# Effect of tcdR Mutation on Sporulation in the Epidemic Clostridium difficile Strain R20291

Brintha Girinathan, Marc Monot, Daniel Boyle, Kathleen N. Mcallister, Joseph A. Sorg, Bruno Dupuy, Revathi Govind

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Brintha Girinathan, Marc Monot, Daniel Boyle, Kathleen N. Mcallister, Joseph A. Sorg, et al.. Effect of tcdR Mutation on Sporulation in the Epidemic Clostridium difficile Strain R20291. MSphere, 2017, 2 (1), pp.00383-16. 10.1128/mSphere.00383-16. pasteur-01613368

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# Effect of *tcdR* Mutation on Sporulation in the Epidemic *Clostridium difficile* Strain R20291

Brintha P. Girinathan,ª Marc Monot,ᵇ Daniel Boyle,ª Kathleen N. McAllister,ᶜ

Joseph A. Sorg,ᶜ Bruno Dupuy,ᵇ Revathi Govindª

Division of Biology, Kansas State University, Manhattan, Kansas, USAª; Department of Microbiology, Laboratoire Pathogenèse des Bactéries Anaérobie, Institut Pasteur, Paris, Franceᵇ; Department of Biology, Texas A&M University, College Station, Texas, USAᶜ

**ABSTRACT** *Clostridium difficile* is an important nosocomial pathogen and the leading cause of hospital-acquired diarrhea. Antibiotic use is the primary risk factor for the development of *C. difficile*-associated disease because it disrupts normally protective gut flora and enables *C. difficile* to colonize the colon. *C. difficile* damages host tissue by secreting toxins and disseminates by forming spores. The toxin-encoding genes, *tcdA* and *tcdB*, are part of a pathogenicity locus, which also includes the *tcdR* gene that codes for TcdR, an alternate sigma factor that initiates transcription of *tcdA* and *tcdB* genes. We created a *tcdR* mutant in epidemic-type *C. difficile* strain R20291 in an attempt to identify the global role of *tcdR*. A site-directed mutation in *tcdR* affected both toxin production and sporulation in *C. difficile* R20291. Spores of the *tcdR* mutant were more heat sensitive than the wild type (WT). Nearly 3-fold more taurocholate was needed to germinate spores from the *tcdR* mutant than to germinate the spores prepared from the WT strain. Transmission electron microscopic analysis of the spores also revealed a weakly assembled exosporium on the *tcdR* mutant spores. Accordingly, comparative transcriptome analysis showed many differentially expressed sporulation genes in the *tcdR* mutant compared to the WT strain. These data suggest that regulatory networks of toxin production and sporulation in *C. difficile* strain R20291 are linked with each other.

**IMPORTANCE** *C. difficile* infects thousands of hospitalized patients every year, causing significant morbidity and mortality. *C. difficile* spores play a pivotal role in the transmission of the pathogen in the hospital environment. During infection, the spores germinate, and the vegetative bacterial cells produce toxins that damage host tissue. Thus, sporulation and toxin production are two important traits of *C. difficile*. In this study, we showed that a mutation in *tcdR*, the toxin gene regulator, affects both toxin production and sporulation in epidemic-type *C. difficile* strain R20291.

**KEYWORDS** *Clostridium difficile*, sporulation, toxin gene regulation

*Clostridium difficile* is a Gram-positive, spore-forming, anaerobic bacillus and is the leading cause of hospital-acquired diarrheal diseases (1, 2). Nearly 50% of all patients carry *C. difficile* asymptomatically after hospitalization (2, 3). Nearly 10% of all *C. difficile*-infected patients develop pseudomembranous colitis, and 3% develop severe, life-threatening complications such as fulminant colitis and toxic megacolon (4). *C. difficile* infection (CDI) is commonly acquired from *C. difficile* spores present in the hospital environment, and individuals become infected when the normal colonic microbiota is suppressed by antibiotic therapy (5). In the gut, *C. difficile* spores germinate to the toxin-producing vegetative form in response to certain bile acids, e.g., taurocholic acid (TA), and amino acids. *C. difficile* toxins A (TcdA) and B (TcdB) are then

Received 22 December 2016 Accepted 29 January 2017 Published 15 February 2017

**Citation** Girinathan BP, Monot M, Boyle D, McAllister KN, Sorg JA, Dupuy B, Govind R. 2017. Effect of *tcdR* mutation on sporulation in the epidemic *Clostridium difficile* strain R20291. mSphere 2:e00383-16. https://doi.org/10.1128/mSphere.00383-16.

