A novel mechanism for bacterial sporulation based on programmed peptidoglycan degradation

Reviewer #2 (Public review):

The authors initial goal was to demonstrate loss of PG during the slow sporulation process of Myxococcus xanthus, with examination of the PG degradation products in order to implicate possible enzymes involved. Upon finding a predominance of LTG products, they examined sporulation in strains lacking each of the 14 candidate LTGs encoded in the genome, leading to the identification of two sporulation-linked LTGs. An extensive characterization of the roles played by these LTGs. One LTG is responsible for the slow sporulation PG degradation, while another is required for the rapid sporulation process. Interestingly, the "slow" LTG seems to provide an important regulatory brake on the rapid enzyme. Single molecule fluorescent tracking of these enzymes was used to develop a model for their interaction with PG that mimics their observed activity. The rate of PG synthesis activity was also shown to impact the rate of PG degradation, suggesting potential interplay between the synthetic and degradative enzymes.

Strengths:

The genetic analysis to identify sporulation-linked LTGs and their effects on growth sporulation, and spore properties was well done and productive. The fluorescence microscopy to track LTG mobility, presumably tied to activity, produced a convincing argument about the mechanism of regulation of one LTG by another. The authors have responded well to most points of the previous review.

Weaknesses:

While the impact of LTGs on sporulation was clearly demonstrated, the PG analysis that resulted in the study of LTGs raised some important unanswered questions. The analyses suggest that the PG is degraded to quite small fragments, which would normally be lost during the purification of PG. The conclusions concerning the PG degradation during sporulation needs to be clarified, as described below. The authors suggest a "new mechanism of sporulation" when they have actually simply identified an important factor (PG degradation by LTGs) within a complex "process of sporulation". This needs to be reflected also in title of the paper.

Author response:

The following is the authors’ response to the original reviews.

Public Reviews:

Reviewer #1 (Public review):

Summary:

Ramirez Carbo et al. use the powerful M. xanthus spore morphogenesis model to address fundamental mechanisms in coordinated peptidoglycan remodeling and degradation. As peptidoglycan is an essential macromolecule and difficult to study in vivo, the authors use indirect but important methodology. The authors first identify two lytic transglycosylase (Ltg) enzymes necessary for spore morphogenesis using mutant phenotypic studies. They characterize these mutants for their role in coordinating spore morphogenesis induced either in fruiting bodies (starvation-dependent) or in liquid-rich media conditions (chemical-dependent). They conclude from these phenotypic and epistatic analyses that LtgA is necessary for morphogenesis during chemical-induced sporulation, and LtgB appears to be necessary to coordinate LtgA activity by interfering with LtgA function. Under starvation-induced sporulation, the absence of LtgB interferes with the building of fruiting bodies. LtgA does not appear to play a primary role in promoting aggregation into fruiting bodies, nor in degradation of peptidoglycan as assayed by loss of signal in anti-PG immunofluorescence. The authors demonstrate that the purified periplasmic domain of LtgA is highly active in degrading purified PG sacculi in vitro, while that of LtgB is highly reduced (relative to LtgA or lysozyme). The authors use photoactivated mCherry Lyt fusions and PALM to track the fusion protein mobility, which they state correlates with activity as immobilization results from PG binding. They demonstrate that in vegetative cells, a greater proportion of LtgA-PAmCh is more immobile (more active) than LtgB-PAmCh, but that directly after chemical-induction of sporulation, LtgB-PAmCh becomes more immobile (active). These analyses in the partner mutant backgrounds suggest that LtgA-PAmCh is more immobile (less active) in the absence of LtgB, but the reverse is not observed. Finally, the authors demonstrate that overexpression of LtgA in vegetative conditions leads to cell rounding, likely because of uncontrolled PG degradation, while overexpression of LtgB displays no phenotype.

