Nutrient-regulated proteolysis of MrpC halts expression of genes important for commitment to sporulation during Myxococcus xanthus development.

Nutrient-Regulated Proteolysis of MrpC Halts Expression of Genes Important for Commitment to Sporulation during Myxococcus xanthus Development Ramya Rajagopalan, Lee Kroos

Starved Myxococcus xanthus cells glide to aggregation centers and form fruiting bodies in which rod-shaped cells differentiate into ovoid spores. Commitment to development was investigated by adding nutrients at specific times after starvation and determining whether development halted or proceeded. At 24 h poststarvation, some rod-shaped cells were committed to subsequent shape change and to becoming sonication-resistant spores, but nutrients caused partial disaggregation of fruiting bodies. By 30 h poststarvation, 10-fold more cells were committed to becoming sonication-resistant spores, and compact fruiting bodies persisted after nutrient addition. During the critical period of commitment around 24 to 30 h poststarvation, the transcription factors MrpC and FruA cooperatively regulate genes important for sporulation. FruA responds to short-range C-signaling, which increases as cells form fruiting bodies. MrpC was found to be highly sensitive to nutrient-regulated proteolysis both before and during the critical period of commitment to sporulation. The rapid turnover of MrpC upon nutrient addition to developing cells halted expression of the dev operon, which is important for sporulation. Regulated proteolysis of MrpC appeared to involve ATP-independent metalloprotease activity and may provide a mechanism for monitoring whether starvation persists and halting commitment to sporulation if nutrients reappear.

M

yxococcus xanthus is a bacterium that undergoes multicellular development and provides an excellent model to study mechanisms of signaling, gene regulation, and cell fate determination (1). M. xanthus cells move in packs that feed on available prey bacteria and organic matter (2). When starved, the population undergoes development, including aggregation of cells into mounds that become fruiting bodies as some of the rod-shaped cells differentiate into dormant, ovoid spores capable of withstanding harsh environmental conditions (3). However, not all cells in the population become spores. Some cells persist outside fruiting bodies as peripheral rods (4, 5), while the majority of the population undergoes cellular lysis (6–8). Hence, M. xanthus cells adopt one of three distinct fates during the developmental process. At some point poststarvation, a cell may “commit” to a particular fate irreversibly, adopting that fate even if nutrients reappear. Early studies of developmental commitment to cellular lysis and spore formation were carried out with M. xanthus before work on the signaling and gene regulatory network began. Commitment to cellular lysis was shown to occur in 25% of the population by 36 h after being placed on starvation agar (8). Commitment to the formation of optically refractile spores was also explored (9). Many of the rod-shaped cells harvested after 24 to 32 h on starvation agar appeared to be in an intermediate state, failing to form spores if placed directly into nutrient medium, but forming spores if incubated in magnesium phosphate buffer for 6 h prior to incubation in nutrient medium. By 36 h poststarvation, ca. 10% of the rod-shaped cells formed spores if placed directly into nutrient medium. These early studies established that commitment to cellular lysis and spore formation occur, but insight into the molecular mechanisms that might be involved required a better understanding of the signaling and gene regulatory network that controls development. A complex network of signals and gene expression regulates M.

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xanthus development (10) (see Fig. S1 in the supplemental material). Expression of early developmental genes is triggered by the intracellular production of (p)ppGpp in response to starvation (11, 12). This is followed by production of the A-signal, which is a mixture of amino acids and peptides released by the activity of extracellular proteases (13, 14). A-signaling has been proposed to play a quorum-sensing role that at a high enough cell density stimulates expression of certain genes and causes cells to begin building mounds (15, 16). Like A-signal, production of extracellular C-signal depends on starvation-linked (p)ppGpp accumulation inside cells and on Asg proteins, but C-signal is produced more slowly (17, 18), accumulates on the surface of cells (19, 20), and its effects are manifest slightly later as cells align during aggregation into mounds (21–24). C-signaling stimulates expression of many genes beginning at about 6 h poststarvation (24) and is required for cells to form compact mounds that mature into fruiting bodies as spores form inside (25). Abundant contacts between cells in nascent fruiting bodies have been proposed to facilitate C-signaling, and a high level of C-signaling is necessary for expression of genes required for sporulation (26–30). Therefore, C-signaling may be involved in committing cells to spore formation. It may also be involved in commitment to cellular lysis since mutants unable to produce C-signal are defective for developmental lysis (31–33).

Received 25 March 2014 Accepted 12 May 2014 Published ahead of print 16 May 2014 Address correspondence to Lee Kroos, [email protected]. Supplemental material for this article may be found at http://dx.doi.org/10.1128 /JB.01692-14. Copyright © 2014, American Society for Microbiology. All Rights Reserved. doi:10.1128/JB.01692-14

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August 2014 Volume 196 Number 15

Downloaded from http://jb.asm.org/ on March 18, 2015 by The University of Iowa Libraries

Department of Biochemistry and Molecular Biology, Michigan State University, East Lansing, Michigan, USA

MrpC Proteolysis and Myxococcus Sporulation Commitment

MATERIALS AND METHODS Bacterial strains, plasmids, and primers. Strains, plasmids, and primers used in the present study are listed in Table S1 in the supplemental material. To generate pRR010 that was used in the RNA analysis, a 110-bp internal fragment of mrpC was amplified using the primers mrpC qPCR F and mrpC qPCR R with chromosomal DNA from M. xanthus strain DK1622 as the template. The fragment was cloned into pCR2.1-TOPO


(Invitrogen) as described by the manufacturer. The presence of the correct fragment was verified by DNA sequencing. Growth and development. Escherichia coli containing plasmids were grown at 37°C in Luria-Bertani (LB) medium supplemented with 50 ␮g of kanamycin sulfate/ml as required. M. xanthus strains were grown at 32°C in CTTYE liquid medium (1% Casitone, 0.2% yeast extract, 10 mM TrisHCl [pH 8.0], 1 mM KH2PO4-K2HPO4, 8 mM MgSO4 [final pH 7.6]) with shaking or on CTT agar plates (1.5% agar, 1% Casitone, 10 mM Tris-HCl [pH 8.0], 1 mM KH2PO4-K2HPO4, 8 mM MgSO4 [final pH 7.6]) supplemented with 40 ␮g of kanamycin sulfate/ml as required. Fruiting body development was performed in submerged culture with MC7 (10 mM morpholinepropanesulfonic acid [MOPS; pH 7.0], 1 mM CaCl2) as the starvation buffer (44). Briefly, log-phase cultures of M. xanthus growing in CTTYE liquid medium were collected by centrifugation and resuspended in MC7 at a concentration of 1,000 Klett units. Either 247 ␮l of the cell suspension plus 1.65 ml of MC7 was added to a 3.5-cmdiameter well of a six-well plastic plate or in the case of samples for RNA preparation, 1.5 ml of cell suspension plus 10.5 ml of MC7 was added to an 8.5-cm-diameter plastic petri plate. Upon incubation at 32°C, cells adhere to the bottom of the plate. At the indicated times, the culture supernatant was replaced with fresh MC7 starvation buffer or CTTYE nutrient medium supplemented with inhibitors as required. For the collection of samples, cells were scraped from the plate bottom using a sterile 1-ml plastic pipette tip, and the entire contents were aspirated into a 15-ml centrifuge tube. Samples were mixed thoroughly by vortexing for 15 s, followed by pipetting the mixture up and down fifteen times, and this was done a total of three times. For observation of the cellular shape change, the tubes were then left undisturbed for 5 to 10 min to allow cell aggregates to settle to the bottom of the tube. The aggregates were then removed for microscopic observation. For measurement of sonication resistance, 400 ␮l of the thoroughly mixed sample was transferred to a microcentrifuge tube, sonicated using a model 450 sonifier (Branson) at output setting 2 for 10-s intervals three times with cooling on ice in between, and ovoid spores were counted microscopically using a Neubauer counting chamber. To quantify mature spores that are heat- and sonication-resistant and capable of germination, 400 ␮l of the thoroughly mixed sample was heated at 50°C for 60 min before sonication as described above; the sample was then serially diluted 10-fold in MC7 buffer, and 100-␮l samples were spread onto CTT soft (1.0%) agar plates. Colonies were counted after incubation at 32°C for 5 to 7 days. Microscopy. Low-resolution images of fruiting bodies were obtained with a Leica Wild M8 microscope equipped with an Olympus E-620 digital camera. High-resolution images of presumed nascent fruiting bodies to visualize cellular shape change were obtained with an Olympus BX51 microscope using a differential interference contrast filter and a 100⫻ oil immersion objective lens and equipped with an Olympus DP30BW digital camera. Total protein concentration and immunoblot analysis. Thoroughly mixed developmental samples were prepared as described above and stored at ⫺20°C in 400-␮l aliquots. One aliquot was used to determine the total protein concentration by sonicating the sample as described above, centrifuging it at 10,000 ⫻ g for 1 min to sediment cell debris, and using the supernatant in a Bradford (45) assay (Bio-Rad Laboratories). A second aliquot was mixed with an equal volume of 2⫻ sample buffer (0.125 M Tris-HCl [pH 6.8], 20% glycerol, 4% sodium dodecyl sulfate [SDS], 0.2% bromophenol blue, 0.2 M dithiothreitol) and boiled for 5 min. Samples (equal amounts of total protein; typically, 1 ␮g) were subjected to SDSPAGE and immunoblotting as described previously (46). For the detection of MrpC and MrpC2, anti-MrpC antibodies were used at a 1:10,000 dilution. Anti-FruA antibodies were used at a 1:2,000 dilution. Proteinantibody complexes were detected using horseradish peroxidase-conjugated goat anti-rabbit immunoglobulin G (Bio-Rad) at 1:5,000 dilution, and chemiluminescence (Western Lightning; Perkin-Elmer) according to the manufacturers’ instructions. Signals were detected using a LAS 3000 imager (Fujifilm) with exposure times short enough to ensure the signals

