fold and 1.9-fold (p = 0.02 and p = 0.0051), respectively (Fig. 6B, bottom) . In contrast, P espC -M2* tdTom expression showed only a modest 1.3-fold reduction which was not considered significant (p = 0.17). This pattern persisted at 48 hours with tdTom from P espC -M1* , P espC - M1*/M2* , and P espC -M2* reduced by 1.6-fold (ns, p = 0.075), 1.8-fold (p = 0.023), and 1.4-fold (ns, p = 0.25), respectively, compared to P espC WT (Fig. 6B, bottom) . These analyses suggests that espC -M1 is the primary site mediating positive regulation of espC expression, while espC -M2 plays a minor role, at least under our standard developmental conditions in the lab. FruA regulates espC but not espA FruA was recovered in the proteins binding to the espC probe, but not the espA probe ( Table 1 ). To assess a potential regulatory effect of FruA on espC expression, we first examined EspC protein levels in a fruA mutant. Developmental cell lysates were harvested from the wild type and the fruA mutant and subject to immunoblot analysis with anti-EspC antibodies. Interestingly, EspC protein displayed a significant 1.5 ± 0.12-fold (p = 0.0027) increase in accumulation levels in the fruA mutant relative to wild type as measured at 24 hours of development (Fig. 7A) . In contrast, anti-EspA immunoblots revealed no significant difference in EspA protein levels relative to wild type (0.89 ± 0.05-fold, p = 0.07, at 24 hours) (Fig. S4A). To determine if this pattern was reflected at the transcriptional level, qRT-PCR was used to examine espC gene expression in a fruA mutant. RNA was harvested from 24 hour developmental cells, converted to cDNA, and subject to qPCR. Consistently, espC transcripts were significantly higher in the fruA strain with a 1.85 ± 0.14-fold (p = 0.0052) increase, relative to wild type (Fig. 7B). espA transcript levels in the fruA mutant were not significantly altered compared to wild type (data not shown). Consistently, a putative FruA binding site with 100% sequence identity to the previously defined GGGYR-n 4-6 -YGGG FruA consensus sequence (Yoder-Himes and Kroos, 2006; Viswanathan et al. , 2007) could be identified centered at +1 bp relative to the espC start codon ( espC -F1) (Fig. 2 and Fig. S4B) ; no convincing FruA binding sequence could be identified in the espA promoter region. To test if FruA directly bound the espC promoter region, affinity tagged full length FruA (FruA FL His 6 ) or affinity tagged DNA binding domain of FruA (FruA DBD His 8 ) were purified and utilized with EMSA analysis with the 600 bp espC probe . We observed diffuse shifts in the presence of FruA DBD and FruA FL (Fig. 7C) . If FruA and MrpC were added together, migration of the two distinct bands observed for MrpC binding were both retarded (Fig. 7C) . These observations suggested that FruA and MrpC could simultaneously bind to the espC promoter region. However, FruA FL binding to a 30 bp probe consisting of the predicted FruA binding site ( espC -F1) could not be detected by EMSA analysis, although efficent binding to a fmgD positive control probe (Yoder-Himes and Kroos, 2006; Mittal and Kroos, 2009b) was detected (Fig S4C) . Thus, it is not currently clear whether FruA represses espC expression by binding to the espC -F1 site in vivo . Collectively, however, our data suggests that FruA acts to repress espC expression, but does not regulate espA expression. Xre0228 induces espA and represses espC expression The xenobiotic transcriptional regulator homolog, Xre0228, was recovered from both the espA and espC promoter probe pull down assays (Table 1) . Xre0228 has been previously characterized as important for regulation of the locus necessary for synthesis of the secondary metabolite/yellow pigment DKxanthene in M. xanthus (Dziewanowska et al. , 2014). In the DZ2 background under standard laboratory conditions, the ∆ xre0228 mutant does not have an obvious developmental phenotype (Kasto and Higgs, unpublished) and no significant effect on espA and espC expression was observed (data not shown). Xre0228 is likely not expressed in all cells in the population (Furusawa et al. , 2011; Dziewanowska et al. , 2014), and Xre0228-dependent effects on esp gene expression may be difficult to detect. We therefore forced xre0228 expression in all cells by placing it under the expression of a vanillate-inducible promoter, generating P vanillate xre0228 ( xre0228 ++ ). This construct was integrated at an exogenous locus in strains bearing either the espA or espC reporters, generating xre0228 ++ P espA WT mCh or xre0228 ++ espC - tdTom strains, respectively. These strains were induced to develop in 96 well plates in the absence or presence of vanillate, and total population fluorescence was detected in a microplate reader over the course of 48 hours. Fluorescence detected in the uninduced xre0228 ++ on P espA WT mCh strain was similar to that in the wild type background throughout development (Fig. 8A, top). For the first ~24 hours of development, mCherry fluorescence was similar under uninduced and induced conditions; thereafter, however, fluorescence increased in the induced strain. Specifically, the fluorescence captured from the induced strain was an average 1.3-fold higher at 36 (p = 0.125) and 48 (p = 0.086) hours of development (Fig. 8A, top). This expression pattern was confirmed by analysis of single-cell fluorescence. mCherry under inducing conditions was upregulated ~1.5-fold at 24 (p = 0.078) and 48 (p = 0.13) hours relative to the uninduced (Fig. 8A, bottom) . These results suggested that Xre0228 may act to upregulate espA during development. The same approach was used to examine the effect of inducing xre0228 ++ on espC - tdTom reporter expression. U nder inducing conditions in the total population, reporter fluorescence was slightly reduced 1.2- (p = 0.003) and 1.3-fold (p = 0.04) at 36 and 48 hours of development , respectively, relative to the uninduced strain (Fig. 8B, top) . This trend was magnified in the individual cell analysis where tdTomato fluorescence was 2- (p = 0.015) and 3-fold (p = 0.057) reduced at 24 and 48 hours, respectively, relative to the uninduced (Fig. 8B, bottom) . These data suggest that during development, Xre0228 may act to repress expression of espC. Interestingly, Xre0228 exerts opposite effects on espA versus espC expression. In contrast to MrpC and FruA, which play major roles in the core developmental program, we speculate Xre0228 may serve to fine-tune the espA / espC balance in reponse to environmental cues not necessarily present in our standard developmental assays. Discussion During the M. xanthus developmental program, cells that do not undergo cell lysis segregate into peripheral rods or fruiting bodies. We have long postulated that MrpC plays a key role in this process, because threshold levels of MrpC have been proposed to trigger distinct developmental transitions: sub-threshold MrpC levels are associated with peripheral rods, intermediate levels of MrpC initiates aggregation into fruiting bodies, and higher levels of MrpC promote commitment to sporulation (Lee et al. , 2012; Rajagopalan and Kroos, 2014; Marcos-Torres et al. , 2020; McLaughlin and Higgs, 2023). MrpC does not appear to require a native ligand, but its accumulation is subject to complex transcriptional (Sun and Shi, 2001; Nariya and Inouye, 2006; McLaughlin et al. , 2018; Marcos-Torres et al. , 2020), and post-translational control (Nariya and Inouye, 2006; Schramm et al. , 2012; Rajagopalan and Kroos, 2014; Feeley et al. , 2019). Perturbation of these regulatory mechanisms can influence cell fate segreation (Cho and Zusman, 1999; Higgs et al. , 2008), induce novel fates (McLaughlin and Higgs, 2023), and even cause loss of developmental robustness (Feeley et al. , 2019). Here we focus on the EspAC signaling system to reveal one mechanism by which MrpC can be induced to accumulate to different levels within each of the developmental subpopulations. First, we find that expression of espA and esp C , and therefore likely their protein products, are enriched in peripheral rod cells relative to cells that have migrated into aggregation centers. As we have previously observed that MrpC levels are depleted in the peripheral rod population (Lee et al. , 2012; McLaughlin et al. , 2018; McLaughlin and Higgs, 2023), we suggest that EspAC contribute to keeping MrpC levels low in this population. MrpC sits near the top of a regulatory hierarchy (Kroos, 2017), i.e. it is required for expression of at least two other major regulators (CsgA and FruA) (Ueki and Inouye, 2003; Higgs and Mataczynski, unpublished) that are essential to induce cells to aggregate into fruiting bodies (Shimkets and Rafiee, 1990; Ogawa et al. , 1996). Thus, low levels of MrpC are likely insufficient to induce efficient FruA and CsgA accumulation, aggregation is not stimulated, and cells are stabilized in the peripheral rod developmental trajectory. Consistently, in the espAC mutant, the peripheral rod population appears to be depleted and some of the peripheral rods have sporulated ( Fig. 1B ). We suggest that in the espAC mutant, MrpC is allowed to accumulate and additional cells are switched to the fruiting body trajectory such that aggregates are observed earlier than in the wild type. We propose that EspAC also regulate appropriate accumulation of MrpC in cells in the fruiting body developmental trajectory, but that the presumably lower levels of EspAC induced in this population allow less efficient MrpC turnover. The result is a gradual