Strengths:

This paper capitalizes on a novel spore morphogenesis mechanism to define proteins and mechanisms involved in peptidoglycan reorganization. The authors use the powerful PALM microscopy technique to assess Ltg activity in vivo by assaying for immobility as a proxy for PG binding. The authors elucidate a novel mechanism by which two Ltg's function together- with one (LtgB) seeming to regulate the activity of the other (the primary Ltg).

Despite some weaknesses, there is no question that this study provides important insight into mechanisms of peptidoglycan remodeling- a difficult but highly impactful area of study with implications for the development of novel therapeutics and the discovery of mechanisms of fundamental bacterial physiology.

Weaknesses:

In many places, the authors do not adequately justify interpretations of their assays, leading to some apparently unjustified conclusions. Many of these are minor and may just require citations to demonstrate that the interpretations are justified by previous studies (detailed in recommendations below), but two bigger concerns are as follows:

(1) It is not clear how the muropeptides listed in Figure 1 were assigned, and it is missing in the methods. In the sporulating conditions, the spectra look like combinations of multiple peaks, and the data, as stated, is not convincing to the non-specialist eye.

We thank the reviewer for raising this point. We've expanded the Methods section to give a fuller account of how muropeptides were identified. In particular, we now describe the chromatographic separation and the assignment process, which relies on comparison with published data, fragmentation patterns, retention times, and accurate mass values. We acknowledge that the chromatograms show several peaks; nevertheless, only those muropeptides that we could clearly identify by MS/MS analysis were annotated in Figure 1. This clarification is now made explicit in the figure legend in the revised version of the manuscript.

(2) The observation that the lytB mutant prevents appropriate aggregation into fruiting bodies does not allow the interpretation that the absence of LtgB prevents PG morphogenesis in the starvation-induced sporulation pathway, per se. It is more likely that in the LtgB mutant, the morphogenesis program is not even triggered. This is because signaling proteins and regulators (specifically, C-signal accumulation/activated FruA), which are dependent on increased cell-cell signaling in the fruiting body, do not accumulate appropriately in shallow aggregates. C-signal/FruA are necessary to trigger the sporulation program in FBs. BTW: A hypothesis to explain the indirect effect of ltgB absence on aggregation could be that UDP-precursors are not regulated appropriately (unregulated LtyA (LtgA [sic])??), so polysaccharides necessary for motility are not properly produced.

Along these lines, fruiting body formation does not equal sporulation, and even "darkened" fruiting bodies can be misleading, as some mutants form polysacchariderich fruiting bodies (that appear dark under certain light conditions in the stereomicroscope) but do not sporulate efficiently. The wording in the text suggests that the authors assume that sporulation levels are normal because fruiting bodies are produced (see specific comments for details).

We deeply appreciate this question. Seeking the answer, we repeated the fruiting body assay and found that both the ΔltgA and ΔltgB mutants formed dark aggregates that were comparable to wild-type fruiting bodies. However, these “fruiting body-like” aggregates did not contain sonication-resistant spores. Thus, regardless of the signals, either glycerol or starvation, sporulation requires both LtgA and LtgB. We have corrected the mistakes in the first submission.

(3) The authors repeatedly state that production of spore coat polysaccharides likely affects the PG IP staining (see below), but this is not well justified. A citation is needed if this has already been directly shown, or the language needs to be softened.

We agree with the reviewer. We have softened our language as “However, we cannot exclude the possibility that the polysaccharide spore coats (Voelz & Dworkin, 1962) hinder antibody access to PG.”

(4) Better justification for the immobility of Ltg proteins in vivo as an assay for activity may be required. If this is well known in the field, it should be explicitly stated. The authors address this better in the discussion - but still state it is a correlation.

We elaborated the justification, “Thus, when diffusive enzymes bind to PG, their mobility decreases (Lee et al., 2016; Zhang et al., 2023). For instance, DacB, another PG hydrolase, reduces its single-particle mobility in the conditions where its activity is activated (Zhang et al., 2023). By tracking single fluorescently-labeled enzyme particles, we can approximate their PG-binding in different physiological conditions and genetic backgrounds (Ramirez Carbo et al., 2024, Zhang et al., 2023, Ramírez Carbó & Nan, 2026).”