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Recently, several C-signal-dependent genes have been shown to be under the combinatorial control of two transcription factors, MrpC and FruA, which bind cooperatively in promoter regions (34–37). Different arrangements of binding sites with different affinities influence the timing of promoter activity and its dependence on C-signaling. It was not surprising to find that FruA directly regulates C-signal-dependent genes since FruA was known to be similar to DNA-binding response regulators of two-component signal transduction systems and mutations in fruA interfered with the response to C-signaling (38–40). On the other hand, MrpC had not been implicated as a direct regulator of C-signaldependent genes, although it was known to be similar to transcription factors of the cyclic AMP receptor protein family (41) and an N-terminally truncated form of MrpC appeared to directly activate fruA transcription (42). The discovery that MrpC directly regulates C-signal-dependent genes led to the hypothesis that the abundance of MrpC is responsive to nutrient conditions as development proceeds (35) because several mechanisms link MrpC activity to starvation (see Fig. S1 in the supplemental material). This hypothesis predicts that nutrient addition to developing cells would diminish MrpC activity. Only if cells were both starving (to allow sufficient MrpC activity) and efficiently C-signaling within a nascent fruiting body (to allow sufficient FruA activity) would C-signal-dependent genes be expressed, committing the cell to form a spore. Consistent with the possible involvement of MrpC, C-signal, and FruA in cell fate determination, all three proteins were found to be more abundant in aggregating cells than in nonaggregating cells of a starving population (33). Also, mutants that accumulate MrpC and FruA, or C-signal, earlier than normal during development exhibit premature aggregation and sporulation (27, 43). We report here that many M. xanthus cells commit to shape change and sonication resistance between 24 and 30 h poststarvation. During this critical period, MrpC is present and FruA accumulates, but if nutrients are added before or during this period, MrpC declines rapidly. The decline is more rapid than observed when chloramphenicol is added to inhibit protein synthesis, indicating that MrpC undergoes nutrient-regulated proteolysis (see Fig. S1 in the supplemental material). The mrpC transcript declines less rapidly than MrpC protein, but transcriptional control also appears to be involved in the response to nutrients. Importantly, the decline of MrpC halts the expression of direct target genes important for sporulation. Experiments with cell extracts implicate ATP-independent metalloprotease activity in nutrientregulated turnover of MrpC, which appears to provide a mechanism for sensing whether nutrients reappear before the critical period around 24 to 30 h poststarvation. If starvation persists, the MrpC level remains high and, together with a high level of activated FruA due to C-signaling between cells in close proximity in the nascent fruiting body, the two transcription factors coordinately regulate genes important for commitment to the spore fate.

Rajagopalan and Kroos

were not saturated. Signal intensities were quantified using Multi-Gauge (Fujifilm) software. Protein half-lives were calculated assuming a firstorder kinetic degradation reaction as described previously (47). Signal intensities, after subtracting the background, were normalized to the relevant intensity at t ⫽ 0, and the natural log of the resulting values was plotted versus minutes after nutrient, chloramphenicol, or protease inhibitor/sonication treatment. The slope of a linear fit of the data was used to calculate the protein half-life. For MrpC, only the upper band corresponding to the full-length protein was quantified and used to calculate the half-life since lower bands could be MrpC2 and/or other proteolytic fragments of MrpC. RNA extraction and analysis. Immediately prior to harvesting the culture, its supernatant was replaced with a mixture of 0.5 ml of RNase stop solution (5% phenol [pH ⬍ 7] in ethanol) (48) and 4.5 ml of MC7 buffer. Cells were scraped from the bottom of the plate as described above, and the entire contents were aspirated into a 15-ml centrifuge tube, which was subjected to flash freezing in a dry ice-ethanol bath and stored at ⫺80°C. After thawing and centrifugation at 8,700 ⫻ g for 10 min at 4°C, RNA was extracted using the hot-phenol method and digested with DNase I (Roche) as described previously (43). Total RNA (1 ␮g) was used to synthesize cDNA in a 20-␮l reaction with Superscript III reverse transcriptase (Lifetech) and random primers (Promega) according to the manufacturers’ instructions. Quantitative reverse transcription-PCR (qRTPCR) was performed in 20-␮l reactions using SYBR green PCR master mix (Applied Biosystems), 2 ␮l of 10-fold-diluted cDNA product, and primers (20 pmol each) in a model 7300 real-time PCR system (Applied Biosystems). qRT-PCRs were performed in triplicate for each biological replicate. Control reactions were done with reverse transcriptase omitted at the cDNA synthesis step prior to the qPCRs. A standard curve for each set of qPCRs was generated using 10-fold serial dilutions of plasmid pRR010 or DK1622 chromosomal DNA, and gene expression was quantified using the relative standard curve method (user bulletin 2; Applied Biosystems). 16S rRNA was used as the internal standard. The primers used to quantify mrpC, dev, and 16S rRNA transcripts are listed in Table S1 in the supplemental material. Protein degradation assays. For the experiments shown in Fig. 7 and for Fig. S5 and S7 in the supplemental material, we modified a procedure described previously (17). At 18 or 24 h poststarvation in submerged culture, the supernatant was replaced with fresh starvation buffer or nu-

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trient medium as described above for 15 min at 32°C. The supernatant was then replaced with phosphate-buffered saline, and samples were collected immediately and mixed thoroughly as described above. Samples (400 ␮l) were transferred to microcentrifuge tubes, and protease inhibitors were added as follows: Complete PIC (Roche) at the concentration recommended by the manufacturer, EDTA (5 mM), Pefabloc (0.4 mM), aprotinin (1.5 ␮M), leupeptin (20 ␮M), E-64 (28 ␮M), or pepstatin (40 ␮M). Samples were sonicated as described above. Alternatively, apyrase (2 ␮M) was added after sonication. After 0, 15, 45, and 105 min at 25°C, 40 ␮l of sample was mixed with 20 ␮l of 3⫻ sample buffer (containing PIC) and heated at 96°C for 5 min. Samples were stored at ⫺20°C. Equal volumes of the samples were subjected to SDS-PAGE and immunoblot analysis. For the experiment shown in Fig. S6 in the supplemental material, samples were prepared at 24 h poststarvation as described above with sonication in the absence of protease inhibitor or in the presence of EDTA (5 mM). To one set of extracts prepared in the absence of protease inhibitor, apyrase (2 ␮M) was added, and the sample was placed at 4°C for 10 min. Recombinant His10-MrpC (60 ng in 5 ␮l) or His10-MrpC2 (60 ng in 5 ␮l), expressed in E. coli and purified as described previously (49, 50), was added to 5 ␮l of a 2-fold dilution series of extract in phosphate-buffered saline, and the 10-␮l reactions were incubated at 25°C for 10 min, mixed with 5 ␮l of 3⫻ sample buffer, heated, stored, and subjected to SDS-PAGE (1 ␮l per well) and immunoblot analysis as described above.

RESULTS

Commitment to fruiting body formation and cellular shape change. To investigate commitment to morphological changes during M. xanthus development, we used a submerged culture in which growing cells are resuspended in starvation buffer; they attach to the bottom of the culture vessel and undergo development (44). If left undisturbed, aggregation centers resolved into small mounds by 18 h (Fig. 1A). Darkening of some nascent fruiting bodies was observed by 36 h poststarvation, with more growing darker by 42 h. A similar developmental pattern was observed if the culture supernatant was replaced with fresh starvation buffer at different times (Fig. 1B). In this and subsequent figures, 18S-24, for example, means the culture supernatant was replaced with



FIG 1 Effect of nutrient addition on fruiting body formation. (A) Fruiting body formation by M. xanthus DK1622 under submerged culture conditions. Aggregates form compact fruiting bodies by 24 h (an arrow points to one), which darken by 36 to 42 h. Spaces between fruiting bodies become clearer over time. The culture supernatant was replaced with fresh starvation buffer (designated “S”) (B) or nutrient medium (designated “N”) (C) at 18, 24, 30, or 36 h and observed at the indicated times (e.g., 18S-24 means the culture supernatant was replaced with fresh starvation buffer at 18 h and the culture was photographed at 24 h). Bar, 100 ␮m. Similar results were observed in at least two biological replicates.