accumulation of MrpC which leads to larger, well-organized aggregates, before sporulation is induced. In the espAC mutant, we suggest the aggregates are smaller and less well organized, because a rapid accumulation of MrpC allows cells to begin to sporulate before they have finished aggregating ( Fig. 1A ). The regulatory network which controls espA and espC expression helps explain how EspAC can be expressed to different levels in the two populations. First, we demonstrated that MrpC is required to induce expression of espA and espC. Since EspAC mediates MrpC turnover, this constitutes a composite negative feedback network motif containing a relatively slow transcriptional regulation associated with rapid protein degradation (Alon, 2007). For example, in E. coli and Salmonella enterica , the stress reponse sigma factor, RpoS, regulates its own turnover by inducing expression of an adaptor protein, SprE, that targets RpoS to the ClpXP protease (Gibson and Silhavy, 2000; Ruiz et al. , 2001; Bouillet et al. , 2024). The RpoS/SprE feedback loop has been proposed to maintain low levels of RpoS when it is not needed, because overaccumulation can be potentially fatal to the cells by diverting them into a SOS response (Zambrano et al. , 1993; Pratt and Silhavy, 1998; Merrikh et al. , 2009). In M. xanthus , it is possible that this type of motif could contribute to stable reduced MrpC levels in the peripheral rod population, which is rapidly reversible if EspAC are inactivated, perhaps in response to environmental stimuli (Fig. 9A). A second set of regulatory interactions involving MrpC, FruA, and espC may explain how distinct MrpC levels can be controlled in fruiting bodies. First, we observed that FruA slightly (~1.5-fold) represses EspC ( Fig. 7 ). Consistently, upregulation of espC in a fruA mutant was observed in RNA-Seq studies with an average increase of et al. , 2025). Despite a candidate FruA binding site (100% consensus match; Fig. S4B and Fig. 2 ) in a position consistent with repression (Rojo, 1999) ( espC -F1, +1 bp relative to start codon), we could not verify direct FruA binding to this site by EMSA analysis. We could, however, detect in vitro binding of FruA to the fmgD binding site ( Fig. S4C ). We cannot rule out the possiblility that the FruA consensus sequence requires refinement, because the current FruA consensus has been derrived from a limited set of promoters (Yoder-Himes and Kroos, 2006; Viswanathan et al., 2007), and the fmgD binding site is only an 80% match to this sequence ( Fig. S4B ). However, we consider it likely that unactivated FruA simply binds with higher affinity to fmgD in vitro . fmgD expression is abolished in fruA or csgA mutants (Kroos and Kaiser, 1987; Lee et al. , 2011). In contrast, the modest FruA-dependent repression of espC may require substantialy higher levels of activated FruA, which we cannot recapitulate in vitro . These observations raise the possibility that FruA repression of espC may operate primarily within fruiting bodies where activated FruA is concentrated as a result of C-signaling (Ellehauge et al. , 1998; Saha et al. , 2019; Hoang et al. , 2021) ( Fig. 9B ). The interactions observed between MrpC, FruA and espC fit a common type I incoherent feed-forward loop (I1-FFL) motif (Milo et al. , 2002; Mangan and Alon, 2003). A well characterized example of a I1-FFL is observed in the E. coli galactose utilization ( gal) system (Weldemichael et al. , 2022). Transcriptional activation of the galETK operon is regulated by CRP. CRP also activates galS, and high levels of GalS bind to and represses galETK (Weickert and Adhya, 1993). The gal I1-FFL has been shown to lead to accelerated response times and generation of a tunable non-monotonic response (Mangan et al. , 2006; Kaplan et al. , 2008). We propose an analogous I1-FFL operates with MrpC as the primary activator, FruA as the repressor, and espC as the target. As MrpC accumulates, espC and fruA are both expressed, but espC is subsequently repressed as activated FruA accumulates, yielding a fast initial expression of espC and a tunable non-monotonic response. Because activated FruA is C-signal dependent, and C-signal is produced upon cell-cell contact (Ellehauge et al. , 1998; Lobedanz and Søgaard-Andersen, 2003; Saha et al. , 2019; Hoang et al. , 2021), espC is likely primarly repressed in aggregating cells. As EspA and EspC are both required to induce MrpC turnover (Schramm et al. , 2012), repression of EspC prevents efficient turnover of MrpC, allowing MrpC to accumulate and stiumulate the fruiting body trajectory of the developmental program ( Fig. 9B ). Our detailed analysis of MrpC-dependent activation of espA and espC, suggests MrpC activates both genes through a promoter proximal site, approximately 103 bp and 99 bp upstream from the start codon, respectively. The position of these binding sites is consistent with a mechanism defined for CRP class I activation in Escherichia coli in which CRP directly contacts the αCTD subunit of RNA polymerase to activate transcription (Ushida and Aiba, 1990; Gaston et al. , 1990; Zhou et al. , 2014). It was recently demontrated that genes strongly activated by MrpC are associated with MrpC Chip-seq occupancy in a region of 60-105 bp upstream of the TSS (Robinson et al. , 2014; Kuzmich et al. , 2022; A. Farrugia et al. , 2025). Interestingly, espC is additionally regulated by a distal MrpC binding site 453 bp upstream of the start codon ( Fig. 2 ), suggesting that MrpC may participate in a long range regulatory mechanism. It has been previously reported that CRP-family regulators can act at distal sites. For example, Vibrio cholerae CRP contibutes to control of the ompT gene from both proximal and distal binding sites. In this example, CRP activates expression from the distal site (via DNA looping) but represses expression from the proximal sites; complex signals related to growth phase govern the selectivity for each site (Schleif, 1992; Li et al. , 2002). In M. xanthus , MrpC has been proposed to mediate negative autoregulation through DNA looping to form a closed promoter complex that prevents σ 54 binding (McLaughlin et al. , 2018). Under standard laboratory conditions, mutation of the distal site reproducibly lowered expression of espC reporter activity, although the effect did not reach statistical significance likely owing to reporter noise. Regardless, the impact of the espC -M2 distal element may become more apparent under alternative environmentally-relevant conditions, suggesting it may be a fine tuning mechanism that may also alter EspC levels, relative to EspA, providing an additional mechanim to tune MrpC accumulation in some cells. Surprisingly, we also identied Xre0228 as interacting with both espA and espC promoter regions ( Table 1 ), and demonstrated it has opposite effects on the expression of espA and espC ( Fig. 8 ). Xre0228 is a transcription factor previously characterized for its role in vegetative “yellow/tan” phenoytypic heterogeneity. Xre expression is enriched in yellow cells and is necessary for expression of the yellow pigment DKxanthene (Furusawa et al. , 2011; Dziewanowska et al. , 2014). We could not confirm direct Xre0228 binding to espA or C probes. Despite purification of milligram quantities of affinity tagged Xre0228, the protein was poorly behaved in EMSAs (data not shown), suggesting it may require an interacting partner or activation signal. Additionally, no binding sequence has been identified for Xre0228. Nevertheless, forced expression of Xre0228 revealed an interesting converse regulation where espA expression was induced, while that of espC was repressed, especially in the later stages of development ( Fig. 8 ). We interpret this observation to suggest that Xre0228 is not normally present/active in the later stages of development, such that its effect on espA/C is most readily observed by overexpresison in this period. Unlike MrpC and FruA, which are core developmental regulators, we suggest that Xre0228 primarily functions to fine-tune EspA/C ratios likely in reponse to environmental signals. First, a ∆ xre0228 mutant displayed no discernable developmental phenotype, and the esp reporters were not signifcantly affected in the ∆ xre0228 background under standard laboratory conditions (data not shown). These observations suggest that Xre0228 is likely limited in most cells, only present in a subset of the population, and/or only significantly activated under non-laboratory conditions. We propose that activation of Xre0228, in reponse to environmental conditions, generates an imbalance of EspA/EspC. Because EspA and EspC act cooperatively, skewing their stoichiometry reduces effective activation of the MrpC targeted proteolysis, allowing MrpC to accumulate to the aggregation threshold and initiating aggregation in additonal cells ( Fig. 9C ). Indeed, the porportion of cells in the peripheral rod population varies depending upon the level of nutrients in the environment (O’Connor and Zusman, 1991b), raising the intriging possibility that Xre0228 may be a factor that helps tune the PR / FB segregation balance in reponse to environmental conditions. We are currently exploring the hypothesis that espAC regulation is linked, through Xre0228, to the proportion of yellow and tan cells that enter into the developmental program. Intrigingly, the yellow tan ratio can be skewed by the availability of iron (Dziewanowska et al. , 2014), population density, and nutrient availability (Burchard et al. , 