We further discussed the correlation in discussion, “The simultaneous occurrence of reduced LtgA mobility and PG degradation during glycerol-induced sporulation indicates that the molecular dynamics of LtgA accurately mirrors its enzymatic activity. Such correlation between decreased particle mobility and increased enzymatic activity applies to many other PG-related enzymes, including multiple PG polymerases in E. coli and the endopeptidase DacB in M. xanthus (Lee et al., 2016, Zhang et al., 2023, Yang et al., 2021).”

Reviewer #2 (Public review):

Summary:

The authors' initial goal was to demonstrate loss of PG during the slow sporulation process of Myxococcus xanthus, with examination of the PG degradation products in order to implicate possible enzymes involved. Upon finding a predominance of LGT products, they examined sporulation in strains lacking each of the 14 candidate LTGs encoded in the genome, leading to the identification of two sporulation-linked LTGs. An extensive characterization of the roles played by these LTGs. One LTG is responsible for the slow sporulation PG degradation, while another is required for the rapid sporulation process. Interestingly, the "slow" LTG seems to provide an important regulatory brake on the rapid enzyme. Single-molecule fluorescent tracking of these enzymes was used to develop a model for their interaction with PG that mimics their observed activity. The rate of PG synthesis activity was also shown to impact the rate of PG degradation, suggesting potential interplay between the synthetic and degradative enzymes.

Strengths:

The genetic analysis to identify sporulation-linked LTGs and their effects on growth, sporulation, and spore properties was well done and productive. The fluorescence microscopy to track LTG mobility, presumably tied to activity, produced a convincing argument about the mechanism of regulation of one LTG by another.

Weaknesses:

While the impact of LTGs on sporulation was clearly demonstrated, the PG analysis that resulted from the study of LTGs raised some important unanswered questions. The analyses suggest that the PG is degraded to quite small fragments, which would normally be lost during the purification of PG. How these small fragments were thus detected is unclear, and this suggests a more complex story concerning PG metabolism during sporulation. An anti-PG antibody is used to quantify PG in the spores, but it is not made clear what the specificity of this antibody is, and thus whether it would recognize the LTG -altered PG of the spore. The authors suggest a "new mechanism of sporulation" when they have actually simply identified an important factor (PG degradation by LTGs) within a complex "process of sporulation".

Recommendations for the authors:

Reviewer #1 (Recommendations for the authors):

Details on places in the text that could be improved:

(1) Line 77: more appropriate "homologs of the sporulation genes"

Corrected.

(2) Line 137-139. Where are the 14 KO mutant data? Only showing the ones with phenotypes? Needs a citation if published elsewhere.

The phenotypes of other mutants are shown in the new Fig. S1.

(3) Line 144 and later: why "ORF"? Are they not homologous to Ltg's??. Also, the orf designation is not in the figure, so it is hard to follow the data the authors are presenting.

Following the reviewer’s recommendation, we deleted “ORF” and presented the ORF designation in the figure legend.

(4) Figure 2B - add the y-axis legend to the figure (also S1B).

Added

(5) Line 156: What does "same below" mean?

Deleted “same below”.

(6) Line 161: emtA is not indicated on Figure 2D - can you reword or add to the legend??

Changed to “MltE, an LTG encoded by Escherichia coli emtA”

(7) Line 163: "opposite roles" could be a bit better defined... do you mean ltgA induces rounding and ltgB delays rounding???

We agree with the reviewer. “opposite” was replaced by “different”.

(8) Line 172: But did the ltgA mutant make the same number of viable spores (not just FBs)? Can't conclude "the slow sporulation pathway only requires ltgB" if ltgA mutant was not tested.

We agree with the reviewer. In the revised manuscript, we quantified the starvation-induced spores and found that both LtgA and LtgB are required for forming mature spores. The data are shown in Figure 3 and supplement figures.

(9) Line 202: might be appropriate to indicate the degree of homology (% identity over protein length).

Added.