added. Other cells appear to be committed by 18 h to subsequently lower their protein concentration or undergo lysis. If starvation persists during the period between 24 and 30 h, many cells in nascent fruiting bodies morph into spores (or are committed to do so by 36 h). Since spores are nonmotile, nascent fruiting bodies persist and darken even if nutrients are added at 30 h. A computational simulation suggested that formation of nonmotile spores stabilizes nascent fruiting bodies (51). Commitment to sonication resistance and formation of mature spores. Mature spores are resistant to heat and sonication, and they are capable of germinating in the presence of nutrients and growing to produce a colony. We reasoned that cells might change shape and become sonication resistant before they become fully mature spores. To quantify spores that were resistant to sonication but not necessarily fully mature, we sonicated submerged culture samples to disperse spores so they could be counted microscopically. Sonication-resistant spores were present by 30 h poststarvation and increased ⬃10-fold by 36 h (Fig. 3A, no replacement), a result consistent with our observations of cellular shape change reported above. The number of sonication-resistant spores rose more gradually thereafter. Replacing the culture supernatant with fresh starvation buffer did not affect development of sonication resistance (Fig. 3A, starvation). In striking agreement with our observations of cellular shape change reported above, replacing the culture supernatant with nutrient medium at 18 h but not at 24 h prevented cells from becoming sonication resistant (Fig. 3A, nutrient). Therefore, many cells that are not yet sonication resistant at 24 h are nevertheless committed to become sonication-resistant spores by 30 h (Fig. 3A, 24N-30). Interestingly, the addition of nutrients at 24 h blocked the large increase in the number of sonication-resistant spores between 30 and 36 h (24N-36). The addition of nutrients at 30 h did not block this increase, indicating that a large number of cells that are not yet sonication resistant at 30 h are committed to become so by 36 h (30N-36). The number of sonication-resistant spores decreased slightly after prolonged exposure to nutrients (30N-54 and 36N54), perhaps due to a small amount of spore germination. To measure the number of mature spores, samples were subjected to heat treatment and sonication, followed by plating on nutrient agar to allow spore germination and colony formation. Mature spores were observed starting at 36 h poststarvation, and the number gradually went up more than 1,000-fold by 96 h (Fig. 3B, no replacement). Formation of mature spores lags behind development of sonication resistance (Fig. 3A, no replacement). The addition of nutrients at 18 h poststarvation prevented the development of mature spores by 36 h (Fig. 3B, 18N-36). However, the addition of nutrients at 24 h did not stop the formation of mature spores when analyzed at 48 h (24N-48), although the number of mature spores formed was much lower than the number at 48 h in the undisturbed culture (48) or after replacement of the culture supernatant with fresh starvation buffer (24S-48). Thus, at least some cells in a starving population are committed by 24 h to becoming mature spores. Similarly, nutrient addition at 30 h did not stop the formation of a small number of mature spores by 54 h (30N-54). Nutrient addition at 36 h allowed more cells to form mature spores by 42 h (36N-42), resulting in only ⬃10-fold less than in the undisturbed culture (42). Waiting longer after nutrient addition at 36 h resulted in slight decreases in the number of mature spores (36N-48 and 36N-54), perhaps because a small number of spores begin to germinate in the presence of nutrients.

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fresh starvation buffer at 18 h, and the culture was photographed at 24 h. The area between developing fruiting bodies became clearer over time, presumably due to movement of cells into fruiting bodies and cellular lysis. The total protein concentration of developing cultures showed a substantial decrease as time progressed (see Fig. S2A in the supplemental material), with a 6-fold decrease by 30 h and a 50-fold decrease by 66 h. This suggests that developing cells contain less protein or they lyse, since only a small proportion of the input cells form spores (that would not release protein upon sonication) inside fruiting bodies. Strikingly, if the culture supernatant was replaced with nutrient medium at 18 or 24 h poststarvation, aggregates failed to progress to compact, darkened fruiting bodies and instead became less compact over time (Fig. 1C, compare 24N-30 and 24N-36). Also, the areas between fruiting bodies failed to clear normally. On the other hand, nutrient addition at 30 or 36 h poststarvation did not stop the formation of compact, darkened fruiting bodies (Fig. 1C), indicating that a starving culture commits to fruiting body formation by 30 h. Nutrient addition at all time points tested (18, 24, 30) did not prevent the eventual reduction of total protein concentration, although it did not decrease as rapidly (see Fig. S2B in the supplemental material). These results suggest that some cells in the population commit to lowering their protein concentration or undergoing lysis before 18 h, but nutrient addition delays these events. Delayed cellular lysis and movement of cells out of fruiting bodies may account for the failure of areas between nascent fruiting bodies to clear when nutrients were added at 18 or 24 h. Commitment to cellular shape change was examined by microscopic observation after gentle dispersion. Developing cells were scraped from the substratum, along with the culture supernatant, and samples were subjected to mild dispersion by vortexing and pipetting. This treatment was designed to preserve cells in the process of converting from rods into spores but perhaps not yet resistant to sonication. Indeed, thickened rods and ovoid spores were first observed at 24 h poststarvation within or at the edge of large cell aggregates (presumably nascent fruiting bodies) (Fig. 2A). By 30 h a larger percentage of cells had changed shape, and by 36 h the majority of cells in aggregates had morphed into spores. Darkening of cell aggregates was observed at 36 h, a finding consistent with the darkening of nascent fruiting bodies at 36 h seen under lower magnification (Fig. 1A). Replacing the culture supernatant with fresh starvation buffer did not affect the normal progression of cellular shape change (Fig. 2B and D). In contrast, replacing the culture supernatant with nutrient medium at 18 h prevented cellular shape change (Fig. 2C and data not shown for later times after nutrient addition). However, the addition of nutrients at 24 h did not stop a large proportion of the cells in some aggregates from changing shape, although a delay was observed, with few cells having changed shape by 30 h (Fig. 2E, 24N-30) but many cells having changed shape by 36 h (Fig. 2E, 24N-36, left half). Therefore, at least some cells in a developing culture commit to cellular shape by 24 h. Other aggregates lacked spores but contained some rods and what appeared to be cell debris, as if some cells had undergone lysis (Fig. 2E, 24N-36, right half). This is consistent with the observations noted above at lower magnification. We conclude from these data that in submerged culture under our conditions a few cells are changing from rods to spores by 24 h poststarvation, and many more cells are committed by this time to subsequently change their shape by 36 h, even if nutrients are


However, spore germination cannot account for the relatively small numbers of mature spores in the 24N-48, 30N-54, and 36N-54 samples (Fig. 3B, nutrient), since germination would render cells sensitive to sonication, and the number of sonicationresistant spores remained high in those samples (Fig. 3A, nutrient). We conclude that commitment to formation of sonicationresistant spores, like commitment to cellular shape change, occurs primarily during a critical period at 24 to 30 h poststarvation. In contrast, only a small number of cells commit to forming a mature spore during this period, and even at 36 h poststarvation, when a large number of sonication-resistant spores have formed (Fig. 3A, no replacement or starvation), very few of these spores are committed to maturation (Fig. 3B, nutrient). The number of mature

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spores increases sharply during the period between 36 and 48 h poststarvation (Fig. 3B, no replacement or starvation); we were unable to define a shorter period of commitment to spore maturation. Therefore, we focused on molecular events before and during the critical period between 24 and 30 h poststarvation when cells commit to subsequent shape change and sonication resistance. MrpC and FruA are required for commitment to development and persist through the period of commitment to shape change and sonication resistance. To begin testing possible involvement of MrpC and FruA in commitment to development, we examined mutants under our conditions of submerged culture. Neither the mrpC mutant nor the fruA mutant formed compact nascent fruiting bodies (see Fig. S3A in the supplemental mate-



FIG 2 Effect of nutrient addition on cellular shape change. (A) Cellular shape change in M. xanthus DK1622 under submerged culture conditions. Developing cultures were subjected to mild dispersion and examined microscopically. Photos show densely packed cell aggregates presumed to be nascent fruiting bodies and insets show the edge of a cell aggregate. Arrows indicate thickened rods or ovoid spores. The culture supernatant was replaced with fresh starvation buffer (designated “S”) (B) or nutrient medium (designated “N”) (C) at 18 or 24 h and observed at the indicated time (e.g., 24N-36 means the culture supernatant was replaced with nutrient medium at 24 h and the culture was photographed at 36 h). The panel labeled 24N-36 shows a cell aggregate with many spores (left half) and a cell aggregate lacking spores but with rods and what appears to be cell debris (right half). Bar, 5 ␮m. Similar results were observed in at least two biological replicates.


under submerged culture conditions for the time indicated by the first number. The culture supernatant was left undisturbed or replaced with fresh starvation buffer (designated “S” and indicated by the bracket labeled “starvation”) or nutrient medium (designated “N” and indicated by the bracket labeled “nutrient”), and then the culture was harvested for the measurement of sonication-resistant or mature spores at the time indicated by the second number (e.g., 24N-36 means the culture supernatant was replaced with nutrient medium at 24 h and the culture was harvested at 36 h). Values (log10) are the average of at least three biological replicates, and error bars represent one standard deviation from the mean.