1977). It is noteworthy that regulation of espC appears to be subject to additional layers of regulation over that of espA ; while espA is induced by MrpC (through CRP class I activation) and likely by Xre0228, MrpC-dependent regulation of espC involves CRP class I activation and an additional distal mechanism, and espC is additionally repressed by FruA and Xre0228. Interestingly, in the alternate DK1622 M. xanthus background (Wall et al. , 1999), deletion of espC does not produce an obvious phenotype (Shi et al. , 2008), and the peripheral rod population may not display reduced MrpC accumulation (Kuzmich et al. , 2022). The DK1622 developmental program occurs ~12 hrs more rapidly than that of DZ2 (Lee et al. , 2012), and the population contains significantly more yellow cells than that of DZ2 (Laue and Gill, 1994; S. Kasto and P.I. Higgs, unpublished results). We speculate that the EspC regulatory mechanism may be perturbed in the DK1622 background. An intriguing hypothesis is that DK1622 lacks a subset of peripheral rods which are influenced by EspC in the DZ2 background; less peripheral rods likely explains why DK1622 produces visible aggregates earlier than DZ2. We have long speculated that there may be distinct subpopulations of peripheral rods that may be influenced by different signaling systems. For instance, disruption of the Red (Jagadeesan et al. , 2009), SinK (Glaser and Higgs, 2019), or TodK (Rasmussen and Søgaard-Andersen, 2003) signaling systems each influence MrpC activity and cause innappropriate sporulation of some peripheral rods (B. Lee, C. Mataczynksi, and P.I. Higgs, unpublished results). We have demonstrated that double mutants of these systems produce strinkingly additive early development phenotypes (Lee, 2009), and a ∆ red todK ∆ sinK ∆ espAC mutant completely lacks peripheral rods (Lee, 2009; Lall et al. , 2024). We provide evidence for a genetic regulatory framework that influences cell fate segregation via the EspAC signaling system. While we have identified some mechanisms by which cell fate segregation can be tuned to environmental conditions, and perhaps to the yellow/tan cell states, it is obvious that this framework captures only a subset of the regulatory mechanims involved. For instance, the activity of EspA and EspC are certainly regulated by, as yet unknown, ligands/stimuli (Cho and Zusman, 1999; Schramm et al. , 2012), and MrpC itself is subject to an extremely complex set of transcriptional and post-translational regulatory mechanisms that are influenced by multiple additional signaling systems. Experimental Procedures Bacterial strains, growth, and development conditions- The bacterial strains and plasmids used in this study are listed in Table 2 . Escherichia coli strains were grown at 37°C in LB broth with shaking at 220 rpm, or on LB agar, each containing 50 µg mL -1 kanamycin or 20 µg mL -1 tetracycline, where necessary (Green and Sambrook, 2012). M. xanthus strains were grown under vegetative conditions at 32°C in CYE broth with shaking at 220 rpm, or on CYE 1.5% agar plates (Campos and Zusman, 1975) supplemented with 100 µg mL -1 kanamycin, 10 µg mL -1 oxytetracycline, and/or 0.5 mM vanillate, where necessary. M. xanthus development was induced in submerged culture conditions (Lee et al. , 2010). Briefly, cells were grown overnight in in CYE broth and diluted to 0.035 A 550 in fresh CYE. 16 mL, 2.1 mL, 0.5 mL, or 0.15 mL of diluted culture were seeded into 100 mm petri dishes, confocal dishes (ibiTreated µ-dishes 35 mm, high (Ibidi)), 24 well plates, or 96 well plates respectively, and incubated at 32°C for 24 hours to form a confluent cell layer. To initiate development, the CYE overlay was replaced with an equivalent volume of MMC (10 mM MOPS pH 7.6, 4 mM MgSO 4 , 2 mM CaCl 2 ) and further incubated at 32°C. Construction of plasmids and strains Construction of plasmids is described in Table S1 and primer sequences used are listed in Table S2 . Protocols for overlap PCR and restriction enzyme/ligation cloning techniques have been previously described in detail (Lee et al. , 2010). For plasmids constructed by Gibson assembly (Avilan, 2023), the respective fragments indicated in Table S1 were designed with a 15-25 bp overlap. Fragment assembly was performed using NEBuilder® HiFi DNA Assembly Master Mix (New England Biolabs) as per manufacturer’s reccomendations. Assembled plasmids were transformed into E. coli strain Top10 and selected with relevant antibiotics. The relevant regions of plasmids were sequenced to confirm the absence of PCR-derived errors. Suicide plasmids were introduced into M. xanthus DZ2 backgrounds by electoporation, as previously described (Lee et al. , 2010). Plasmid integration by homologous recomination was confirmed using PCR; primer sequences used to confirm integration are available upon request. At least three independent M. xanthus clones were tested to confrim identical developmental phenotypes and/or reporter expression profiles. Bioinformatic analyses Percent identity of MrpC or FruA binding sites to the respective consensus sequence was calculated as follows: For sequences of equal length, percent sequence identity = [2L/(L y + L z )] x 100, where L is the number of aligned bp with same identity, L y is the total length of the binding site sequence, and L z is the total length of the consensus sequence. For sequences of unequal length, percent sequence identity = (L/L x )x 100, where L x is the length of the shorter of the two sequences (Stothard, 2000). Candidate binding sites for MrpC and FruA were identified by manually inspecting upstream regions (approximately -500 to +100 bp relative to the start codons) for motifs similar to the previously defined MrpC (Nariya and Inouye, 2006; Robinson et al. , 2014) or FruA (Yoder-Himes and Kroos, 2006; Viswanathan et al. , 2007) consensus sequences. Putative SigA (RpoD; σ⁷⁰) -35 and -10 promoter elements were assigned using the E. coli σ 70 / Bacillus subtilis σ 70 consensus sequences as per (Biran and Kroos, 1997). Calls were retained only when orientation and spacing were plausible for initiation at the nearby espA or espC transcription start sites (TSS) as mapped previously by Cappable-Seq (Kuzmich et al. , 2022). Confocal microscopy For analysis of single-cell espA and espC expression in the developing populations, M. xanthus strains bearing P vanillate - mNeonGreen and either P espA WT mCh or espC-tdTom were diluted 1:9 with an unlabeled wild type strain (DZ2). Strains were induced to develop in confocal dishes (ibiTreated µ-dishes 35 mm, high ) and imaged with a Leica TCS SP8 inverted confocal microscope as described previously (McLaughlin and Higgs, 2023), except as outlined below. mNeonGreen fluorescence was detected using a 488 nm wavelength excitation laser (5% power), a 500-540 nm emission spectra window with 750V gain and 0.34% offset. mCherry fluorescence was detected using a 552 nm wavelength excitation laser (5% power), a 585-630 nm emission spectra window with 933.9V gain and 0.28% offset. tdTomato fluorescence was detected using a 552 nm wavelength excitation laser (5% power), a 577-641 nm emission spectra window with 1100V gain and -0.11% offset. For each replicate of the peripheral rod populations, a minimum of three distinct fields were imaged. For fruiting bodies, 3-5 z-stacks (5 µM steps) were recorded starting from the base. All images used a line average of 4 with 1024 x 1024 resolution. As no significant differences in red-green ratios were observed between the different z-stacks, the data was combined. The final data were compiled from two independent biological replicates. Peripheral rod and fruiting body images were analyzed in ImageJ (Schindelin et al. , 2012) as previously described (McLaughlin and Higgs, 2023). Images containing fruiting bodies were cropped to include only the fruiting body. For fruiting bodies and peripheral rods at least 2500 and 330 cells were imaged for each timepoint in each strain, respectively. Quantification of single cell fluorescence was performed in Image J as described previously (McLaughlin and Higgs, 2023). Briefly, non-specific fluorescence was defined using unlabeled strains. Regions of interest (ROIs) were defined from single cells in the green channel using the threshold values derived from unlabeled control strains and analyze particles function. ROIs were then transferred to the corresponding mCherry or tdTomato channel. Green and red fluorescence was quantified using integrated density. The red-green (RG) ratios were plotted, and outlying points identified by Grubb’s test (p < 0.5) were removed. RG ratios from each population at each time point were averaged. To assay enrichment of expression in either subpopulation, the average peripheral rod RG ratio (PR RG ) was divided by the average fruiting body RG ratio (FB RG ). PR RG /FB RG ratios equal to 1 indicated equivalent expression in both populations, whereas ratios greater than 1 indicate enriched expression in the PRs. For brightfield imaging of peripheral rods, strains were induced to develop in confocal dishes as described above for 72 hours. Images were recorded on the Leica TCS SP8 inverted confocal microscope with a gain of 225 V and all images were cropped and adjusted in ImageJ for brightness/contrast using the same settings. DNA pull down assay The DNA pull down assay was performed as previously described (Volz et al. , 2012), except as outlined below. Briefly, probes were generated by amplifying the putative promoter regions of espA and espC or an intragenic region of mrpC using 5’ biotinylated primers ( Table S2 ) and DZ2 genomic DNA as template. 