Editor Craig D. Ellermeier, University of Iowa

**Copyright** © 2017 Girinathan et al. This is an open-access article distributed under the terms of the Creative Commons Attribution 4.0 International license.

Address correspondence to Revathi Govind, rgovind@ksu.edu.
secreted from the vegetative cell and cause tissue damage, necrosis, and inflammation
and are the main reasons for this disease outcome (6).

In *C. difficile*, the toxin genes, *tcdA* and *tcdB*, are located within a 19-kb pathogenicity
locus (PaLoc) and the *tcdR* gene, located upstream of *tcdB*, is required for expression of
the toxin genes. TcdR is an alternate sigma factor that directs transcription by recruiting
RNA polymerase to the toxin gene promoters and its own promoter (7, 8). Previous
studies have shown that other proteins can regulate toxin gene expression in response
to different environmental stimuli by controlling the transcription of *tcdR*. The sigma
factor SigD positively regulates toxin production by controlling the transcription of *tcdR*
(9). CodY, a global transcriptional regulator, represses the toxin gene expression by
binding with high affinity to the *tcdR* promoter region (10, 11). Finally, in response to
sugar availability, CcpA, a major regulator of carbon catabolite repression, binds to the
promoter region or the 5' ends of several PaLoc genes, with the strongest affinity to the
promoter region of *tcdR* (12, 13).

TcdR was the first member of the group V family of alternative sigma factors to be
described (14). We recently determined that TcsR, a toxin gene regulator in *Clostridium
sordellii*, is also a member of this family of sigma factors (15). Most of these alternative
sigma factors are autoregulated (7, 16) and are induced by environmental stresses, such
as nutritional limitation, DNA damage, or nonoptimal temperatures (8, 14, 17), sug-
gesting that these sigma factors function under these suboptimal growth conditions.

In this study, we created and characterized a mutation in *tcdR* in the epidemic-type
*C. difficile* R20291 strain to determine whether TcdR influenced cellular processes other
than toxin production. We found that the *tcdR* mutant sporulated less efficiently than
the wild-type (WT) strain. Moreover, spores prepared from the *tcdR* mutant were more
heat sensitive and had lower germination efficiency than the wild-type parental strain.
Electron microscopic (EM) analysis of the *tcdR* mutant spores also revealed a weakly
assembled exosporium. In agreement with these findings, comparative transcriptome
sequencing (RNA-seq) analyses of the WT and the *tcdR* mutant strains revealed several
sporulation genes to be affected by the *tcdR* mutation. These results suggested that a
mutation in *tcdR* not only affects toxin production but also influences the sporulation
pathway in the *C. difficile* R20291 strain. Interestingly, however, mutating *tcdR* in the
*C. difficile* 630Δ*erm* strain did not result in this phenotype, suggesting that the TcdR
regulon may be strain specific.

RESULTS

Mutation in *tcdR* affects both toxin production and sporulation in *C. difficile*
strain R20291. To analyze the global role of *tcdR* in *C. difficile* strain R20291, we used
a Clostron system (18) to inactivate the *tcdR* gene. Insertion of the group II intron into
the target gene (see Fig. S1A in the supplemental material) was verified by PCR using
intron-specific primers and *tcdR* gene-specific primers (Fig. S1B and Table S1 in the
supplemental material). Southern blotting confirmed the single chromosomal insertion
of the intron in the *tcdR* gene (Fig. S1C). Growth kinetics analyses were performed and
indicated that the inactivation of the *tcdR* gene did not affect the normal growth of the
bacterium (Fig. 1A). A toxin enzyme-linked immunosorbent assay (ELISA) was per-
formed with the cytosolic protein extracts of the *tcdR* mutant and the WT strain. We
observed a dramatic reduction in toxin production (Fig. 1B) in the mutant compared to
the WT, supporting the concept of the previously known function of TcdR as a positive
regulator of the toxin genes (7, 8, 16). Further, we measured the sporulation efficiency
of the *tcdR* mutant at the 24-h time point. A nearly 3-fold reduction in the level of
ethanol-resistant spores was observed in the *tcdR* mutant compared to the WT strain
(Fig. 2A). A similarly reduced sporulation rate (~2.6-fold) was observed when the
number of sporulation cells in the population was counted microscopically (Fig. 2B). We
then complemented the *tcdR* mutant by cloning and expressing *tcdR* from its own
promoter. Toxin production in the complemented strain was fully recovered (Fig. 1B),
whereas the effect on sporulation could be restored only partially (Fig. 2). Unlike toxin
gene regulation (where TcdR directly regulates *tcdA* and *tcdB* transcription), sporulation
<figure><img src="image_3.png" /></figure>