rial), as expected based on previous reports of their developmental phenotypes on starvation agar plates (41, 52). Neither mutant initiated detectable cellular shape change (see Fig. S3B in the supplemental material). Hence, the absence of MrpC or FruA prevented detectable commitment to development. Also, mature spores failed to form (data not shown), as reported previously (38, 41, 52). Next, we examined MrpC and FruA levels in wild-type cells developing under our submerged culture conditions. The levels of these proteins in different “wild-type” backgrounds have been examined previously under various conditions of development (38, 39, 43, 50), but it was important to measure the levels in our wild-type strain under our conditions of development for comparison to the results described above. Immunoblots showed that MrpC and its smaller form, MrpC2, were not detected at 0 h, when cells were sedimented from nutrient medium and resuspended in starvation buffer. Both forms were detected at 12 h and increased by 15 h, with MrpC persisting through the period of commitment to shape change and sonication resistance (i.e., 24 to 30 h poststarvation), at least until 42 h, and MrpC2 decreasing reproducibly between 30 and 36 h (Fig. 4A). FruA was barely detectable at 12 h, and its accumulation lagged behind that of MrpC by about 6 h (Fig. 4B), as expected since transcription of fruA depends on MrpC (42). The FruA level rose steadily throughout the period of commitment. Thus, both MrpC and FruA accumulate prior to the time we observed commitment to cellular shape


change (24 h) (Fig. 2) and persist at least until 42 h (Fig. 4), the period during which cells in the developing population commit to forming compact nascent fruiting bodies (Fig. 1) and sonicationresistant and mature spores (Fig. 3). MrpC declines rapidly upon nutrient addition to developing cells. What roles do MrpC and FruA play in commitment to development? The discovery that the two proteins cooperatively regulate developmental genes suggested that positional information through C-signaling and FruA is integrated with nutrient sensing by regulating the abundance of MrpC (35). Therefore, we hypoth-

FIG 4 (A and B) Immunoblot analysis of MrpC (A) and FruA (B) during development. M. xanthus DK1622 was starved under submerged culture conditions. Samples were collected at the indicated times and equal amounts of protein were analyzed by immunoblot using anti-MrpC or anti-FruA antibodies. Intervening lanes were removed between the 36 and 42 h samples in panel A. Representative immunoblots are shown. Similar results were observed in at least two biological replicates.

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FIG 3 (A and B) Effect of nutrient addition on development of sonication resistance (A) and mature spores (B). M. xanthus DK1622 was subjected to starvation


esized that the role of MrpC in commitment is to have its abundance regulated in response to nutrients. This hypothesis predicts that MrpC would decline rapidly upon nutrient addition to developing cells. In agreement with our hypothesis, immunoblots showed a rapid decline in MrpC upon nutrient addition at 6, 12, 18, 24, or 30 h poststarvation (Fig. 5). Nutrient addition caused the MrpC level to decrease faster than addition of chloramphenicol (Cm), an inhibitor of protein synthesis, indicating that nutrient addition triggers MrpC turnover rather than simply halting MrpC synthesis. The signals were quantified and used to calculate the half-life of MrpC (Fig. 5, right). Reproducibly, the MrpC halflife was shorter after nutrient addition than after Cm addition at all times tested (compare t1/2 for N with S⫹Cm). Moreover, the simultaneous addition of nutrient and Cm at 24 h poststarvation caused MrpC to decline at least as rapidly as addition of nutrient alone (Fig. 5D, compare N⫹Cm to N). Therefore, protein synthesis is not required for nutrients to activate one or more preexisting proteases to rapidly degrade MrpC. We note that although nutrient-regulated proteolysis of MrpC occurs at 24 h poststarvation (Fig. 5D), it does not stop a large proportion of the cells in some aggregates from subsequently changing shape by 36 h (Fig. 2E, 24N-36, left panel), presumably because MrpC-dependent genes required for cellular shape change had already been expressed. Likewise, nutrient-regulated MrpC proteolysis occurs at 30 h (Fig. 5E), but does not prevent formation of compact nascent fruiting bodies that subsequently darken by 42 h (Fig. 1C, 30N-42) and does not prevent a

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FIG 5 Effect of nutrient addition on the MrpC level during development. M. xanthus DK1622 was starved under submerged culture conditions, and after 6 (A), 12 (B), 18 (C), 24 (D), or 30 h (E) the culture supernatant was replaced with fresh starvation buffer (S), nutrient medium (N), fresh starvation buffer with 200 ␮g of chloramphenicol/ml (S⫹Cm), or nutrient medium with 200 ␮g of chloramphenicol/ml (N⫹Cm). At the indicated times (in minutes) after replacement, cells were harvested and equal amounts of total protein were analyzed by immunoblotting with anti-MrpC antibodies. Filled and unfilled arrowheads indicate MrpC and MrpC2, respectively. Intervening lanes were removed from the image in panel A. Representative immunoblots are shown. The half-life in minutes (t1/2 [in minutes]) of MrpC after the indicated treatment is shown to the right. A similar ratio of the half-life after nutrient treatment to that after chloramphenicol treatment was observed in at least two biological replicates.

large increase in the number of sonication-resistant cells by 36 h (Fig. 3A, 30N-36). Presumably, MrpC-dependent genes required for these morphological and physical changes are expressed between 24 and 30 h. It appears that MrpC must persist after 30 h in order for genes to be expressed that are important for mature spores to form in large numbers after 36 h (Fig. 3B). In this context, it is worth noting that very little of the smaller form, MrpC2, was present after 30 h (Fig. 4A), suggesting that MrpC2 is not required for spore maturation. We also note that very little MrpC2 was present at any of the times tested in our nutrient addition experiments (Fig. 5, unfilled arrowheads). In these experiments, the culture supernatant was replaced with fresh starvation buffer prior to collecting samples, whereas the culture was undisturbed prior to collecting samples for the experiment shown in Fig. 4. Although replacement with fresh starvation buffer greatly diminished MrpC2, there was no apparent effect on fruiting body formation (Fig. 1B), cellular shape change (Fig. 2B and D), or formation of sonication-resistant or mature spores (Fig. 3, starvation). MrpC2 has been reported to bind sites in the mrpC and fruA promoter regions with higher affinity than MrpC (50), but individual roles of the two proteins during development have not been dissected. The level of FruA also decreased after nutrient and/or Cm addition (Fig. S4), but the effect of nutrient addition was not as profound as for MrpC (Fig. 5). When nutrients were added at 18 h poststarvation, MrpC was nearly undetectable after 1 h (Fig. 5C), but FruA was still visible (Fig. S4A). Nutrient addition at 24 h poststarvation caused a more rapid and complete loss of MrpC (Fig. 5D) than FruA (Fig. S4B). At both times, nutrient addition caused a more rapid loss of MrpC than Cm addition (Fig. 5C and D, compare N to S⫹Cm and the corresponding t1/2 values), but this was not the case for FruA (see Fig. S4 in the supplemental material). The half-life of FruA was similar after nutrient or Cm addition at 18 h, and it was longer after nutrient addition than after Cm addition at 24 h. Therefore, we cannot be certain whether FruA is subject to nutrient-regulated proteolysis. Since transcription of fruA depends on MrpC (42), synthesis of FruA is expected to decline rapidly upon nutrient addition, although probably not as rapidly as upon Cm addition. We conclude from the data in Fig. S4 in the supplemental material that FruA might be subject to nutrient-regulated proteolysis, in addition to its expected decline as fruA transcription ceases due to the declining MrpC level. Since the decline of MrpC appeared to be the primary response of developing cells to nutrient addition, we focused on this response and its consequences in subsequent experiments. The mrpC transcript increases, then decreases, upon nutrient addition to developing cells. To explore further the response of developing cells to nutrient addition prior to commitment of most cells to sonication resistance, we measured mrpC transcript levels after nutrient addition to 24-h developing cells (Fig. 6A). Nutrient addition resulted in more than a 3-fold increase in the mrpC transcript level after 15 min (Fig. 6, N⫹15), followed by a decline to the original level after 30 min (N⫹30), and a further decline to about half the original level by 120 min (N⫹120). As a control, the culture supernatant was replaced with fresh starvation buffer rather than nutrient medium. Surprisingly, the mrpC transcript level increased ⬃2-fold after 15 min (S⫹15) and remained high after 30 min (S⫹30), as if an inhibitor of mrpC transcript accumulation had been removed, followed by a decline to the original level after 60 min (S ⫹ 60) or 120 min (S⫹120). In con-


and the dev transcript level (B) during development. M. xanthus DK1622 was starved under submerged culture conditions, and after 24 h the culture supernatant was replaced with fresh starvation buffer (S), nutrient medium (N), fresh starvation buffer with 50 ␮g of rifampin/ml (S⫹Rif), or nutrient medium with 50 ␮g of rifampin/ml (N⫹Rif). At the indicated times (in minutes) after replacement (e.g., “S⫹15” indicates that the culture supernatant was replaced with fresh starvation buffer and the culture was harvested 15 min later), cultures were harvested, and RNA was isolated and subjected to qRT-PCR analysis to determine the mrpC or dev transcript level. Values are the average of three technical replicates for at least three biological replicates and are reported relative to the untreated 24-h sample (labeled “None”). Error bars indicate one standard deviation from the mean.

trast, if the fresh starvation buffer contained rifampin (Rif), which blocks transcription, the mrpC transcript level declined ⬃10-fold after 15 min (S⫹Rif⫹15) and was barely detectable after 30 min (S⫹Rif⫹30). The result with rifampin indicates that the mrpC transcript is unstable in 24-h developing cells. Nutrient addition, like the addition of fresh starvation buffer, may remove an inhibitor of mrpC transcript accumulation, resulting in the observed increase after 15 min. However, the mrpC transcript level remains high 30 min after the addition of fresh starvation buffer but declines to the original level 30 min after nutrient addition, indicating that nutrients inhibit mrpC transcript accumulation between 15 and 30 min after their addition. The inhibition, however, returns the mrpC transcript to its original level, which does not explain the observed decrease in MrpC protein level by 30 min after nutrient addition to 24-h developing cells (Fig. 5D), a finding consistent with our interpretation above that nutrients activate one or more proteases that rapidly degrade MrpC. The rapid decline of MrpC may account for the decline of the mrpC transcript, since MrpC has been reported to positively autoregulate its own expression, likely at the transcriptional level (41). Since the primary effect of nutrients prior to the commitment of most cells to sonication resistance appeared to be rapid degradation of MrpC, we focused on gaining further insight into this process. Nutrient addition halts expression of MrpC target genes important for sporulation. To test whether nutrient-regulated proteolysis of MrpC affects transcription of its target genes that code