2 mg of Dynabeads® M-280 Streptavidin (Invitrogen) were transferred to a 2 mL microcentrifuge tube and ~100 pmol DNA probe was bound according to manufactures instructions. Cell lysates were prepared from approximately 10 plates (~3.2 x 10 10 cells) of cells developing for 19 hours under submerged culture in 100mm dishes. Cells were harvested at 4696 x g for 30 minutes at 4°C, the supernatant was removed and cells were processed, and lysates were incubated with DNA probes, washed, and eluted as described previously (Volz et al. , 2012). Eluted proteins were identified via mass spectrometry at the Wayne State University Proteomic Core Facility. Samples in solution containing 0.5 M NaCl, 5% glycerol and buffered with 10 mM Tris, pH=7.5 were acidified by addition of 10% of the original volume of 12% phosphoric acid. Proteins were precipitated with 5 volumes 90% Methanol (MeOH) with 100 mM Triethylammonium bicarbonate (TEAB, Honeywell Fluka cat# 60-044-974) followed by overnight incubation at -20 o C. Precipitates were recovered at 10,000 x g for 5 minutes, washed one time with 80% MeOH/10mM TEAB, air dried, and then resuspended in 50 ul of 40 mM TEAB buffer containing 5 mM DL-Dithiothreitol (DTT, Sigma cat# D5545) and 0.4 ug of trypsin (Promega, V5113) for reduction of cysteine residues and Tryptic digest of proteins. Digestion proceeded for 1 hour at 47 o C followed by 3 hours at 37 o C. Alkylation of cysteine residues was initiated by addition of 15 mM Iodoacetamide (IAA, Sigma cat# I1149), followed by incubation at room temperature for 30 minutes in the dark. The alkylation reaction was stopped by addition of 5mM DTT. Mass spectrometry was performed on a Thermo Vanquish Neo UHPLC chromatography system with an Acclaim PepMap 100 C18 trap, 75um x 2cm and EasySpray PepMap RSLC, 75um x 25cm column (Thermo scientific). LC-MS/MS was performed using Data Dependent Analysis on an Orbitrap Eclipse. MS1 spectra were acquired at 120,000 resolution and MS2 in the ion trap. Data were analyzed using Proteome Discoverer 2.4 searching a Myxococcus xanthus DZ2 database (accession CP070500) downloaded on March 22, 2023 with 7,377 entries. Results were exported to Scaffold 5 for additional analysis. Proteome Discoverer analysis was performed using Sequest NT and Percolator algorithms accepting 2 missed cleavages by trypsin digestion. Carbamidomethylation of cysteine was a fixed modification. Deamidation of asparagine and glutamine, oxidation of methionine and N-terminal acetylation were allowed dynamic modifications. A protein False Discovery Rate (FDR) was set at 0.01 for high confidence matches in both PD and Scaffold analyses and Scaffold required a two peptide minimum. Overexpression and purification of recombinant proteins His 6 -MrpC was purified as described previously (McLaughlin et al. , 2018). To purify FruA DBD -His 8 or FruA FL -His 6, the respective overexpression plasmids were transformed into BL21(λDE3) or GJ1158, respectively. For each culture, 3 resulting colonies were grown overnight in 5 mL cultures and then subcultured 1:100 into 500 mL LB broth containing appropriate antibiotic. Cultures were induced mid-log for 3 hours at 37°C with 1 mM isopropyl 1-thio-β-D-galactopyranoside (IPTG) (BL21(λDE3)) or with 0.3 M NaCl (GJ1158). Cultures were pelleted at 9936 x g for 20 min at 4°C, supernatant was removed, and pellets were stored at -20°C until needed. Pellets were resuspended in lysis buffer (50 mM HEPES, pH 7.4, 500 mM NaCl, 20 mM imidazole) supplemented with 1X c0mplete mini protease inhibitors (Roche) and lysed via FRENCH® Press (SLM Instruments) 3 times at 18,000 psi. Lysates were clarified at 600 x g for 30 min at 4°C. Affinity-tagged proteins were purified by gravity flow at 4°C on 2 mL of 50% slurry Ni-NTA resin (Qiagen) pre-equilibrated with lysis buffer. The resin was washed with 5 column volumes (CV) of wash buffer (50 mM HEPES, pH 7.4, 500 mM NaCl, 20 mM imidazole). Elution was performed in three steps of 3 CV containing 100 mM, 250 mM and then 500 mM imidazole in elution buffer (50 mM HEPES, pH 7.4, 500 mM NaCl). Elution fractions containing peak eluted protein were pooled and dialyzed overnight at 4 °C in 1 L dialysis buffer 1 (50 mM HEPES, pH 7.5, 250 mM NaCl, 1 mM dithiothreitol (DTT) and 10% glycerol (v/v)), followed by overnight 4 °C in 1 L dialysis buffer 2 (dialysis buffer 1, except 100 mM NaCl and 20% glycerol). Dialyzed proteins were aliquoted and stored at -20°C. Electrophoretic mobility shift assays (EMSA) Oligonucleotides used to generate probes are listed in Table S2. For the extended (~600 bp) probes, a Cy3- labeled primer was paired with an unlabeled primer to PCR amplify espA or espC probes using DZ2 genomic DNA as a template. Probes were purified using GeneJET PCR Purification Kit (Thermo Scientific) and quantified by a Nanodrop 2000 spectrophotometer (Thermo Scientific). The method (Jullien and Herman, 2011). Purified His 6 -MrpC was generated and EMSAs performed as previously described (McLaughlin et al. , 2018). Briefly, 5 µL of each reaction containing 50 nM probe and purified His 6 -MrpC or FruA DBD -His 8 or FruA FL -His 6 (as described in each figure legend) was resolved on a non-denaturing, degassed, 5% (extended probes), 8% (FruA) or 10% (MrpC) polyacrylamide gel prepared with 0.5x TBE (45 mM Tris-borate, 1 mM EDTA, pH 8.0) and run at 100V in 0.5x TBE buffer at 4°C. Gels were imaged using either Cy3 or Cy5 settings on an iBright 1500 imaging system (Invitrogen). Immunoblot analysis Protein lysates for immunoblot analysis were prepared from M. xanthus strains developed in 100 mm petri dishes. At the indicated timepoints, overlay media was removed from one plate, 1.5 mL of ice-cold MMC was added, and cells were transferred to a 2 mL microcentrifuge tube. To precipitate proteins, 80% trichloroacetic acid was added to a final concentration of 13% followed by incubation on ice for 15 mins. Protein precipitates were recovered at 17,000 x g for 5 minutes at 4°C, rinsed with 500 µL ice-cold acetone, centrifuged as above, and then rinsed with 1 mL 100mM Tris, pH 8.0. Protein pellets were resuspended in 100 µL of 100 mM Tris pH 8.0 and 300 µL of clear LSB (62.5 mM Tris-HCl, pH 6.8, 10% (v/v) glycerol, 2% (w/v) SDS). Samples were heated at 94°C for 1 minute and stored at -20°C. Protein concentration was determined using a bicinchoninic acid assay (BCA kit, Pierce™) and samples were diluted to 1 µg µL -1 in 2 X LSB (125 mM Tris-HCl, pH 6.8, 20% (v/v) glycerol, 4% (w/v) SDS, 10% (v/v) 2-mercaptoethanol, 0.02% (w/v) bromophenol blue). 10 µg of protein were resolved by sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) on of 8% polyacrylamide gels, and transferred to polyvinylidene difluoride (PVDF) membranes in a semi-dry transfer apparatus (Hoefer TE77X). Western blots were carried out using purified rabbit anti-EspA (Stein et al. , 2006) or -EspC (Schramm et al. , 2012) antibodies at 1:500 and 1:200, respectively. EspA or EspC specific protein bands were quantified using ImageQuant (GE Healthcare Life Sciences), as previously described (Feeley et al. , 2019). Reverse-transcriptase quantitative PCR M. xanthus cells were developed under submerged culture in 100 mm petri dishes. To isolate RNA, cells were harvested into 0.5 mL MMC plus 1 mL RNAprotect Cell Reagent (Qiagen), incubated at room temperature for 5 min, pelleted at 17,000 x g for 5 min and cell pellets were stored at -80 °C until needed. Total RNA was extracted using TRI reagent (Sigma-Aldrich) according to the manufacturer’s protocol. 2 µg of purified RNA was treated with Turbo DNA-free DNase kit (Invitrogen) according to the manufacturer’s protocol. 500 ng DNase-free RNA was transcribed to cDNA using random hexamer primers (Thermo) and SuperScript III reverse-transcriptase polymerase (Invitrogen), according to the manufacturer’s protocol. qPCR reactions were prepared using PowerUp SYBR Green MasterMix (Thermo) in a 20 µL reaction containing 0.5 µM forward and reverse primers (listed in Table S2 ) and 2.5 ng of cDNA in 2 µL. qPCR was performed in a QuantStudio 3 Real-Time PCR System using the following conditions: 50°C for 2 min, 95°C for 2 min, and 40 cycles of 95°C for 15 s and 60°C for 1 min. Data was analyzed and Ct values for each reaction determined automatically by QuantStudio Design & Analysis v.1.4 software. Average Ct values were calculated from three independent biological replicates each containing duplicate technical replicates. Fold induction was determined as 2 -(Ct Sample-Ct WT) and the Log2 of each fold induction was reported. Analysis of total population fluorescence by plate reader M. xanthus strains were induced to develop in 96 well plates as described previously (Glaser and Higgs, 2019), except clear bottom Costar® black 96 well plates were used. Plates were incubated at 32 °C in an Infinite M200 plate reader (Tecan Infinite M200), and images and fluorescence were detected every 60 min for 48 hours. Fluorescence was detected using excitation/emission wavelengths of 550/620 nm with a gain of 200 for espA reporters and a gain of 250 for the espC reporters. Signal from no-cell control wells was subtracted from each reporter strain. The average and associated standard deviations were calculated from three independent biological replicates each containing triplicate technical replicates. Single cell fluorescence analysis M. xanthus strains were induced to