<figure><img src="image_4.png" /><figcaption>**FIG 1** Effect of *tcdR* inactivation on bacterial growth kinetics and toxin production. (A) Growth curve of R20291 and R20291::*tcdR* in TY medium. (B) TcdA and TcdB levels in cytosolic fractions after 10 h of growth. *C. difficile* strains were grown in TY medium, and toxins were quantified using ELISA. The data represent the averages of the results of three independent assays. Error bars in both panel A and B correspond to the standard errors of the means. The asterisks (**) in panel B indicate statistical difference at a *P* value of <0.005.</figcaption></figure>

is regulated by multiple transcription factors and alternative RNA polymerase sigma factors (19–21). Sporulation also involves finely tuned spatially and temporally regulated gene expression programs and may not be mimicked exactly in the complemented strain. All of these regulatory mechanisms could result in partial complementation of the sporulation. Another explanation could be that, when the TcdR sigma factor is overexpressed, the availability of RNA core polymerase for other sigma factors needed for sporulation could be limited and that limitation could result in partial complementation of the sporulation phenotype.

**Transcriptome analysis of tcdR mutant.** The global regulators *ccpA* and *codY* are known to influence both sporulation and toxin production in *C. difficile* (10, 11). We performed quantitative reverse transcription-PCR (qRT-PCR) analysis and found no significant change in their transcript levels in the *tcdR* mutant compared to the WT strain (Fig. S2). Since this initial analysis failed to explain the reasons behind the unexpected phenotype of the *tcdR* mutant, we decided to perform a transcriptome study using RNA-seq analysis. RNAs were prepared from stationary-phase cultures of the *tcdR* mutant (mutant R20291::*tcdR*) and the WT strain (strain R20291) and were subjected to RNA-seq analysis. The data observed for selected genes were confirmed by performing qRT-PCR analysis (Fig. S3 and S4). RNA-seq analysis of the *tcdR* mutant showed that most of the genes were underexpressed and revealed that two major classes of genes were particularly affected, i.e., the PaLoc genes and the sporulation-associated genes (see NCBI GEO accession number GSE85395). However, few genes were upregulated in the *tcdR* mutant. Among those that were overexpressed, we found the *srlR* gene encoding the regulator of glucitol/sorbitol-specific PTS system (CDR20291_0690 to CDR20291_0696). PaLoc genes (*tcdA*, *tcdB*, *tcdR*, and *tcdE*) were downregulated (33-fold, 12-fold, 5-fold, and 3-fold, respectively) in the *tcdR* mutant, as expected. Autoregulation of TcdR and its need for toxin gene transcription were well characterized previously (7, 8, 16). However, no report was available on the TcdR-mediated transcription of *tcdE* in the PaLoc. TcdE is a holin-like protein and was found to mediate toxin release from *C. difficile* cells (22, 23). Our data suggest that TcdR is also needed to initiate *tcdE* transcription in *C. difficile*.

**Many sporulation-associated genes were significantly repressed in the tcdR mutant.** In addition to the PaLoc genes, many genes in the sporulation pathways were repressed in the *tcdR* mutant compared to the WT (Table 1). Sporulation is a highly complex cellular process regulated by a cascade of events (20, 21, 24). Spo0A is the master regulator of sporulation, and its transcript levels were unchanged in the *tcdR*
<figure><img src="image_5.png" /></figure>