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FIG 6 (A and B) Effect of nutrient addition on the mrpC transcript level (A)

for proteins important for commitment to sporulation, we measured transcript levels from the dev operon after nutrient addition to 24-h developing cells (Fig. 6B), when most cells have not yet committed to sonication resistance (Fig. 3A). MrpC and FruA cooperatively bind in the promoter region of the dev operon and appear to directly activate transcription (53; A. Campbell, P. Viswanathan, B. Son, T. Barrett, and L. Kroos, unpublished data). Products of the dev operon have been shown to be important for sporulation (54–56) and to positively regulate expression of genes that play a major role in maintenance of cellular shape change (57–59). Under our submerged culture conditions, null mutants of devT or devR failed to change cell shape or become sonication resistant by 72 h poststarvation (data not shown). We measured dev transcript levels using qPCR with primers that would amplify from within devT near its 3= end to within devR near its 5= end (Fig. 6B). At 24 h poststarvation, nutrient addition resulted in a steady decline in dev transcripts, which decreased by ⬃3-fold by 120 min (N⫹120) to a level significantly below the starting value (P ⫽ 0.02, Student t test). In the control, where the culture supernatant was replaced with fresh starvation buffer, the dev transcript level remained similar to the starting value. In contrast, if the fresh starvation buffer contained rifampin, the dev transcripts were barely detectable after 15 min (S⫹Rif⫹15), indicating that the transcript is unstable in 24-h developing cells. We conclude that nutrientregulated proteolysis of MrpC halts expression of the dev operon whose products are important for commitment to sporulation. The decline in dev transcripts is less rapid after nutrient addition than after rifampin addition, suggesting that transcription of dev continues for a short time after nutrient addition until MrpC and FruA levels are too low to support further expression. Nutrient-regulated proteolysis of MrpC may involve ATPindependent metalloprotease activity. We investigated the possibility that BsgA (also called LonD) is involved in rapid turnover of MrpC after nutrient addition to developing cells. BsgA is an ATP-dependent serine protease shown previously to be required for development (60, 61). Since a bsgA mutant fails to produce MrpC2 during development, it was suggested that BsgA may be responsible for proteolytic conversion of MrpC to MrpC2 (50) (see Fig. S1 in the supplemental material). Under our conditions of submerged culture development, a bsgA mutant shows a severe developmental defect similar to mrpC or fruA mutants, failing to form fruiting bodies or initiate cellular shape change (see Fig. S3A and S3B in the supplemental material). Immunoblots revealed that addition of nutrients to the bsgA mutant at 18 h poststarvation resulted in a rapid decline of MrpC, more rapid than the addition of Cm (see Fig. S3C in the supplemental material), as observed for wild type (Fig. 5C). The half-life of MrpC was shorter after nutrient addition than after Cm addition (see Fig. S3C, right, in the supplemental material), so BsgA is not required for nutrient-regulated proteolysis of MrpC, but the half-lives were longer in the bsgA mutant than in wild type (Fig. 5C, right), suggesting that BsgA directly or indirectly impacts MrpC proteolysis. As reported previously for a different bsgA mutant (50), we did not detect conversion of MrpC to MrpC2 in the bsgA mutant we used (see Fig. S3C in the supplemental material). To identify the type of protease that might be involved in rapid turnover of MrpC after nutrient addition to developing cells, various protease inhibitors were tested for the ability to slow the degradation of MrpC in cell extracts. Immunoblots showed that MrpC was degraded rapidly in sonic extracts of developing cells


of MrpC in extracts of developing cells. M. xanthus DK1622 was starved under submerged culture conditions, and after 18 h the culture supernatant was replaced with fresh starvation buffer (S) or nutrient medium (N) for 15 min. Cultures were harvested and protease inhibitor(s) was added prior to sonication, or no protease inhibitor was added as a control (⫺). After sonication, samples were taken at the indicated times (in minutes). Apyrase was added immediately after sonication. Equal sample volumes were analyzed by immunoblotting with anti-MrpC antibodies. Gaps in the image indicate samples analyzed on separate immunoblots. Representative immunoblots are shown. The half-life in minutes (t1/2 [in minutes]) of MrpC under the indicated condition is shown to the right. A similar ratio of the half-life in the presence of PIC to that with EDTA was observed in at least two biological replicates.

prepared at 18 h (Fig. 7) or 24 h (see Fig. S5 in the supplemental material) poststarvation, covering time points both before and at the beginning of the critical period of commitment to sporulation. Loss of MrpC appeared to be more rapid when the culture supernatant was replaced with fresh starvation buffer for 15 min prior to sonication [designated S(⫺), with the minus indicating the absence of protease inhibitors] than when it was replaced with nutrient medium [designated N(⫺)]. We expected the opposite to be the case since nutrient addition caused MrpC to decline rapidly in developing cells (Fig. 5); however, we loaded equal volumes rather than equal amounts of total protein in the experiments shown in Fig. 7 and see Fig. S5 in the supplemental material, and there could be a larger burst of MrpC synthesis shortly after nutrient addition than after starvation buffer addition. In any case, the half-life of MrpC could not be determined because it was too unstable in sonic extracts without protease inhibitor. The addition of a protease inhibitor cocktail (PIC) during sonication greatly stabilized full-length MrpC in samples to which nutrients had been added before sonication (N⫹PIC), and to a lesser degree in samples to which fresh starvation buffer had been added (S⫹PIC) (Fig. 7 and see Fig. S5 in the supplemental material). The greater stabilization of MrpC in extracts of developing cells to which nutrients had been added suggests that protease activity is altered in these cells. Perhaps nutrients activate a protease that is effectively inhibited by the cocktail, and this directly or indirectly protects MrpC from attack by other proteases. Since the PIC was a mixture of protease inhibitors at proprietary concentrations, we tested individual protease inhibitors at standard concentrations. Only the metalloprotease inhibitor EDTA stabilized MrpC, albeit typically not quite as well as the PIC, as judged from the half-lives (Fig. 7 and see Fig. S5 in the supple-

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DISCUSSION

Our examination of the consequences of adding nutrients to developing M. xanthus revealed progressive commitment to differentiation and established a foundation for investigating the molecular basis of commitment to particular cell fates. At between 18 and 24 h poststarvation, mounds become well defined, a few cells begin to change shape, and some cells commit to subsequent shape change and sonication resistance. Nevertheless, nutrient addition at 24 h poststarvation still profoundly changes the developmental outcome, resulting in 10-fold-less sonication-resistant spores (as measured 6 or 12 h later) than if starvation persists until 30 h. Hence, the period from around 24 to 30 h poststarvation, when nascent fruiting bodies become well defined and cells are close packed in mounds, is a critical period for commitment to cellular shape change and sonication resistance. During this period, short-range C-signaling is proposed to reach a maximum and stimulate expression of genes required for sporulation (26– 30). The transcription factor FruA was known to be involved in the response to C-signaling (38–40), and yet each of several C-signal-dependent genes or operons studied was found to be under coordinate control by MrpC (34–37). Here, we have shown that MrpC is rapidly degraded upon nutrient addition to developing cells before and during the critical period of commitment, halting expression of direct target genes in the dev operon that are important for sporulation (see Fig. S1 in the supplemental material). Therefore, we propose that nutrient-regulated proteolysis of MrpC provides a mechanism for monitoring whether starvation persists and halting commitment to sporulation if nutrients reappear before the critical period from around 24 to 30 h poststarvation. Advantages of delayed, stepwise commitment to development. Sporulation is a costly decision both in terms of lost time for reproduction and in terms of resources spent to produce a cell capable of surviving a long period without nutrients in a harsh environment. Bacillus subtilis and related endospore formers de-



FIG 7 Effects of nutrient addition and protease inhibitors on the degradation

mental material). Inhibitors of other types of proteases did not stabilize MrpC. Like PIC, EDTA stabilized full-length MrpC in samples to which nutrients had been added before sonication (N⫹EDTA) to a greater degree than in samples to which fresh starvation buffer had been added (S⫹EDTA). Apyrase, an enzyme that hydrolyzes ATP, was also tested, to see if proteolysis of MrpC depends on ATP. Addition of apyrase immediately after sonication did not stabilize MrpC (Fig. 7 and see Fig. S5 in the supplemental material). Neither did preincubation of sonic extracts with apyrase inhibit their ability to produce a discrete breakdown product of purified recombinant His10-MrpC or His10-MrpC2, although the addition of EDTA during sonication did inhibit breakdown product formation (see Fig. S6 in the supplemental material). Taken together, our results suggest that nutrient-regulated proteolysis of MrpC may involve ATP-independent metalloprotease activity. Since the FruA level also decreased after nutrient addition (see Fig. S4 in the supplemental material), we examined FruA degradation in cell extracts. As with MrpC, the addition of PIC or EDTA during sonication stabilized FruA in samples at 18 or 24 h poststarvation, and the addition of apyrase immediately after sonication did not stabilize FruA (see Fig. S7 in the supplemental material). These results suggest that proteolysis of FruA in developing cells may also involve ATP-independent metalloprotease activity.