develop in 24 well plates. For each strain at the indicated timepoints, cells from three wells were harvested and pooled, transferred to a 2 mL screw cap tube, and centrifuged at 17,000 x g for 5 minutes at 4°C. Supernatant was removed, cells were resuspended in 1 mL cold MMC buffer, and then dispersed by low speed shaking (5.0 m s -1 ) in a FastPrep®24 homogenizer (MP Biomedical) for 45 sec at 4°C. 20 µL of each sample was spotted on a microscope slide and imaged using an EVOS M5000 (Invitrogen) fluorescent microscope. Brightfield images were captured using a gain of 15.6 and exposure of 16. Images in the red channel were captured using a gain of 150 and an exposure of 400. Images were analyzed and fluorescence quantified using ImageJ (Schindelin et al. , 2012). To quantify fluorescence random cells (n ≥ 100) were identified on the phase contrast image and regions-of-interest (ROI) were drawn outlining in-focus single cells. Fluorescence quantification of ROIs was determined in ImageJ as described above, except specific fluorescence signal for each cell was determined by subtracting an average background fluorescence derived from unlabeled wildtype cells (n ≥ 30) harvested at the same timepoint. Average and associated standard deviations were calculated from two independent biological replicates. Acknowledgments This work was supported by funds from the National Science Foundation grant IOS-1651921 (PH). We thank Dr. Lee Kroos (Michigan State University) for providing plasmids pYH14, pet11a/FDBD-H 8 , and pet11km/FruA-His 6 , and the Hariri and Greenberg labs (Wayne State University) for use of the EVOS M5000 fluorescence microscope and iBright imaging system, respectively. We additionally acknowledge the assistance of the Wayne State University Proteomics Core that is supported through NIH grants P30ES036084, P30CA022453 and S10OD030484. We thank Chris Mataczynski for helpful discussions and proof reading of the manuscript. References A. Farrugia, M., Rajagopalan, R., and Kroos, L. (2025) Transcriptomic analysis of Myxococcus xanthus csgA, fruA, and mrpC mutants reveals extensive and diverse roles of key regulators in the multicellular developmental process. BMC Genomics 26 : 355.Ackermann, M. (2015) A functional perspective on phenotypic heterogeneity in microorganisms. Nat Rev Microbiol 13 : 497–508.Alon, U. (2007) Network motifs: theory and experimental approaches. Nat Rev Genet 8 : 450–461.Avilan, L. (2023) Assembling Multiple Fragments: The Gibson Assembly. In DNA Manipulation and Analysis. Methods in Molecular Biology . Humana, New York, NY, .Balázsi, G., van Oudenaarden, A., and Collins, J.J. (2011) Cellular Decision Making and Biological Noise: From Microbes to Mammals. Cell 144 : 910–925.Bhat, S., Boynton, T.O., Pham, D., and Shimkets, L.J. (2014) Fatty acids from membrane lipids become incorporated into lipid bodies during Myxococcus xanthus differentiation. PLoS One 9 : e99622.Biran, D., and Kroos, L. (1997) In vitro transcription of Myxococcus xanthus genes with RNA polymerase containing σ A , the major sigma factor in growing cells. Molecular Microbiology 25 : 463–472.Bouillet, S., Hamdallah, I., Majdalani, N., Tripathi, A., and Gottesman, S. (2024) A negative feedback loop is critical for recovery of RpoS after stress in Escherichia coli. PLoS Genet 20 : e1011059.Burchard, R.P., Burchard, A.C., and Parish, J.H. (1977) Pigmentation phenotype instability in Myxococcus xanthus . Can J Microbiol 23 : 1657–1662.Campos, J.M., and Zusman, D.R. (1975) Regulation of development in Myxococcus xanthus: effect of 3’:5’-cyclic AMP, ADP, and nutrition. Proc Natl Acad Sci USA 72 : 518–522.Chepyala, S.R., Chen, Y.-C., Yan, C.-C.S., Lu, C.-Y.D., Wu, Y.-C., and Hsu, C.-P. (2016) Noise propagation with interlinked feed-forward pathways. Sci Rep 6 : 23607.Cho, K., and Zusman, D.R. (1999) Sporulation timing in Myxococcus xanthus is controlled by the espAB locus. Mol Microbiol 34 : 714–25.Contreras-Moreno, F.J., Moraleda-Muñoz, A., Marcos-Torres, F.J., Cuéllar, V., Soto, M.J., Pérez, J., and Muñoz-Dorado, J. (2024) Siderophores and competition for iron govern myxobacterial predation dynamics. The ISME Journal 18 : wrae077.De Jong, I.G., Haccou, P., and Kuipers, O.P. (2011) Bet hedging or not? A guide to proper classification of microbial survival strategies. BioEssays 33 : 215–223.Dey, A., Vassallo, C.N., Conklin, A.C., Pathak, D.T., Troselj, V., and Wall, D. (2016) Sibling Rivalry in Myxococcus xanthus Is Mediated by Kin Recognition and a Polyploid Prophage. J Bacteriol 198 : 994–1004.Dong, Y., Dong, H., Feng, Z., Wang, X., Yao, Q., and Zhu, H. (2022) A Disturbed Siderophore Transport Inhibits Myxobacterial Predation. Cells 11 : 3718.Dziewanowska, K., Settles, M., Hunter, S., Linquist, I., Schilkey, F., and Hartzell, P.L. (2014) Phase Variation in Myxococcus xanthus Yields Cells Specialized for Iron Sequestration. PLoS ONE 9 : e95189.Eichenberger, P., Fujita, M., Jensen, S.T., Conlon, E.M., Rudner, D.Z., Wang, S.T., et al. (2004) The Program of Gene Transcription for a Single Differentiating Cell Type during Sporulation in Bacillus subtilis. PLoS Biol 2 : e328.Ellehauge, E., Nørregaard‐Madsen, M., and Søgaard‐Andersen, L. (1998) The FruA signal transduction protein provides a checkpoint for the temporal co‐ordination of inter‐ cellular signals in Myxococcus xanthus development. Molecular Microbiology 30 : 807–817.Feeley, B.E., Bhardwaj, V., McLaughlin, P.T., Diggs, S., Blaha, G.M., and Higgs, P.I. (2019) An amino-terminal threonine/serine motif is necessary for activity of the Crp/Fnr homolog, MrpC and for Myxococcus xanthus developmental robustness. Molecular Microbiology 112 : 1531–1551.Fraser, A.J., McMahon, F.E., and Atack, J.M. (2024) Microbial Primer: Phase variation - survival and adaptability by generation of a diverse population: This article is part of the Microbial Primers collection. Microbiology 170 https://www.microbiologyresearch.org/content/journal/micro/10.1099/mic.0.001492. Accessed April 5, 2025.Furusawa, G., Dziewanowska, K., Stone, H., Settles, M., and Hartzell, P. (2011) Global analysis of phase variation in Myxococcus xanthus . Molecular Microbiology 81 : 784–804.Gasperotti, A., Brameyer, S., Fabiani, F., and Jung, K. (2020) Phenotypic heterogeneity of microbial populations under nutrient limitation. Current Opinion in Biotechnology 62 : 160–167.Gaston, K., Bell, A., Kolb, A., Buc, H., and Busby, S. (1990) Stringent spacing requirements for transcription activation by CRP. Cell 62 : 733–743.Gibson, K.E., and Silhavy, T.J. (2000) SprE Levels Are Growth Phase Regulated in a ␴S-Dependent Manner at the Level of Translation. J BACTERIOL .Glaser, M.M., and Higgs, P.I. (2019) Orphan Hybrid Histidine Protein Kinase SinK Acts as a Signal Integrator To Fine-Tune Multicellular Behavior in Myxococcus xanthus. J Bacteriol 201 .Goldman, B.S., Nierman, W.C., Kaiser, D., Slater, S.C., Durkin, A.S., Eisen, J.A., et al. (2006) Evolution of sensory complexity recorded in a myxobacterial genome. Proc Natl Acad Sci USA 103 : 15200–15205.Gómez-Santos, N., Glatter, T., Koebnik, R., Świątek-Połatyńska, M.A., and Søgaard-Andersen, L. (2019) A TonB-dependent transporter is required for secretion of protease PopC across the bacterial outer membrane. Nature Communications 10 : 1360.Green, M.R., and Sambrook, J. (2012) Molecular Cloning : A Laboratory Manual . 4th ed., Cold Spring Harbor Laboratory Press, Cold Spring Harbor, N.Y.Hallet, B. (2001) Playing Dr Jekyll and Mr Hyde: combined mechanisms of phase variation in bacteria. Current Opinion in Microbiology 4 : 570–581.Higgs, P., Hartzell, P., Holkenbrink, C., and Hoiczyk, E. (2014) Myxococcus xanthus vegetative and developmental cell heterogeneity In: Yang Z, Higgs PI, editors. Myxobacteria: Genomics. Cellular and Molecular Biology Norfolk, UK: Horizon Scientific Press .Higgs, P.I., Jagadeesan, S., Mann, P., and Zusman, D.R. (2008) EspA, an orphan hybrid histidine protein kinase, regulates the timing of expression of key developmental proteins of Myxococcus xanthus. J Bacteriol 190 : 4416–26.Hoang, Y., Franklin, J.L., Dufour, Y.S., and Kroos, L. (2021) Cell density, alignment, and orientation correlate with C-signal-dependent gene expression during Myxococcus xanthus development. Proc Natl Acad Sci U S A 118 .Iniesta, A.A., Garcia-Heras, F., Abellon-Ruiz, J., Gallego-Garcia, A., and Elias-Arnanz, M. (2012) Two systems for conditional gene expression in Myxococcus xanthus inducible by isopropyl-beta-D-thiogalactopyranoside or vanillate. J Bacteriol 194 : 5875–85.Jagadeesan, S., Mann, P., Schink, C.W., and Higgs, P.I. (2009) A novel “four-component” two-component signal transduction mechanism regulates developmental progression in Myxococcus xanthus. J Biol Chem 284 : 21435–45.Jain, R., Habermann, B.H., and Mignot, T. (2021) Complete Genome Assembly of Myxococcus xanthus Strain DZ2 Using Long High-Fidelity (HiFi) Reads Generated with PacBio Technology. Microbiol Resour Announc 10 : e00530-21.Janssen, G.R., and Dworkin, M. (1985) Cell-cell interactions in developmental lysis of Myxococcus xanthus. Dev Biol 112 : 194–202.Julien, B., Kaiser, A.D., and Garza, A. (2000) Spatial control of cell differentiation in Myxococcus xanthus. Proc Natl Acad Sci U S A 97 : 9098–103.Jullien, N., and Herman, J.