(10) Line 178, 181, and thereafter: DIC microscopy.

Corrected.

(11) Line 182/183 and 190: What is the basis for the conclusion that "polysaccharides sustained unflattened structures"??

We changed the description to “likely due to the deposition of spore coat polysaccharides that sustained unflattened cell structures (Wartel et al., 2013; Holkenbrink et al., 2014).”

(12) Line 185/186: What is the basis for the conclusion that ltgB only contained a small amount of PG? If it is based on reduced intensity, it is not obvious from the images presented. Quantitative analysis would better support this statement.

We changed the description to “Sacculi from the ∆ltgB pseudospores still contained PG but showed lower fluorescence intensity”. We also updated the figure to show the typical fluorescence intensities.

(13) Figure legend 3 line 245/6: It is not totally clear what the difference is, in that the white arrows are pointing to in ltgA vs ltgB mutant. Is it the proportion of large spherical objects in ltgB? What is the significance of the puncta in A vs. B?

We updated the description in the text as “While the remaining PG sacculi of these pseudospores were largely spherical, they lost integrity during purification, with many sacculi displaying irregular shapes in the fluorescence channel (Figure 4A).” In the figure legend, we changed the description to White arrows point to the sacculi in irregular shapes.”

(14) Line 190-192. Again, not really seeing what the authors are identifying to conclude "irregular shapes and ruptures" could authors pin-point more specifically and quantify such structures?

We updated the description in the text as “While the remaining PG sacculi of these pseudospores were largely spherical, they lost integrity during purification, with many sacculi displaying irregular shapes in the fluorescence channel (Figure 4A).”

(15) Line 194: if the authors want to make such a strong conclusion, they really should demonstrate this. Anti-spore coat antibodies are available in the field. Or take out these statements.

We took this statement out.

(16) Line 210: either they lack PG, OR the antibody can't gain access? How to conclude both? Might be more accurate to say "although we can't rule out that the polysaccharide spore coat prevents access to PG"

Following the reviewer’s recommendation, we changed the description to “These fruiting body spores lacked PG-specific fluorescence (Fig. 3A), consistent with their markedly reduced PG content (Fig. 1). However, we cannot exclude the possibility that the polysaccharide spore coats (Voelz & Dworkin, 1962) hinder antibody access to PG.

(17) Line 217-19: This is often stated in the literature, but full glycerol-induced spore maturation requires much more than 2 hours (note the authors are using O/N glycerol induction in Figure 1). And the long starvation-induced sporulation process is likely due to the differential start of sporulation, because the cells don't all enter the aggregate at the same time.... so they are not triggered to induce sporulation at the same time.

We agree with the reviewer on the first statement and changed “two hours” to “four hours”. For the second statement, we do not completely agree. Much research showed that spore development in fruiting bodies is well synchronized, for example, in (Dworkin & Voelz, 1962), starvation-induced spores did not at 48 h.

(18) Line 222: What is the evidence that it is really variation in production from the van promoter? Could it also just be due to differences in the cell cycle in the population? The authors may be correct, but should be less definitive about those conclusions since they haven't measured LtgA levels directly.

We agree with the reviewer. We changed the description to, “Upon induction with 200 μM vanillate, cells exhibited heterogeneous morphology, likely resulting from variations in LtgA expression or differences in cell cycle stages within the population.” Following this suggestion, we also changed the description on murA overexpression, “Similar to the cells that overexpressed LtgA (Fig. 3B), this heterogeneity likely reflects variable murA induction or the unsynchronized growth stages within the population.”

(19) Line 224: Where is the over-expressed LtgA data in Figure 2A? Is it that the authors are referring to over-expressed murA data, and the point is that overexpression of this kind of gene can lead to veg cell rounding? If the latter, this should be specifically stated in the text.

We apologize for this mistake. The data were shown in Fig. 3B, rather than Fig. 2A, 3B.

(20) Line 224: Did the authors test that LtgB is stably overproduced in veg cells? It could be that LtgB is turned over while LtgA is not. From the purified protein blot in Figure 3C, it does look like LtgB contains a degradation product (which is perhaps inhibiting the LtgB activity).