growth and division so that differentiation proceeds even if nutrients become available. In B. subtilis, the SpoIIP and SpoIIQ proteins play key roles in commitment to endospore formation (73), but how they block growth and division is unknown. In M. xanthus, proteins that block growth and division, as well as proteins directly involved in peptidoglycan rearrangement that presumably accompanies cell shape change, remain to be identified. Regulation of MrpC as a mechanism to sense nutrients and escape commitment. As a key component of a feed-forward loop that appears to govern dev transcription, MrpC is well positioned to halt commitment to sporulation. Our results show that MrpC is rapidly degraded upon nutrient addition to developing cells before and during the critical period of commitment. Further, nutrient addition to developing cells at 24 h poststarvation resulted in a steady decline in dev transcripts to a low level within 2 h. We conclude that nutrient-regulated proteolysis of MrpC halts the expression of direct target genes important for sporulation, and we propose that this provides a rapid mechanism to escape commitment if nutrients reappear. Regulated proteolysis in bacteria is used to control adaptive, cell cycle, and developmental events by degrading or activating regulatory proteins, or by producing signals (74). During M. xanthus development, extracellular proteases produce A-signal (13, 14) and a proteolytic cascade produces C-signal (17, 75). The protease BsgA has been implicated in production of the unidentified B-signal (60) and in conversion of MrpC to MrpC2 (50) (see Fig. S1 in the supplemental material). Our results show that replacing the developing culture supernatant with fresh starvation buffer markedly diminishes MrpC2 accumulation but has no detectable effect on development, suggesting that MrpC2 need not accumulate continuously for development to occur. A bsgA mutant failed to accumulate MrpC2, as expected (50), and the level of MrpC was reduced compared to the wild type, but the addition of nutrients resulted in a rapid decline of MrpC (Fig. S3C), indicating that BsgA is not responsible for nutrient-regulated proteolysis of MrpC. Instead, our experiments involving addition of protease inhibitors to sonic extracts of developing cells implicate ATP-independent metalloprotease activity. Analysis of the M. xanthus genome (76) reveals at least 29 genes predicted to code for metalloproteases that are likely to be ATP-independent and cytosolic. These putative metalloproteases are candidates for involvement in nutrient-regulated proteolysis of MrpC. The Esp signaling system negatively regulates MrpC stability (see Fig. S1 in the supplemental material) to delay M. xanthus development (43) and has been proposed to do so by activating a protease or protease targeting factor to stimulate MrpC turnover (47). However, the signal(s) sensed by the Esp system is unknown. Transcriptional regulation of mrpC plays a secondary role in the response of developing cells to nutrient addition. Our results show that the mrpC transcript level increases transiently after nutrient addition to developing M. xanthus at 24 h poststarvation, returns to the original level, and then decreases (Fig. 6A). The decrease in transcript level occurs well after the decrease in MrpC protein level (Fig. 5D). Indeed, the loss of MrpC is expected to cause the mrpC transcript level to decrease since MrpC has been reported to positively autoregulate and to positively regulate the mrpAB operon, whose products positively regulate mrpC, forming a positive feedback loop (see Fig. S1 in the supplemental material). Hence, nutrient-regulated proteolysis destroys MrpC rapidly and

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lay commitment to sporulation by secreting toxins that enable cells to cannibalize siblings (62, 63). M. xanthus may likewise feed off lysing siblings, although whether this involves autocide production (64–66), programmed cell death (6, 33), or both, is unclear. Our results agree with previous work showing that M. xanthus does not commit to sporulation until it has been starving for about 24 h (9). The obvious advantage of delaying commitment to sporulation is that cells could resume growth if nutrients reappear. Our results suggest a temporal progression of M. xanthus in terms of commitment to differentiation, with some cells committing to shape change and sonication resistance during the critical period around 24 to 30 h, followed by gradual commitment to formation of mature spores. The advantage of gradual stepwise commitment to development may be to provide adaptability under changing environmental conditions. Single-cell analysis of gene expression and modeling of different stress conditions suggests that B. subtilis cells remain responsive to environmental fluctuations as they proceed toward commitment to endospore formation (67). Methods have not yet been developed to track gene expression in individual cells during M. xanthus fruiting body formation. However, in a recent study, aggregating cells were separated from nonaggregating cells, and MrpC, C-signal, and FruA were found to be more abundant in the aggregating cells (33). Therefore, high levels of the three proteins are correlated with the subpopulation in which cells commit to sporulation. Molecular basis of commitment to particular cell fates. In a wide variety of systems, commitment to particular cell fates is governed by key signals and gene regulators with characteristic features (68–70). For example, many are part of a positive-feedback loop that results in bistable expression with some cells in a population producing a much higher level of the key molecule than other cells. MrpC, C-signal, and FruA are in positive feedback loops (see Fig. S1 in the supplemental material). These positive-feedback loops likely result in bistable expression of MrpC, C-signal, and FruA, accounting for their higher abundance in aggregating cells than in nonaggregating cells (33). We propose that MrpC, C-signal, and FruA are key regulators of commitment to sporulation. C-signal accumulates gradually during development, and distinct thresholds have been observed for aggregation, sporulation, and gene expression (27–29). FruA mediates the response to C-signal (38) and binds cooperatively with MrpC to promoter regions of C-signal-dependent genes (34– 37), including the dev operon (56; Campbell et al., unpublished data) (see also Fig. S1 in the supplemental material). Since MrpC appears to activate transcription of fruA (42), and MrpC and FruA together appear to activate dev transcription, the expression of the dev operon appears to be controlled by a coherent feed-forward loop (71) expected to delay full expression until FruA* (i.e., FruA activated in response to C-signaling by an unknown mechanism) reaches a sufficient concentration. Products of the dev operon are important for sporulation (53–55), including cellular shape change and development of sonication resistance (the present study), and they positively regulate the exo operon, whose products are necessary to maintain spore differentiation (57, 59). Also, transcription of the exo operon appears to be directly activated by FruA* (72). So, together MrpC, C-signal, and FruA appear to activate targets, including the dev and exo operons whose products are important for committing cells in a nascent fruiting body to the spore fate. Commitment to sporulation also involves blocking


ACKNOWLEDGMENTS We thank Lotte Sogaard-Andersen and her coworkers for sharing their analysis of predicted proteases in the M. xanthus genome, Dale Kaiser and Wenyuan Shi for providing bacterial strains, and Penelope Higgs for suggesting primer sequences. This research was supported by NSF grant MCB-0744343 and by Michigan State University AgBioResearch.

REFERENCES 1. Whitworth DE. 2008. Myxobacteria: multicellularity and differentiation. ASM Press, Washington, DC. 2. Berleman JE, Kirby JR. 2009. Deciphering the hunting strategy of a bacterial wolfpack. FEMS Microbiol. Rev. 33:942–957. http://dx.doi.org /10.1111/j.1574-6976.2009.00185.x. 3. Kaiser D, Robinson M, Kroos L. 2010. Myxobacteria, polarity, and multicellular morphogenesis. Cold Spring Harb. Perspect. Biol. 2:a000380. http://dx.doi.org/10.1101/cshperspect.a000380. 4. O’Connor KA, Zusman DR. 1991. Behavior of peripheral rods and their role in the life cycle of Myxococcus xanthus. J. Bacteriol. 173:3342–3355. 5. O’Connor KA, Zusman DR. 1991. Development in Myxococcus xanthus involves differentiation into two cell types, peripheral rods and spores. J. Bacteriol. 173:3318 –3333. 6. Nariya H, Inouye M. 2008. MazF, an mRNA interferase, mediates programmed cell death during multicellular Myxococcus development. Cell 132:55– 66. http://dx.doi.org/10.1016/j.cell.2007.11.044. 7. O’Connor KA, Zusman DR. 1988. Reexamination of the role of autolysis in the development of Myxococcus xanthus. J. Bacteriol. 170:4103– 4112. 8. Wireman JW, Dworkin M. 1977. Developmentally induced autolysis during fruiting body formation by Myxococcus xanthus. J. Bacteriol. 129: 796 – 802. 9. Wireman JW. 1979. Developmental induction of Myxococcus xanthus myxospores. J. Bacteriol. 140:147–163. 10. Rajagopalan R, Sarwar Z, Garza AG, Kroos L. 2014. Developmental gene regulation, p 105–126. In Yang Z, Higgs P (ed), Myxobacteria: genomics, cellular and molecular biology. Caister Academic Press, Norfolk, United Kingdom. 11. Harris BZ, Kaiser D, Singer M. 1998. The guanosine nucleotide (p) ppGpp initiates development and A-factor production in Myxococcus xanthus. Genes Dev. 12:1022–1035. http://dx.doi.org/10.1101/gad.12.7.1022. 12. Singer M, Kaiser D. 1995. Ectopic production of guanosine penta- and tetraphosphate can initiate early developmental gene expression in Myxococcus xanthus. Genes Dev. 9:1633–1644. http://dx.doi.org/10.1101/gad.9 .13.1633. 13. Kuspa A, Plamann L, Kaiser D. 1992. Identification of heat-stable A-factor from Myxococcus xanthus. J. Bacteriol. 174:3319 –3326. 14. Plamann L, Kuspa A, Kaiser D. 1992. Proteins that rescue A-signaldefective mutants of Myxococcus xanthus. J. Bacteriol. 174:3311–3318. 15. Kuspa A, Kroos L, Kaiser D. 1986. Intercellular signaling is required for developmental gene expression in Myxococcus xanthus. Dev. Biol. 117: 267–276. http://dx.doi.org/10.1016/0012-1606(86)90369-6. 16. Kuspa A, Plamann L, Kaiser D. 1992. A-signalling and the cell density requirement for Myxococcus xanthus development. J. Bacteriol. 174:7360 – 7369.