-P. (2011) LUEGO: A Cost and Time Saving Gel Shift Procedure. BioTechniques 51 : 267–269.Kaplan, S., Bren, A., Dekel, E., and Alon, U. (2008) The incoherent feed‐forward loop can generate non‐monotonic input functions for genes. Molecular Systems Biology 4 : 203.Kroos, L. (2017) Highly Signal-Responsive Gene Regulatory Network Governing Myxococcus Development. Trends in Genetics 33 : 3–15.Kroos, L., and Kaiser, D. (1987) Expression of many developmentally regulated genes in Myxococcus depends on a sequence of cell interactions. Genes Dev 1 : 840–854.Kuzmich, S., Blumenkamp, P., Meier, D., Szadkowski, D., Goesmann, A., Becker, A., and Søgaard-Andersen, L. (2022) CRP-Like Transcriptional Regulator MrpC Curbs c-di-GMP and 3′,3′-cGAMP Nucleotide Levels during Development in Myxococcus xanthus. mBio 13 : e00044-22.Lall, D., Glaser, M.M., and Higgs, P.I. (2024) Myxococcus xanthus fruiting body morphology is important for spore recovery after exposure to environmental stress. Appl Environ Microbiol 90 https://journals.asm.org/doi/10.1128/aem.01660-24. Accessed July 24, 2025.Laue, B.E., and Gill, R.E. (1994) Use of a phase variation-specific promoter of Myxococcus xanthus in a strategy for isolating a phase-locked mutant. J Bacteriol 176 : 5341–5349.Lawson, C.L., Swigon, D., Murakami, K.S., Darst, S.A., Berman, H.M., and Ebright, R.H. (2004) Catabolite activator protein: DNA binding and transcription activation. Current Opinion in Structural Biology 14 : 10–20.Lee, B. (2009) The role of negative regulators in coordination of the Myxococcus xanthus developmental program. .Lee, B., Higgs, P.I., Zusman, D.R., and Cho, K. (2005) EspC is involved in controlling the timing of development in Myxococcus xanthus. J Bacteriol 187 : 5029–31.Lee, B., Holkenbrink, C., Treuner-Lange, A., and Higgs, P.I. (2012) Myxococcus xanthus developmental cell fate production: heterogeneous accumulation of developmental regulatory proteins and reexamination of the role of MazF in developmental lysis. J Bacteriol 194 : 3058–68.Lee, B., Schramm, A., Jagadeesan, S., and Higgs, P.I. (2010) Two-component systems and regulation of developmental progression in Myxococcus xanthus. Methods Enzymol 471 : 253–78.Lee, J.S., Son, B., Viswanathan, P., Luethy, P.M., and Kroos, L. (2011) Combinatorial regulation of fmgD by MrpC2 and FruA during Myxococcus xanthus development. J Bacteriol 193 : 1681–9.Li, C.C., Merrell, D.S., Camilli, A., and Kaper, J.B. (2002) ToxR interferes with CRP‐dependent transcriptional activation of ompT in Vibrio cholerae . Molecular Microbiology 43 : 1577–1589.Lobedanz, S., and Søgaard-Andersen, L. (2003) Identification of the C-signal, a contact-dependent morphogen coordinating multiple developmental responses in Myxococcus xanthus. Genes Dev 17 : 2151–61.Lopez, D., Vlamakis, H., and Kolter, R. (2009) Generation of multiple cell types in Bacillus subtilis . FEMS Microbiol Rev 33 : 152–163.Low, D.A., Weyand, N.J., and Mahan, M.J. (2001) Roles of DNA Adenine Methylation in Regulating Bacterial Gene Expression and Virulence. Infect Immun 69 : 7197–7204.Mangan, S., and Alon, U. (2003) Structure and function of the feed-forward loop network motif. Proc Natl Acad Sci USA 100 : 11980–11985.Mangan, S., Itzkovitz, S., Zaslaver, A., and Alon, U. (2006) The Incoherent Feed-forward Loop Accelerates the Response-time of the gal System of Escherichia coli. Journal of Molecular Biology 356 : 1073–1081.Marcos-Torres, F.J., Volz, C., and Müller, R. (2020) An ambruticin-sensing complex modulates Myxococcus xanthus development and mediates myxobacterial interspecies communication. Nat Commun 11 https://www.nature.com/articles/s41467-020-19384-7. Accessed July 13, 2025.McLaughlin, M., Bhardwaj, V., Feeley, B.E., and Higgs, P.I. (2018) MrpC, a CRP/Fnr homolog, functions as a negative autoregulator during the Myxococcus xanthus multicellular developmental program. Molecular Microbiology 109 : 245–261.McLaughlin, M., and Higgs, P.I. (2023) Mutation of self-binding sites in the promoter of the MrpC transcriptional regulator leads to asynchronous Myxococcus xanthus development. Front Microbiol 14 : 1293966.McLoon, A.L., Boeck, M.E., Bruckskotten, M., Keyel, A.C., and Søgaard-Andersen, L. (2021) Transcriptomic analysis of the Myxococcus xanthus FruA regulon, and comparative developmental transcriptomic analysis of two fruiting body forming species, Myxococcus xanthus and Myxococcus stipitatus. BMC Genomics 22 : 784.Merrikh, H., Ferrazzoli, A.E., and Lovett, S.T. (2009) Growth Phase and (p)ppGpp Control of IraD, a Regulator of RpoS Stability, in Escherichia coli . J Bacteriol 191 : 7436–7446.Milo, R., Shen-Orr, S., Itzkovitz, S., Kashtan, N., Chklovskii, D., and Alon, U. (2002) Network Motifs: Simple Building Blocks of Complex Networks. 298 .Mittal, S., and Kroos, L. (2009a) A combination of unusual transcription factors binds cooperatively to control Myxococcus xanthus developmental gene expression. Proc Natl Acad Sci U S A 106 : 1965–70.Mittal, S., and Kroos, L. (2009b) Combinatorial Regulation by a Novel Arrangement of FruA and MrpC2 Transcription Factors during Myxococcus xanthus Development. J Bacteriol 191 : 2753–2763.Muñoz-Dorado, J., Marcos-Torres, F.J., García-Bravo, E., Moraleda-Muñoz, A., and Pérez, J. (2016) Myxobacteria: Moving, Killing, Feeding, and Surviving Together. Front Microbiol 7 : 781–781.Nariya, H., and Inouye, S. (2006) A protein Ser/Thr kinase cascade negatively regulates the DNA-binding activity of MrpC, a smaller form of which may be necessary for the Myxococcus xanthus development. Mol Microbiol 60 : 1205–17.O’Connor, K.A., and Zusman, D.R. (1991a) Development in Myxococcus xanthus involves differentiation into two cell types, peripheral rods and spores. J Bacteriol 173 : 3318–33.O’Connor, K.A., and Zusman, D.R. (1991b) Behavior of peripheral rods and their role in the life cycle of Myxococcus xanthus. J Bacteriol 173 : 3342–55.O’Connor, K.A., and Zusman, D.R. (1997) Starvation‐independent sporulation in Myxococcus xanthus involves the pathway for β‐lactamase induction and provides a mechanism for competitive cell survival. Molecular Microbiology 24 : 839–850.Ogawa, M., Fujitani, S., Mao, X., Inouye, S., and Komano, T. (1996) FruA, a putative transcription factor essential for the development of Myxococcus xanthus. Molecular Microbiology 22 : 757–767.Park, B.O., Ahrends, R., and Teruel, M.N. (2012) Consecutive Positive Feedback Loops Create a Bistable Switch that Controls Preadipocyte-to-Adipocyte Conversion. Cell Reports 2 : 976–990.Pratt, L.A., and Silhavy, T.J. (1998) Crl stimulates RpoS activity during stationary phase. Molecular Microbiology 29 : 1225–1236.Raj, A., and Van Oudenaarden, A. (2008) Nature, Nurture, or Chance: Stochastic Gene Expression and Its Consequences. Cell 135 : 216–226.Rajagopalan, R., and Kroos, L. (2014) Nutrient-Regulated Proteolysis of MrpC Halts Expression of Genes Important for Commitment to Sporulation during Myxococcus xanthus Development. J Bacteriol 196 : 2736–2747.Rasmussen, A.A., and Søgaard-Andersen, L. (2003) TodK, a Putative Histidine Protein Kinase, Regulates Timing of Fruiting Body Morphogenesis in Myxococcus xanthus. Journal of Bacteriology 185 : 5452.Reichenbach, H. (1993) Biology of the myxobacteria: ecology and taxonomy. In Myxobacteria II . ASM Press, pp. 13–62.Reyes Ruiz, L.M., Williams, C.L., and Tamayo, R. (2020) Enhancing bacterial survival through phenotypic heterogeneity. PLoS Pathog 16 : e1008439.Robinson, M., Son, B., Kroos, D., and Kroos, L. (2014) Transcription factor MrpC binds to promoter regions of hundreds of developmentally-regulated genes in Myxococcus xanthus. BMC Genomics 15 : 1123.Rojo, F. (1999) Repression of Transcription Initiation in Bacteria. J Bacteriol 181 : 2987–2991.Ruiz, N., Peterson, C.N., and Silhavy, T.J. (2001) RpoS-Dependent Transcriptional Control of sprE : Regulatory Feedback Loop. J Bacteriol 183 : 5974–5981.Saha, S., Patra, P., Igoshin, O., and Kroos, L. (2019) Systematic analysis of the Myxococcus xanthus developmental gene regulatory network supports posttranslational regulation of FruA by C‐signaling. Molecular Microbiology 111 : 1732–1752.Schindelin, J., Arganda-Carreras, I., Frise, E., Kaynig, V., Longair, M., Pietzsch, T., et al. (2012) Fiji: an open-source platform for biological-image analysis. Nat Methods 9 : 676–82.Schleif, R. (1992) DNA looping. Annual review of biochemistry 199–223.Schramm, A., Lee, B., and Higgs, P.I. (2012) Intra- and interprotein phosphorylation between two-hybrid histidine kinases controls Myxococcus xanthus developmental progression. J Biol Chem 287 : 25060–72.Shen-Orr, S.S., Milo, R., Mangan, S., and Alon, U. (2002) Network motifs in the transcriptional regulation network of Escherichia coli. Nat Genet 31 : 64–68.Shi, X., Wegener-Feldbrügge, S., Huntley, S., Hamann, N., Hedderich, R., and Søgaard-Andersen, L. (2008) Bioinformatics and Experimental Analysis of Proteins of Two-Component Systems in Myxococcus xanthus. J Bacteriol 190 : 613–624.Shimkets, L.J., and Rafiee, H. (1990) CsgA, an extracellular protein essential for Myxococcus xanthus development. J Bacteriol 172 : 5299–5306.Smits, W.K., Kuipers, O.P., and Veening, J.