(21) Line 229: AgmT? Do the authors mean Ltg?

Corrected.

(22) Line 238: Is "rate" the right word? Technically, kinetics haven't been measured. Could state "LtgB less active" or "less efficient"?

Changed.

(23) Line 255: "ltgB shows a slight increase in expression during slow sporulation". Do the authors mean over the entire dev time course or specifically during the sporulation phase (which will be different timing for DZ2 vs DK1622)?

We clarified the description as “Consistent with its role in PG degradation, in a microarray-based transcriptome analysis, ltgA transcription was found to increase about twofold during rapid sporulation (4 h) but remain unchanged during slow sporulation (96 h). in the closely related DK1622 strain (Muller et al., 2010). Conversely, ltgB expression gradually rises during slow sporulation, reaching 1.8 times the vegetative level at 96 h, while remaining stable during rapid sporulation (Muller et al., 2010, Munoz-Dorado et al., 2019).”

(24) Line 278: This needs to be corrected. The authors have shown that production of fruiting bodies is not affected by the fusions. (also in Fig. S1 legend). This is really not the same as sporulation efficiency.

We changed the description to “the PAmCherry tags did not affect the formation of either glycerol-induced spores or starvation-induced fruiting bodies”.

(25) Figure S1A: panel A: Is there a deg product partially cut off at the bottom of the gel? (or is this a non-specific cross-reactive band). What is the predicted molecular mass for both proteins with the PAmCherry fusion? What conditions were these lysates generated from: veg prior to glycerol induction? I understand there are probably no antibodies available to LtgA or B, but it is important to note that it is not possible to know if there is simultaneously wt LtgA or B produced (by cleavage and degradation of the mCh fusion). Panel B: Were the differences in l/w between wt and the fusion strains tested to see if there really were no significant differences? Please state in the text (it looks like the data variance is higher in the fusion strains relative to the wt at 1 hr).

The bands at the bottom of the gel are the running front that appear in both lanes. They are not mCherry because there estimated molecular weight is much lower than that of mCherry (26.3 kDa). To avoid confusion, we cut these bands from the figure. The expression of both fusion proteins was detected from vegetative cells, which was clarified in the legend. The predicted molecular weights of them were provided in the legend too.

(26) Figure S1C: The ability to make fb is not the same as the production of spores. The authors should test the number of viable spores produced under starvation conditions, if they want to state starvation-induced sporulation is not affected by the fusions.

We agree with the reviewer. We quantified starvation-induced sporulation in the revised manuscript.

(27) Line 399: Figure S2 looks at fb formation in the absence of vanillate, not sporulation.

We changed the description to “cells grown without vanillate progressed normally through glycerol-induced sporulation and starvation-induced fruiting body formation”.

We also quantified starvation-induced sporulation in the revised manuscript.

(28) Line 281: How many fold is the ltgA transcript reduced compared to the ltgB? (i.e., If it is 1.2 fold reduced that may not be as worth mentioning as if it was 5-10 fold reduced).

ltgA transcription in vegetative cells was detected in a microarray (Muller et al., 2010) but not reported in RNAseq (Munoz-Dorado et al., 2019), significantly different from that of ltgB. We pointed this out in the revised manuscript.

(29) Figure 4B legend line 358. Define D (should it be italicized?).

Corrected.

(30) Line 363: Please define how significance was calculated. Is this the p-value?

We deleted the word “significant”.

(31) Line 298: How do the authors know immobility is from binding to PG? Provide a reference if this is well-known.

The rationale and references have been mentioned at the beginning of this section.

(32) Line 302 and thereafter: suggest "2.62 × 10-2 (plus minus) 2.0 × 10-3 μm2/s" is presented as "2.62 (plus minus) 0.20 × 10-2 μm2/s" for easier reading; switch to past tense (Were not are).

Changed following the reviewer’s recommendation.

(33) Line 310: "suggest" not "indicate", because binding of PG was directly tested.