2746

jb.asm.org

17. Konovalova A, Lobach S, Sogaard-Andersen L. 2012. A RelA-dependent two-tiered regulated proteolysis cascade controls synthesis of a contactdependent intercellular signal in Myxococcus xanthus. Mol. Microbiol. 84: 260 –275. http://dx.doi.org/10.1111/j.1365-2958.2012.08020.x. 18. Konovalova A, Wegener-Feldbrugge S, Sogaard-Andersen L. 2012. Two intercellular signals required for fruiting body formation in Myxococcus xanthus act sequentially but nonhierarchically. Mol. Microbiol. 86:65– 81. http://dx.doi.org/10.1111/j.1365-2958.2012.08173.x. 19. Lobedanz S, Sogaard-Andersen L. 2003. Identification of the C-signal, a contact-dependent morphogen coordinating multiple developmental responses in Myxococcus xanthus. Genes Dev. 17:2151–2161. http://dx.doi .org/10.1101/gad.274203. 20. Shimkets LJ, Rafiee H. 1990. CsgA, an extracellular protein essential for Myxococcus xanthus development. J. Bacteriol. 172:5299 –5306. 21. Kim SK, Kaiser D. 1990. Cell motility is required for the transmission of C-factor, an intercellular signal that coordinates fruiting body morphogenesis of Myxococcus xanthus. Genes Dev. 4:896 –905. http://dx.doi.org /10.1101/gad.4.6.896. 22. Kim SK, Kaiser D. 1990. Cell alignment required in differentiation of Myxococcus xanthus. Science 249:926 –928. http://dx.doi.org/10.1126 /science.2118274. 23. Kroos L, Hartzell P, Stephens K, Kaiser D. 1988. A link between cell movement and gene expression argues that motility is required for cell-cell signaling during fruiting body development. Genes Dev. 2:1677–1685. http://dx.doi.org/10.1101/gad.2.12a.1677. 24. Kroos L, Kaiser D. 1987. Expression of many developmentally regulated genes in Myxococcus depends on a sequence of cell interactions. Genes Dev. 1:840 – 854. http://dx.doi.org/10.1101/gad.1.8.840. 25. Shimkets LJ, Gill RE, Kaiser D. 1983. Developmental cell interactions in Myxococcus xanthus and the spoC locus. Proc. Natl. Acad. Sci. U. S. A. 80:1406 –1410. http://dx.doi.org/10.1073/pnas.80.5.1406. 26. Sager B, Kaiser D. 1993. Spatial restriction of cellular differentiation. Genes Dev. 7:1645–1653. http://dx.doi.org/10.1101/gad.7.9.1645. 27. Kruse T, Lobedanz S, Berthelsen NM, Sogaard-Andersen L. 2001. C-signal: a cell surface-associated morphogen that induces and coordinates multicellular fruiting body morphogenesis and sporulation in Myxococcus xanthus. Mol. Microbiol. 40:156 –168. http://dx.doi.org/10 .1046/j.1365-2958.2001.02365.x. 28. Li S-F, Lee B, Shimkets LJ. 1992. csgA expression entrains Myxococcus xanthus development. Genes Dev. 6:401– 410. http://dx.doi.org/10.1101 /gad.6.3.401. 29. Kim SK, Kaiser D. 1991. C-factor has distinct aggregation and sporulation thresholds during Myxococcus development. J. Bacteriol. 173:1722– 1728. 30. Julien B, Kaiser AD, Garza A. 2000. Spatial control of cell differentiation in Myxococcus xanthus. Proc. Natl. Acad. Sci. U. S. A. 97:9098 –9103. http: //dx.doi.org/10.1073/pnas.97.16.9098. 31. Shimkets LJ, Kaiser D. 1982. Murein components rescue developmental sporulation of Myxococcus xanthus. J. Bacteriol. 152:462– 470. 32. Janssen GR, Dworkin M. 1985. Cell-cell interactions in developmental lysis of Myxococcus xanthus. Dev. Biol. 112:194 –202. http://dx.doi.org/10 .1016/0012-1606(85)90133-2. 33. Lee B, Holkenbrink C, Treuner-Lange A, Higgs PI. 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 –3068. http://dx.doi .org/10.1128/JB.06756-11. 34. Lee J, Son B, Viswanathan P, Luethy P, Kroos L. 2011. Combinatorial regulation of fmgD by MrpC2 and FruA during Myxococcus xanthus development. J. Bacteriol. 193:1681–1689. http://dx.doi.org/10.1128/JB .01541-10. 35. Mittal S, Kroos L. 2009. A combination of unusual transcription factors binds cooperatively to control Myxococcus xanthus developmental gene expression. Proc. Natl. Acad. Sci. U. S. A. 106:1965–1970. http://dx.doi .org/10.1073/pnas.0808516106. 36. Mittal S, Kroos L. 2009. Combinatorial regulation by a novel arrangement of FruA and MrpC2 transcription factors during Myxococcus xanthus development. J. Bacteriol. 191:2753–2763. http://dx.doi.org/10.1128 /JB.01818-08. 37. Son B, Liu Y, Kroos L. 2011. Combinatorial regulation by MrpC2 and FruA involves three sites in the fmgE promoter region during Myxococcus xanthus development. J. Bacteriol. 193:2756 –2766. http://dx.doi.org/10 .1128/JB.00205-11.



loss of positive autoregulation and feedback likely accounts for the decline of mrpC transcript observed later. Conclusion. M. xanthus delays commitment to sporulation until a critical period around 24 to 30 h poststarvation. Nutrient addition before or during this period causes a rapid decline of MrpC. The primary response is proteolysis of MrpC that may involve ATP-independent metalloprotease activity, followed by declines in mrpC and dev transcripts, and FruA protein. This response may allow cells to escape commitment to sporulation and resume growth if nutrients reappear in a changing environment. More work is needed to precisely define the signal(s), signal transduction pathway(s), and protease(s) involved in the response and to identify genes acting after the dev operon (and perhaps in parallel with it) that commit a cell to the spore fate if starvation persists.



57.

58.

59.

60. 61. 62. 63. 64. 65. 66. 67.

68. 69. 70. 71. 72.

73. 74. 75.

76.