<figure><img src="image_6.png" /><figcaption>**FIG 2** Mutation in *tcdR* affects the sporulation efficiency in the R20291 strain. (A) Sporulation frequency (CFU per milliliter of ethanol-resistant spores) of R20291 plus pRPF185 (R20291+pRPF185), R20291::tcdR+pRPF185, and R20291::tcdR+pRGL294 (pRPF185 derivative plasmid carrying *tcdR*) strains grown for 24 h in 70:30 sporulation medium. The error bars correspond to standard errors of the means of results from 3 biological replicates. *, *P* < 0.05 (by two-tailed Student's *t* test). At least three independent experiments were performed. (B) Phase-contrast microscopy of paraformaldehyde-fixed R20291, R20291::tcdR+pRPF185, and R20291::tcdR+pRGL294 strains grown for 24 h in 70:30 sporulation plate. Percent sporulation (± standard deviation) was calculated (using the number of spores divided by the total number of spores and vegetative cells) from results from at least three independent experiments. Bar, 10 µm.</figcaption></figure>

mutant as observed in both the RNA-seq and qRT-PCR analyses (Table 1 and Fig. 3). However, we saw that transcripts of specific sporulation sigma factor genes *sigE*, *sigF*, *sigG*, and *sigK* were underexpressed in the *tcdR* mutant (Table 1 and Fig. 3). Even though the levels of transcription of these genes were moderately (1.5-fold to 2-fold) reduced in the *tcdR* mutant compared to the WT strain in the RNA-seq analysis, we observed through qRT-PCR analyses that their transcription levels in the *tcdR* mutant were significantly reduced throughout the time of growth (Fig. 3). RNA-seq analysis also revealed several sporulation genes controlled by *sigE*, *sigG*, and *sigK* to be significantly affected in the *tcdR* mutant (Table 1) (19–21). *SigE* is a mother cell-specific sigma factor responsible for the transcription of early sporulation-specific genes, and the *SigE*-regulated genes identified to be affected in *tcdR* mutant included the following: *spoIVA* (stage IV sporulation protein A); *spmBA* (spore maturation proteins B and A); and *sigK*, the second mother cell-specific sigma factor. *SigG* is the forespore-specific factor that controls the final stages of sporulation. The *SigG*-regulated genes found to be repressed in *tcdR* mutant included the following: *pdaA* (spore specific deacetylase), *sspA* (small acid-soluble protein), and *spoVAC* and *spoVAD* (stage V sporulation proteins). *SigG* and *SigE* activities were previously found to be required for the production of heat-resistant spores (21). The *sigK C. difficile* mutant was able to make heat-resistant spores; however, the level of production was 3 log lower than that seen with the parent
<table><caption>TABLE 1 Differentially expressed sporulation genes in R20291::tcdR mutant<sup>a</sup></caption><thead><tr><th>Gene ID</th><th>Gene name if assigned, known/predicted function</th><th>Fold downregulation in mutant (WT/tcdR mutant)</th><th>Known or predicted sigma factor needed for expression</th></tr></thead><tbody><tr><td>CDR20291_0124</td><td>Cell wall endopeptidase</td><td>3.844</td><td>SigF</td></tr><tr><td>CDR20291_2145</td><td>Hypothetical protein</td><td>5.993</td><td>SigF</td></tr><tr><td>CDR20291_2363</td><td>gpr, germination protease</td><td>4.008</td><td>SigF</td></tr><tr><td>CDR20291_3400</td><td>Spore cortex-lytic enzyme</td><td>5.652</td><td>SigF</td></tr><tr><td>CDR20291_3401</td><td>spolIR, stage II sporulation protein</td><td>4.228</td><td>SigF</td></tr><tr><td>CDR20291_2530</td><td>sigG</td><td>2.14</td><td>SigF</td></tr><tr><td>CDR20291_0125</td><td>spolIID, stage III sporulation protein