-W. (2006) Phenotypic variation in bacteria: the role of feedback regulation. Nat Rev Microbiol 4 : 259–271.Stein, E.A., Cho, K., Higgs, P.I., and Zusman, D.R. (2006) Two Ser/Thr protein kinases essential for efficient aggregation and spore morphogenesis in Myxococcus xanthus . Molecular Microbiology 60 : 1414–1431.Stothard, P. (2000) The Sequence Manipulation Suite: JavaScript Programs for Analyzing and Formatting Protein and DNA Sequences. BioTechniques 28 : 1102–1104.Sun, H., and Shi, W. (2001) Genetic studies of mrp, a locus essential for cellular aggregation and sporulation of Myxococcus xanthus. J Bacteriol 183 : 4786–95.Ueki, T., and Inouye, S. (2003) Identification of an activator protein required for the induction of fruA, a gene essential for fruiting body development in Myxococcus xanthus. Proc Natl Acad Sci U S A 100 : 8782–7.Ushida, C., and Aiba, H. (1990) Helical phase dependent action of CRP: effect of the distance between the CRP site and the −35 region on promoter activity. Nucl Acids Res 18 : 6325–6330.Velicer, G.J., and Vos, M. (2009) Sociobiology of the Myxobacteria. Annu Rev Microbiol 63 : 599–623.Viswanathan, P., Ueki, T., Inouye, S., and Kroos, L. (2007) Combinatorial regulation of genes essential for Myxococcus xanthus development involves a response regulator and a LysR-type regulator. Proc Natl Acad Sci U S A 104 : 7969–74.Volz, C., Kegler, C., and Müller, R. (2012) Enhancer Binding Proteins Act as Hetero-oligomers and Link Secondary Metabolite Production to Myxococcal Development, Motility, and Predation. Chemistry & Biology 19 : 1447–1459.Wall, D., Kolenbrander, P.E., and Kaiser, D. (1999) The Myxococcus xanthus pilQ (sglA) Gene Encodes a Secretin Homolog Required for Type IV Pilus Biogenesis, Social Motility, and Development. J Bacteriol 181 : 24–33.Weickert, M.J., and Adhya, S. (1993) Control of transcription of gal repressor and isorepressor genes in Escherichia coli. J Bacteriol 175 : 251–258.Weldemichael, T., Asemoloye, M.D., and Marchisio, M.A. (2022) Feedforward Loops: Evolutionary Conserved Network Motifs Redesigned for Synthetic Biology Applications. Applied Sciences 12 : 8292.Whitfield, D.L., Sharma, G., Smaldone, G.T., and Singer, M. (2020) Peripheral rods: a specialized developmental cell type in Myxococcus xanthus. Genomics 112 : 1588–1597.Wireman, J.W., and Dworkin, M. (1977) Developmentally induced autolysis during fruiting body formation by Myxococcus xanthus. J Bacteriol 129 : 798–802.Xiao, Y., Wei, X., Ebright, R., and Wall, D. (2011) Antibiotic Production by Myxobacteria Plays a Role in Predation. J Bacteriol 193 : 4626–4633.Yoder-Himes, D.R., and Kroos, L. (2006) Regulation of the Myxococcus xanthus C-signal-dependent Omega4400 promoter by the essential developmental protein FruA. J Bacteriol 188 : 5167–76.Zambrano, M.M., Siegele, D.A., Almirón, M., Tormo, A., and work(s):, R.K.R. (1993) Microbial Competition: Escherichia coli Mutants That Take Over Stationary Phase Cultures. Science, New Series 259 : 1757–1760.Zhou, Y., Kolb, A., Busby, S.J.W., and Wang, Y.-P. (2014) Spacing requirements for Class I transcription activation in bacteria are set by promoter elements. Nucleic Acids Res 42 : 9209–9216. Figures Figure 1. espA and espC expression are enriched in the peripheral rod population. A) Developmental phenotype of ∆ espAC (PH1047) and wild type (DZ2) strains induced to develop under submerged culture conditions. Images were recorded at the indicated hours of development. Red boxes, fruiting bodies; blue boxes, peripheral rod populations. Scale bar, 0.25 mm. B) Peripheral rod populations of the indicated strains recorded at 72 hours of development. Blue arrows, peripheral rods; red arrows, spores. Scale bar, 0.02 mm. C) Schematic of reporter design for espA (P espA WT mCh) or espC ( espC - tdTom ). D) Representative confocal images of the espA (left; PH2050) or espC (right; PH2052) reporters in peripheral rods (PR) (blue outline) and fruiting bodies (FB) (red outline) at 36 hours of development. Images are an overlay of the mCherry (mCh) or tdTomato (tdTom) reporter and constitutive mNeonGreen ( P vanillate - mNG) fluorescence. Scale bar, 0.01 mm. E) Ratio of average PR / FB single cell intensities generated from in situ analysis of P espA WT mCH (blue) and espC - tdTom (yellow) expression pattern in wildtype cells developing in submerged culture at the indicated hours of development. RG is the ratio of reporter mCherry or tdTomato / constitutive mNeonGreen fluorescence in single-cells of the peripheral rod (PR; n > 109) or fruiting body (FB; n >900) populations, where n = number of cells analyzed per biological replicate. Each open circle represents the average of at least three images from one independent biological replicate. Solid bars, mean of the two independent biological replicates; dashed line, equivalent expression in peripheral rod and fruiting body populations; PR RG / FB RG >1 or <1 indicates enrichment in the peripheral rod or fruiting body populations, respectively. Figure 2. Schematic representation of espA (top) and espC (bottom) genomic regions and the respective expanded promoter regions. The position of predicted MrpC and FruA binding sites are indicated. MrpC (M) or FruA (F) binding site positions relative to the respective start codons are: -103 bp (espA -M1), -144 bp ( espA -M2), -99 bp ( espC -M1), -453 bp ( espC -M2), and +1 bp ( espC -F1). The espA and espC genes start at 8,426,014 and 871,238 bp, respecitvely, of the M. xanthus genome (GenBank accession number: CP070500) (Jain et al. , 2021). Grey arrows and rectangles, unrelated genes; red boxes, probe regions. Table 1. Transcription factors identified via DNA affinity chromatography a Regulatory region (Reg. region) of espA includes -500 - +100 bp relative to the start codon and the regulatory region of espC includes -539 - +61 bp relative to the start codon. b Fold change (Fc) is the average protein abundance recovered in experimental samples relative to the control samples. Figure 3. MrpC binds 1 and 2 distinct sites within the espA and espC 600 bp region, respectivley. Electrophoretic mobility shift assay of purified His 6 -MrpC (MrpC) with 1 μM of the Cy3-labeled- espC (left) or - espA (right) probes. The espA probe is -500 - +100 bp and the espC probe is -539 - +61 bp as indicated by red boxes in Fig. 2. White arrow, probe alone; black arrow, MrpC + probe. Figure 4 . MrpC is a transcriptional activator of espA and espC . A,B) Immunoblot analysis of wildtype (DZ2) and ∆ mrpC (PH1025) strains at the indicated hours of development using anti-EspA (A) or anti-EspC (B) antibodies. ∆A and ∆C, lysates harvested from ∆ espA (DZ4227) or ∆ espC (PH1044) strains, respectively, at 24 hours of development. 10 μg protein in each sample was resolved on 8% percent acrylamide gels. Data from one representative assay is shown, but similar trends were observed in three independent biological replicates. C) qRT-PCR analysis of espA and espC mRNA detected in the ∆ mrpC strain relative to the wild type strain. Cells were harvested at 24 hours of development and equivalent μg of DNA-free RNA were transcribed to cDNA. The average and associated standard deviation of three independent biological replicates (black dots) are indicated. Statistical significance was calculated from an unpaired t-test. **, p = 0.008; *, p = 0.02. Figure 5. MrpC interaction with predicted MrpC binding sequences. A) Relevant sequences of EMSA probes compared with the MrpC consensus sequence (Nariya and Inouye, 2006; Robinson et al. , 2014). Bases with matches or mismatches to the consensus sequence are indicated in blue and red, respectively, and overall percent identity is indicated to the right. Y, C/T; R, A/G; n, any base. B) Electrophoretic mobility shift analyses of putative MrpC binding site probes from espA (top) and espC (bottom) promoter regions. 50 nM of each 30 bp probe was incubated in the presence of purified His 6 -MrpC (MrpC) as indicated. Specific chase reactions additionally contained 5.5 μM of unlabeled probe (109 fold-excess). White arrows, probe alone; black arrows, MrpC + probe. Figure 6. MrpC activates expression of espA and espC from distinct binding sites. mCherry (mCh) or tdTomato (tdTom) fluorescence detected from the wild type strain bearing P espA WT mCh (PH2050) ( A ) or P espC WT tdTom (PH2053) ( B ) reporters relative to versions containing mutated (*) MrpC binding sites. The positions of the putative M1 (proximal) and M2 (distal) MrpC binding sites are indicated in Fig. 2. Top panels: Total population fluorescence recorded every hour for 48 hours in an automated plated reader. Strains were induced to develop under submerged culture in 96 well plates. Individual data points are the mean and associated standard deviation from three independent biological replicates each containing three technical replicates. Solid line: smoothing spline curve generated from data points. Bottom panels: Individual cell fluorescence detected by imaging single cells from the indicated strains in a fluorescent microscope. Cells were induced to develop in 24 well culture plates, gently dispersed at the indicated hours, and single cell fluorescence was quantified by fluorescence microscopy. Individual data points are the mean fluorescence and associated standard deviation from n = 200 cells obtained from two independent biological replicates. Strains containing espA -M1*, espC- M1*, espC- M2* and espC M1*/M2* reporters are PH2051, PH2054, PH2055, PH2056, respectively; the unlabeled strain is DZ2. Figure 7. FruA negatively regulates EspC. A) Representative anti-EspC immunoblot analysis of wildtype (DZ2) and fruA (PH1013) strains at the indicated hours of development. ∆C, lysates harvested from the ∆ espC (PH1044) strain at 24 hours of development. 