Corrected.

(34) Line 341: "confirming it restricts access" seems very strong wording. Suggest: may compete with.

Changed.

(35) Line 392: SOME fb are larger- many are significantly smaller.

Because fruiting body sizes do not reflect sporulation efficiency, we removed this description.

(36) Figure S2 legend. Leaky expression; or on fruiting body formation.

Corrected.

(37) Line 436: stationary phase cells decrease PG synthesis- does this increase PG degradation? Perhaps it does lead to the death phase, which is striking in M. xanthus....

How do cells die, either through death phase or under antibiotic stresses, is not well understood (Baquero & Levin, 2021) Very likely, cell death is due to the accumulation of oxidative damages (Kohanski et al., 2007) and cell lysis could be a byproduct of cell death, when cells lose control of the enzymes that break PG. While cell death is a great topic to investigate, it is beyond the scope of this study.

(38) Line 507: washed.

Corrected.

(39) Line 524 (514 [sic]) and thereafter: sacculi.

Corrected.

(40) Line 587: reference for cell lysis procedure?

The procedures of cell lysis, column loading and elusion were described in details “…cells were harvested by centrifugation at 6,000 × g for 20 min and lysed by sonication in buffer A (20 mM Tris-HCl pH 8.0, 200 mM NaCl), (Nan et al., 2010, Nan et al., 2006). Proteins were loaded to an NGC™ Chromatography System (BIO-RAD) and 5-ml HisTrap™ columns (Cytiva) and eluted by buffer B (20 mM Tris-HCl pH 8.0, 200 mM NaCl, 500 mM immidazole) (Pogue et al., 2018, Nan et al., 2010).”

(41) Line 269: by microscopy (or "under THE microscope").

Corrected.

(42) Line 271: THE cell/PG.

“The” added.

(43) Line 603: OR not and.

Corrected.

(44) Line 497 (and elsewhere): Is it really CFU? If determined by OD, then not technically CFU because some cells will not grow into colonies.

We agree with the reviewer. Cell concentrations were determined by OD. “CFU” was deleted.

(45) Line 506: washed.

Same as recommendation (38). Corrected.

(46) Line 521: min.

Corrected.

(47) Line 532: How were peaks assigned?

We described peak assignment in details in the revised manuscript, “Muropeptides were assigned based on: (i) accurate mass matching to theoretical monoisotopic masses of expected M. xanthus PG building blocks (Bui et al., 2009, White et al., 1968) and (ii) comparison of retention times with those reported in previous analysis with similar PG compositions.”

Reviewer #2 (Recommendations for the authors):

(1) If almost all the muropeptides detected in spores are anhydro products of LTGs, then it might be expected that these are all very small peptidoglycan fragments in the spores. If the anhydro units were at the ends of short PG chains, then muramidase digestion would release similar amounts of non-anhydro products, but none are detected. So, is muramidase digestion doing anything to the PG derived from the spores? Is muramidase digestion required to observe the spore muropeptide pattern? A control sample in which muramidase digestion is omitted would answer these questions.

Muramidase digestion is essential for solubilizing PG into different muropeptides for UPLC analysis. In each sample, both anhydro and non-anhydro products were detected. In our original submission, we pointed out that “The two spore types showed similar profiles of a discernible presence of muropeptides that resembled those found in vegetative cells, albeit in significantly reduced quantities (Fig. 1).” We clarified the PG analysis. “The purification procedure yields only sedimentable PG, as all soluble fragments are removed during the washing steps. The resulting sacculi were then digested with muramidase, and the solubilized muropeptides were analyzed by UPLC (see Materials and Methods). The chromatograms in Fig. 1 reflect the muropeptides released specifically from the sedimented sacculus fraction.” Because muramidase release polysaccharides or disaccharides, so it’s digestion does not release nonanhydro GlcNAc species, which is why we did not see equal amounts of anhydro and non-anhydro products. In our case, over 90% of the polysaccharides and disaccharides contain Anhydro-MurNAc. The dominance of anhydro products indicates that LTGs cut very frequently on glycan chains.