lation 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 –7974. http://dx.doi.org/10.1073/pnas.0701569104. Licking E, Gorski L, Kaiser D. 2000. A common step for changing cell shape in fruiting body and starvation-independent sporulation of Myxococcus xanthus. J. Bacteriol. 182:3553–3558. http://dx.doi.org/10.1128/JB .182.12.3553-3558.2000. Muller FD, Treuner-Lange A, Heider J, Huntley SM, Higgs PI. 2010. Global transcriptome analysis of spore formation in Myxococcus xanthus reveals a locus necessary for cell differentiation. BMC Genomics 11:264. http://dx.doi.org/10.1186/1471-2164-11-264. Muller FD, Schink CW, Hoiczyk E, Cserti E, Higgs PI. 2012. Spore formation in Myxococcus xanthus is tied to cytoskeleton functions and polysaccharide spore coat deposition. Mol. Microbiol. 83:486 –505. http: //dx.doi.org/10.1111/j.1365-2958.2011.07944.x. Gill RE, Karlok M, Benton D. 1993. Myxococcus xanthus encodes an ATP-dependent protease which is required for developmental gene transcription and intercellular signaling. J. Bacteriol. 175:4538 – 4544. Tojo N, Inouye S, Komano T. 1993. The lonD gene is homologous to the lon gene encoding an ATP-dependent protease and is essential for the development of Myxococcus xanthus. J. Bacteriol. 175:4545– 4549. Gonzalez-Pastor JE. 2011. Cannibalism: a social behavior in sporulating Bacillus subtilis. FEMS Microbiol. Rev. 35:415– 424. http://dx.doi.org/10 .1111/j.1574-6976.2010.00253.x. Gonzalez-Pastor JE, Hobbs EC, Losick R. 2003. Cannibalism by sporulating bacteria. Science 301:510 –513. http://dx.doi.org/10.1126/science .1086462. Rosenbluh A, Rosenberg E. 1989. Autocide AMI rescues development in dsg mutants of Myxococcus xanthus. J. Bacteriol. 171:1513–1518. Rosenbluh A, Rosenberg E. 1990. Role of autocide AMI in development of Myxococcus xanthus. J. Bacteriol. 172:4307– 4314. Varon M, Cohen S, Rosenberg E. 1984. Autocides produced by Myxococcus xanthus. J. Bacteriol. 160:1146 –1150. Kuchina A, Espinar L, Garcia-Ojalvo J, Suel GM. 2011. Reversible and noisy progression toward a commitment point enables adaptable and reliable cellular decision-making. PLoS Comput. Biol. 7:e1002273. http://dx .doi.org/10.1371/journal.pcbi.1002273. Dubnau D, Losick R. 2006. Bistability in bacteria. Mol. Microbiol. 61: 564 –572. http://dx.doi.org/10.1111/j.1365-2958.2006.05249.x. Eldar A, Elowitz MB. 2010. Functional roles for noise in genetic circuits. Nature 467:167–173. http://dx.doi.org/10.1038/nature09326. Ray JC, Tabor JJ, Igoshin OA. 2011. Nontranscriptional regulatory processes shape transcriptional network dynamics. Nat. Rev. Microbiol. 9:817– 828. http://dx.doi.org/10.1038/nrmicro2667. Mangan S, Zaslaver A, Alon U. 2003. The coherent feed-forward loop serves as a sign-sensitive delay element in transcription networks. J. Mol. Biol. 334:197–204. http://dx.doi.org/10.1016/j.jmb.2003.09.049. Ueki T, Inouye S. 2005. Identification of a gene involved in polysaccharide export as a transcription target of FruA, an essential factor for Myxococcus xanthus development. J. Biol. Chem. 280:32279 –32284. http://dx .doi.org/10.1074/jbc.M507191200. Dworkin J, Losick R. 2005. Developmental commitment in a bacterium. Cell 121:401– 409. http://dx.doi.org/10.1016/j.cell.2005.02.032. Konovalova A, Sogaard-Andersen L, Kroos L. 2014. Regulated proteolysis in bacterial development. FEMS Microbiol. Rev. 38:493–522. http: //dx.doi.org/10.1111/1574-6976.12050. Rolbetzki A, Ammon M, Jakovljevic V, Konovalova A, SogaardAndersen L. 2008. Regulated secretion of a protease activates intercellular signaling during fruiting body formation in Myxococcus xanthus. Dev. Cell 15:627– 634. http://dx.doi.org/10.1016/j.devcel.2008.08.002. Goldman BS, Nierman WC, Kaiser D, Slater SC, Durkin AS, Eisen J, Ronning CM, Barbazuk WB, Blanchard M, Field C, Halling C, Hinkle G, Iartchuk O, Kim HS, Mackenzie C, Madupu R, Miller N, Shvartsbeyn A, Sullivan SA, Vaudin M, Wiegand R, Kaplan HB. 2006. Evolution of sensory complexity recorded in a myxobacterial genome. Proc. Natl. Acad. Sci. U. S. A. 103:15200 –15205. http://dx.doi.org/10.1073/pnas .0607335103.

jb.asm.org 2747


38. Ellehauge E, Norregaard-Madsen M, Sogaard-Andersen L. 1998. The FruA signal transduction protein provides a checkpoint for the temporal coordination of intercellular signals in Myxococcus xanthus development. Mol. Microbiol. 30:807– 817. http://dx.doi.org/10.1046/j.1365-2958.1998 .01113.x. 39. Ogawa M, Fujitani S, Mao X, Inouye S, Komano T. 1996. FruA, a putative transcription factor essential for the development of Myxococcus xanthus. Mol. Microbiol. 22:757–767. http://dx.doi.org/10.1046/j.1365 -2958.1996.d01-1725.x. 40. Sogaard-Andersen L, Slack F, Kimsey H, Kaiser D. 1996. Intercellular C-signaling in Myxococcus xanthus involves a branched signal transduction pathway. Genes Dev. 10:740 –754. http://dx.doi.org/10.1101/gad.10 .6.740. 41. Sun H, Shi W. 2001. Genetic studies of mrp, a locus essential for cellular aggregation and sporulation of Myxococcus xanthus. J. Bacteriol. 183: 4786 – 4795. http://dx.doi.org/10.1128/JB.183.16.4786-4795.2001. 42. Ueki T, 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– 8787. http: //dx.doi.org/10.1073/pnas.1533026100. 43. Higgs PI, Jagadeesan S, Mann P, Zusman DR. 2008. EspA, an orphan hybrid histidine protein kinase, regulates the timing of expression of key developmental proteins of Myxococcus xanthus. J. Bacteriol. 190:4416 – 4426. http://dx.doi.org/10.1128/JB.00265-08. 44. Kuner JM, Kaiser D. 1982. Fruiting body morphogenesis in submerged cultures of Myxococcus xanthus. J. Bacteriol. 151:458 – 461. 45. Bradford M. 1976. A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Anal. Biochem. 72:248 –254. http://dx.doi.org/10.1016/0003 -2697(76)90527-3. 46. Yoder-Himes D, Kroos L. 2006. Regulation of the Myxococcus xanthus C-signal-dependent ⍀4400 promoter by the essential developmental protein FruA. J. Bacteriol. 188:5167–5176. http://dx.doi.org/10.1128/JB .00318-06. 47. Schramm A, Lee B, Higgs PI. 2012. Intra- and inter-protein phosphorylation between two hybrid histidine kinases controls Myxococcus xanthus developmental progression. J. Biol. Chem. 287:25060 –25072. http://dx .doi.org/10.1074/jbc.M112.387241. 48. Overgaard M, Wegener-Feldbrugge S, Sogaard-Andersen L. 2006. The orphan response regulator DigR is required for synthesis of extracellular matrix fibrils in Myxococcus xanthus. J. Bacteriol. 188:4384 – 4394. http: //dx.doi.org/10.1128/JB.00189-06. 49. Nariya H, Inouye S. 2005. Identification of a protein Ser/Thr kinase cascade that regulates essential transcriptional activators in Myxococcus xanthus development. Mol. Microbiol. 58:367–379. http://dx.doi.org/10 .1111/j.1365-2958.2005.04826.x. 50. Nariya H, 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. 60:1205–1217. http://dx.doi.org/10.1111/j.1365-2958.2006.05178.x. 51. Holmes AB, Kalvala S, Whitworth DE. 2010. Spatial simulations of myxobacterial development. PLoS Comput. Biol. 6:e1000686. http://dx .doi.org/10.1371/journal.pcbi.1000686. 52. Kroos L, Kuspa A, Kaiser D. 1990. Defects in fruiting body development caused by Tn5 lac insertions in Myxococcus xanthus. J. Bacteriol. 172:484 – 487. 53. Viswanathan P, Murphy K, Julien B, Garza AG, Kroos L. 2007. Regulation of dev, an operon that includes genes essential for Myxococcus xanthus development and CRISPR-associated genes and repeats. J. Bacteriol. 189:3738 –3750. http://dx.doi.org/10.1128/JB.00187-07. 54. Boysen A, Ellehauge E, Julien B, Sogaard-Andersen L. 2002. The DevT protein stimulates synthesis of FruA, a signal transduction protein required for fruiting body morphogenesis in Myxococcus xanthus. J. Bacteriol. 184:1540 –1546. http://dx.doi.org/10.1128/JB.184.6.1540-1546.2002. 55. Thony-Meyer L, Kaiser D. 1993. devRS, an autoregulated and essential genetic locus for fruiting body development in Myxococcus xanthus. J. Bacteriol. 175:7450 –7462. 56. Viswanathan P, Ueki T, Inouye S, Kroos L. 2007. Combinatorial regu-

Transcription factor MrpC binds to promoter regions of hundreds of developmentally-regulated genes in Myxococcus xanthus.

The dev Operon Regulates the Timing of Sporulation during Myxococcus xanthus Development.

csgA expression entrains Myxococcus xanthus development.

Two-Component Signal Transduction Systems That Regulate the Temporal and Spatial Expression of Myxococcus xanthus Sporulation Genes.

Changes in cyclic AMP levels during development in Myxococcus xanthus.

Phenotypic profiling of ABC transporter coding genes in Myxococcus xanthus.

CsgA, an extracellular protein essential for Myxococcus xanthus development.

Two lipid signals guide fruiting body development of Myxococcus xanthus.

Role of autocide AMI in development of Myxococcus xanthus.

Effects of glucosamine on lysis, glycerol formation, and sporulation in Myxococcus xanthus.

Myxobacterial hemagglutinin: a development-specific lectin of Myxococcus xanthus.

Rifampin-resistant mutants of Myxococcus xanthus defective in development.

Release of a cell surface protein during development of Myxococcus xanthus.

Carbohydrate accumulation during myxospore formation in Myxococcus xanthus.

Production of an extracellular milk-clotting activity during development in Myxococcus xanthus.

Bacillaene and sporulation protect Bacillus subtilis from predation by Myxococcus xanthus.

Cyclic nucleotides, cyclic nucleotide phosphodiesterase, and development in Myxococcus xanthus.

Allopatric integrations selectively change host transcriptomes, leading to varied expression efficiencies of exotic genes in Myxococcus xanthus.

Chromosome replication in Myxococcus xanthus.

Transposon tagging of genes for cell-cell interactions in Myxococcus xanthus.

A versatile class of cell surface directional motors gives rise to gliding motility and sporulation in Myxococcus xanthus.

Defects in contact-stimulated gliding during aggregation by Myxococcus xanthus.

Chloramphenicol resistance in Myxococcus xanthus.

Coats from Myxococcus xanthus: characterization and synthesis during myxospore differentiation.