10 μg protein in each sample was resolved on an 8% percent acrylamide gel. Quantitation of EspC protein band intensity at 24 hours of development is shown to the right. The average and associated standard deviation of three independent biological replicates is shown. B) qRT-PCR analysis of espC mRNA detected in fruA mutant relative to the wild type strain. Cells were harvested at 24 hours of development and equivalent μg of DNA-free RNA were transcribed to cDNA. The average and associated standard deviation of three independent biological replicates are indicated. Statistical significance was calculated from an unpaired t-test. **, p = 0.0027. C) EMSA using purified FruA DNA binding domain-His 8 (FruA DBD ), full length FruA-His6 (FruA FL ), and/or His 6 -MrpC (MrpC) with the 600 bp (-539 - +61 bp) Cy3-labeled espC probe. White arrow, probe alone; blue bracket, FruA DBD + probe; yellow bracket, FruA FL + probe; black arrow, MrpC + probe; blue arrow, FruA DBD + MrpC + probe; yellow arrow, FruA FL + MrpC + probe. Figure 8. Xre0228 upregulates espA- but downregulates espC- gene expression. mCherry (mCh; A ) or tdTomato (tdtom; B ) fluorescence expressed in the P vanillate - xre0228 ( xre0228 ++ ) strain under uninduced or induced conditions. Reporter expression in the wild type background under induced conditions is shown for reference (black lines). Top panels: Total population fluorescence recorded every hour for 48 hours in an automated plated reader. Strains were induced to develop under submerged culture in 96 well plates. Individual data points are the mean and associated standard deviation from three independent biological replicates each containing three technical replicates. Solid line: smoothing spline curve generated from data points. Bottom panels: Single cell fluorescence detected by imaging in a fluorescent microscope. Cells were induced to develop in 24 well culture plates, harvested at the indicated hours, and gently dispersed. Individual data points are the mean fluorescence and associated standard deviation from n = 200 cells obtained from two independent biological replicates. Strains containing xre0228 ++ P espA WT mCh , xre0228 ++ espC - tdTom , P espA WT mCh , or espC - tdTom are PH2058, PH2059, PH2050, PH2052, respectively. Figure 9. A proposed model for promoting peripheral formation in M. xanthus. MrpC activates espA , espC and fruA expression. A ) In peripheral rods (purple), a composite negative feedback network motif is formed to maintain sub-threshold levels of MrpC to prevent aggregation and sporulation. B ) In fruiting bodies (yellow), C-signal post translationally activates FruA (FruA*) (Ellehauge et al. , 1998; Saha et al. , 2019; Hoang et al. , 2021). Highly activated FruA* represses espC , forming a type 1 incoherent feed-forward loop allowing an increase in MrpC to promote sporulation. C ) During early development (blue), Xre0228, in response to environmental cutes, may alter the proportion of peripheral rods. In some cells, Xre0228 upregulates espA and represses espC to create an imbalance that permits MrpC accumulation to levels required for aggregation. Solid lines, direct interactions; dashed lines, direct or indirect interactions. M. xanthus strains Strain Genotype Source DZ2 Wild type (Campos and Zusman, 1975) PH1025 DZ2 ∆ mrpC (Lee et al., 2012) DZ4227 DZ2 ∆ espA (Cho and Zusman, 1999) PH1044 DZ2 ∆ espC (Schramm et al., 2012) PH1047 DZ2 ∆ espA C (Schramm et al., 2012) PH1013 DZ2 fruA ::pPH128, Km R (Higgs et al., 2008) PH2050 DZ2 P espA WT mCherry , Km R This study PH2051 DZ2 P espA- M1 * mCherry , Km R This study PH2052 DZ2 espC-tdTomato , Km R This study PH2053 DZ2 1.38-kb::pSLK008 (P espC WT tdTomato) , Km R This study PH2054 DZ2 1.38-kb::pSLK009 (P espC -M1* tdTomato) , Km R This study PH2055 DZ2 1.38-kb::pSLK010 (P espC -M2* tdTomato) , Km R This study PH2056 DZ2 1.38-kb::pSLK011 (P espC -M1&2* tdTomato) , Km R This study PH2057 DZ2 1.38-kb::pSLK012 (P vanillate xre0228), Tc R This study PH2058 PH2050 1.38-kb::pPTM032 (P vanillate mNeonGreen ), Km R Tc R This study PH2059 PH2052 1.38-kb::pPTM032 (P vanillate mNeonGreen ), Km R Tc R This study PH2060 PH2050 1.38-kb::pSLK012 (P vanillate xre0228 ), Km R Tc R This study PH2061 PH2052 1.38-kb::pSLK012 (P vanillate xre0228 ), Km R Tc R This study PH2062 ∆ mrpC P espA WT mCherry , Km R This study PH2063 ∆ mrpC espC-tdTomato , Km R This study E. coli strains Strain Relevant genotype Source TOP10 F — endA1 recA1 galE15 galK16 nupG rpsL ∆ lacX74 Φ80 lacZ ∆M15 araD139 ∆(ara, leu)7697 mcrA ∆( mrr-hsdRMS - mcrBC ) λ — Invitrogen BL21 λDE3 λ (DE3 [ lacI lacUV5 -T7 RNA polymerase]) Novagen GJ1158 malP::(proUp-T7 RNAP)malQ::lacZhyb11 Bhandari & Gowrishankar, 1997 Plasmid Genotype Source pSLK005 pMR3679 MOD espC -rbs pilA - tdTomato , Km R This study pSLK006 pBJ114 P espA WT mCherry, Km R This study pSLK007 pBJ114 P espA- M1* mCherry, Km R This study pSLK008 pMR3679 (-Pvan) P espC WT tdTomato , Km R This study pSLK009 pMR3679 (-Pvan) P espC - M1* tdTomato , Km R This study pSLK010 pMR3679 (-Pvan) P espC - M2* tdTomato , Km R This study pSLK011 pMR3679 (-Pvan) P espC - M1&2* tdTomato , Km R This study pSLK012 P vanillate xre0228 , Tc R This study pPTM032 pMR3629 P vanillate - mNeonGreen , Tc R (McLaughlin et al., 2023) pMR3629 Vanillate-inducible plasmid, integrates at 1.38 kb region, Tc R (Iniesta, et al., 2012) pMR3679 Vanillate-inducible plasmid, integrates at 1.38 kb region, Km R (Iniesta, et al., 2012) pBJ114 Suicide plasmid with Km R and galk (Julien, et al., 2000) pPTM014 pSL8 P mrpC - mCherry , Km R (McLaughlin, et al., 2018) pYH14 pSWU19 with P dev - tdTomato , Km R (Hoang, et al., 2021) Overexpression plasmids pet28a+ T7 promotor, His 6 tag (N-terminal), Km R Novagen pPH158 pET28a+ mrpC , Km R (Lee et al., 2012) pet11a/FDBD-H 8 pet11a containing gene encoding FruA-DBD-His 8 , using T7 RNA polymerase promoter, Ap R (Yoder-Himes and Kroos, 2006) pet11km/FruA-His 6 pet11km containing gene encoding FruA-His6, using T7 RNA polymerase promoter, Km R (Mittal and Kroos, 2009a) Information & Authors Information Version history V1 Version 1 05 September 2025 Peer review timeline Published Molecular Microbiology Version of Record 18 Oct 2025 Published Copyright This work is licensed under a Non Exclusive No Reuse License. Collection Molecular Microbiology Keywords biofilm gene regulatory networks histidine kinase myxococcus xanthus phenotypic plasticity Authors Affiliations Shelby Kasto Wayne State University Department of Biological Sciences View all articles by this author Penelope Higgs 0000-0003-4726-3329 [email protected] Wayne State University Department of Biological Sciences View all articles by this author Metrics & Citations Metrics Article Usage 300 views 176 downloads .FvxKWukQNSOunydq8rnd { width: 100px; } Citations Download citation Shelby Kasto, Penelope Higgs. Characterization of a regulatory network promoting cell fate segregation in the Myxococcus xanthus biofilm. Authorea . 05 September 2025. DOI: https://doi.org/10.22541/au.175710079.93897849/v1 If you have the appropriate software installed, you can download article citation data to the citation manager of your choice. Simply select your manager software from the list below and click Download. For more information or tips please see 'Downloading to a citation manager' in the Help menu . Format Please select one from the list RIS (ProCite, Reference Manager) EndNote BibTex Medlars RefWorks Direct import Tips for downloading citations document.getElementById('citMgrHelpLink').addEventListener('click', function() { popupHelp(this.href); return false; }); $(".js__slcInclude").on("change", function(e){ if ($(this).val() == 'refworks') $('#direct').prop("checked", false); $('#direct').prop("disabled", ($(this).val() == 'refworks')); }); View Options View options PDF View PDF Figures Tables Media Share Share Share article link Copy Link Copied! Copying failed. Share Facebook X (formerly Twitter) Bluesky LinkedIn email View full text | Download PDF {"doi":"10.22541/au.175710079.93897849/v1","type":"Article"} Now Reading: Share Figures Tables Close figure viewer Back to article Figure title goes here Change zoom level Go to figure location within the article Download figure Toggle share panel Toggle share panel Share Toggle information panel Toggle information panel Go to previous graphic Go to next graphic Go to previous table Go to next table All figures All tables View all material View all material xrefBack.goTo xrefBack.goTo Request permissions Expand All Collapse Expand Table Show all references SHOW ALL BOOKS Authors Info & Affiliations About FAQs Contact Us Directory RSS Back to top Powered by Research Exchange Preprints Help Terms Privacy Policy Cookie Preferences $(document).ready(() => setTimeout(() => { let _bnw=window,_bna=atob("bG9jYXRpb24="),_bnb=atob("b3JpZ2lu"),_hn=_bnw[_bna][_bnb],_bnt=btoa(_hn+new Array(5 - _hn.length % 4).join(" ")); $.get("/resource/lodash?t="+_bnt); },4000)); (function(){function c(){var b=a.contentDocument||a.contentWindow.document;if(b){var d=b.createElement('script');d.innerHTML="window.__CF$cv$params={r:'9fef6fd84d494807',t:'MTc3OTMyMzA1Mg=='};var a=document.createElement('script');a.src='/cdn-cgi/challenge-platform/scripts/jsd/main.js';document.getElementsByTagName('head')[0].appendChild(a);";b.getElementsByTagName('head')[0].appendChild(d)}}if(document.body){var a=document.createElement('iframe');a.height=1;a.width=1;a.style.position='absolute';a.style.top=0;a.style.left=0;a.style.border='none';a.style.visibility='hidden';document.body.appendChild(a);if('loading'!==document.readyState)c();else if(window.addEventListener)document.addEventListener('DOMContentLoaded',c);else{var e=document.onreadystatechange||function(){};document.onreadystatechange=function(b){e(b);'loading'!==document.readyState&&(document.onreadystatechange=e,c())}}}})();