(2) This also raises the question of how these very small peptidoglycan fragments are even retained in the spores. They would be expected to be lost during spore purification or during PG purification prior to muramidase digestion. How do you even purify sacculi when the spores have no PG chains? One could theorize that the polysaccharide coats hold everything in, but then the PG would be protected from muramidase digestion.

We thank the reviewer for this question. Our analysis suggests that M. xanthus spores do not lack PG entirely but retain a residual PG mesh that is still crosslinked. Such crosslinked material sediments during PG purification and remains accessible to muramidase, which hydrolyses internal glycosidic bonds and releases anhydro muropeptides. Non-crosslinked fragments would indeed be washed away during purification steps, so the detected muropeptides reflect the structure of this residual sacculus (Fig. 1).

(3) What is the anti-PG antibody recognizing, the glycan backbone, the peptide side chain, or both? Can the antibody recognize the very short anhydro-containing disaccharides proposed to be predominant in spores? If not, then the PG quantification using the antibody is not accurate.

The structures recognized by the anti-PG antibodies are unknown. Our samples do not contain small degradation products, which was clarified in the revised manuscript, “To answer this question, we purified cell sacculi and used immunofluorescence and an anti-PG serum (de Pedro et al., 1997) to visualize the remaining PG.” We used immunofluorescence to display the PG scaffolds remained in each sample and we did not perform any quantitative analysis based on the images.

(4) Lines 325-327: This is a speculative conclusion and should be stated as such, i.e., "may control the pace.." This conclusion could be somewhat more strongly stated at the end of the next section, around lines 345-350.

Moved following the reviewer’s recommendation.

(5) Lines 403-404: This first sentence of the discussion seems completely dissociated from the topic of the paper; it should be deleted.

Deleted.

(6) Lines 404-405. I am not convinced that these findings "elucidate a new mechanism of sporulation." The study was undertaken because PG degradation was already tied to sporulation in a previous study. Furthermore, I am not sure that PG degradation is a "mechanism of sporulation." It is clearly an important step in sporulation of this species, but is it the driving "mechanism"?

We tuned down our statement as “Our findings demonstrate that M. xanthus, a nonfirmicute bacterium, relies on PG degradation to change cell shape during sporulation”.

(7) Lines 510-516 describe PG purification from vegetative cells. How was this process modified for spores?

We apologize for the confusion. The process was clarified as “For PG analysis, samples were processed as previously described for Gram-negative bacteria (Alvarez et al., 2016; Desmarais et al., 2013). Vegetative cells were harvested at mid-stationary phase by centrifugation (30 min, 8,000 g). Vegetative cells and purified spores (as described in the previous section) were resuspended…”.

Alvarez, L., Hernandez, S.B., de Pedro, M.A., and Cava, F. (2016) Ultra-Sensitive, High-Resolution Liquid Chromatography Methods for the High-Throughput Quantitative Analysis of Bacterial Cell Wall Chemistry and Structure. Methods Mol Biol 1440: 1127.

Baquero, F., and Levin, B.R. (2021) Proximate and ultimate causes of the bactericidal action of antibiotics. Nat Rev Microbiol 19: 123-132.

Bui, N.K., Gray, J., Schwarz, H., Schumann, P., Blanot, D., and Vollmer, W. (2009) The peptidoglycan sacculus of Myxococcus xanthus has unusual structural features and is degraded during glycerol-induced myxospore development. J Bacteriol 191: 494505.

de Pedro, M.A., Quintela, J.C., Holtje, J.V., and Schwarz, H. (1997) Murein segregation in Escherichia coli. J Bacteriol 179: 2823-2834.

Desmarais, S.M., De Pedro, M.A., Cava, F., and Huang, K.C. (2013) Peptidoglycan at its peaks: how chromatographic analyses can reveal bacterial cell wall structure and assembly. Mol Microbiol 89: 1-13.

Dworkin, M., and Voelz, H. (1962) The formation and germination of microcysts in Myxococcus xanthus. J Gen Microbiol 28: 81-85.

Holkenbrink, C., Hoiczyk, E., Kahnt, J., and Higgs, P.I. (2014) Synthesis and assembly of a novel glycan layer in Myxococcus xanthus spores. J Biol Chem 289: 32364-32378.

Kohanski, M.A., Dwyer, D.J., Hayete, B., Lawrence, C.A., and Collins, J.J. (2007) A common mechanism of cellular death induced by bactericidal antibiotics. Cell 130: 797-810.

Lee, T.K., Meng, K., Shi, H., and Huang, K.C. (2016) Single-molecule imaging reveals modulation of cell wall synthesis dynamics in live bacterial cells. Nature communications 7: 13170.

Muller, F.D., Treuner-Lange, A., Heider, J., Huntley, S.M., and Higgs, P.I. (2010) Global transcriptome analysis of spore formation in Myxococcus xanthus reveals a locus necessary for cell diberentiation. BMC Genomics 11: 264.

Munoz-Dorado, J., Moraleda-Munoz, A., Marcos-Torres, F.J., Contreras-Moreno, F.J., MartinCuadrado, A.B., Schrader, J.M., Higgs, P.I., and Perez, J. (2019) Transcriptome dynamics of the Myxococcus xanthus multicellular developmental program. Elife 8.

Nan, B., Liu, X., Zhou, Y., Liu, J., Zhang, L., Wen, J., Zhang, X., Su, X.D., and Wang, Y.P. (2010) From signal perception to signal transduction: ligand-induced dimeric switch of DctB sensory domain in solution. Mol Microbiol 75: 1484-1494.

Nan, B., Zhou, Y., Liang, Y.H., Wen, J., Ma, Q., Zhang, S., Wang, Y., and Su, X.D. (2006) Purification and preliminary X-ray crystallographic analysis of the ligand-binding domain of Sinorhizobium meliloti DctB. Biochim Biophys Acta 1764: 839-841.

Pogue, C.B., Zhou, T., and Nan, B. (2018) PlpA, a PilZ-like protein, regulates directed motility of the bacterium Myxococcus xanthus. Mol Microbiol 107: 214-228.

Ramirez Carbo, C.A., Faromiki, O.G., and Nan, B. (2024) A lytic transglycosylase connects bacterial focal adhesion complexes to the peptidoglycan cell wall. Elife 13.

Ramírez Carbó, C.A., and Nan, B. (2026) Using Single-Particle Fluorescence Microscopy to Quantify Substrate Binding of Peptidoglycan-Modification Enzymes. Bio-protocol 16: e5696.

Voelz, H., and Dworkin, M. (1962) Fine structure of Myxococcus xanthus during morphogenesis. J Bacteriol 84: 943-952.

Wartel, M., Ducret, A., Thutupalli, S., Czerwinski, F., Le Gall, A.V., Mauriello, E.M., Bergam, P., Brun, Y.V., Shaevitz, J., and Mignot, T. (2013) A versatile class of cell surface directional motors gives rise to gliding motility and sporulation in Myxococcus xanthus. PLoS Biol 11: e1001728.

White, D., Dworkin, M., and Tipper, D.J. (1968) Peptidoglycan of Myxococcus xanthus: structure and relation to morphogenesis. J Bacteriol 95: 2186-2197.

Yang, X., McQuillen, R., Lyu, Z., Phillips-Mason, P., De La Cruz, A., McCausland, J.W., Liang, H., DeMeester, K.E., Santiago, C.C., Grimes, C.L., de Boer, P., and Xiao, J. (2021) A two-track model for the spatiotemporal coordination of bacterial septal cell wall synthesis revealed by single-molecule imaging of FtsW. Nat Microbiol 6: 584-593.

Zhang, H., Venkatesan, S., Ng, E., and Nan, B. (2023) Coordinated peptidoglycan synthases and hydrolases stabilize the bacterial cell wall. Nature communications 14: 5357.