D</td><td>5.323</td><td>SigE</td></tr><tr><td>CDR20291_0714</td><td>Stage IV sporulation protein</td><td>12.140</td><td>SigE</td></tr><tr><td>CDR20291_1031</td><td>spolIIAB, stage III sporulation protein AB</td><td>3.600</td><td>SigE</td></tr><tr><td>CDR20291_1032</td><td>spolIIAC, stage III sporulation protein AC</td><td>4.031</td><td>SigE</td></tr><tr><td>CDR20291_1033</td><td>spolIIAD, stage III sporulation protein AD</td><td>4.458</td><td>SigE</td></tr><tr><td>CDR20291_1034</td><td>spolIIAE, stage III sporulation-related protein</td><td>3.733</td><td>SigE</td></tr><tr><td>CDR20291_2147</td><td>cspBA, germination-specific protease</td><td>4.346</td><td>SigE</td></tr><tr><td>CDR20291_2513</td><td>spolVA, stage IV sporulation protein A</td><td>4.773</td><td>SigE</td></tr><tr><td>CDR20291_3376</td><td>spmB, spore maturation protein B</td><td>4.333</td><td>SigE</td></tr><tr><td>CDR20291_3377</td><td>spmA, spore maturation protein A</td><td>5.447</td><td>SigE</td></tr><tr><td>CDR20291_1073</td><td>Hypothetical protein</td><td>4.563</td><td>SigE</td></tr><tr><td>CDR20291_0702</td><td>spoVAC, stage V sporulation protein AC</td><td>5.524</td><td>SigG</td></tr><tr><td>CDR20291_0703</td><td>spoVAD, stage V sporulation protein AD</td><td>5.682</td><td>SigG</td></tr><tr><td>CDR20291_1130</td><td>Small acid-soluble spore protein</td><td>4.816</td><td>SigG</td></tr><tr><td>CDR20291_1131</td><td>dacF, D-alanyl-D-alanine-carboxypeptidase</td><td>5.891</td><td>SigG</td></tr><tr><td>CDR20291_1529</td><td>sodA, superoxide dismutase</td><td>5.714</td><td>SigG</td></tr><tr><td>CDR20291_2576</td><td>sspA, small acid-soluble spore protein A</td><td>4.500</td><td>SigG</td></tr><tr><td>CDR20291_2802</td><td>spoVFB, dipicolinate synthase subunit B</td><td>3.914</td><td>SigG</td></tr><tr><td>CDR20291_3080</td><td>Small acid-soluble spore protein</td><td>4.107</td><td>SigG</td></tr><tr><td>CDR20291_3107</td><td>sspB, small acid-soluble spore protein B</td><td>4.690</td><td>SigG</td></tr><tr><td>CDR20291_0212</td><td>Spore coat protein</td><td>6.600</td><td>SigK</td></tr><tr><td>CDR20291_0316</td><td>Spore coat assembly asparagine-rich protein</td><td>6.101</td><td>SigK</td></tr><tr><td>CDR20291_0337</td><td>Fragment of putative exosporium glycoprotein</td><td>12.666</td><td>SigK</td></tr><tr><td>CDR20291_0522</td><td>cotJB1, spore-coat protein</td><td>8.666</td><td>SigK</td></tr><tr><td>CDR20291_0523</td><td>cotJC1, spore-coat protein</td><td>6.842</td><td>SigK</td></tr><tr><td>CDR20291_2290</td><td>cotJB2, spore-coat protein</td><td>5.679</td><td>SigK</td></tr><tr><td>CDR20291_2291</td><td>cotJC2, spore-coat protein</td><td>5.165</td><td>SigK</td></tr><tr><td>CDR20291_2803</td><td>dpaA, dipicolinate synthase subunit A</td><td>4.291</td><td>SigK</td></tr><tr><td>CDR20291_3090</td><td>bclA2, exosporium glycoprotein</td><td>6.302</td><td>SigK</td></tr><tr><td>CDR20291_3193</td><td>bclA3, exosporium glycoprotein</td><td>12.612</td><td>SigK</td></tr><tr><td>CDR20291_3466</td><td>Cell wall hydrolase</td><td>4.631</td><td>SigK</td></tr><tr><td>CDR20291_0476</td><td>sleC, spore peptidoglycan hydrolase</td><td>5.502</td><td>Partly by SigF, SigK</td></tr><tr><td>CDR20291_2121</td><td>sinR</td><td>20.5</td><td>Unknown</td></tr><tr><td>CDR20291_2122</td><td>sinR like DNA binding protein</td><td>27.25</td><td>Unknown</td></tr><tr><td>CDR20291_0701</td><td>sigF*</td><td>1.23</td><td>SigH</td></tr><tr><td>CDR20291_2531</td><td>sigE*</td><td>1.56</td><td>SigH</td></tr><tr><td>CDR20291_1052</td><td>spoOA*</td><td>1.56</td><td>SigH</td></tr><tr><td>CDR20291_1067B</td><td>sigK*</td><td>1.78</td><td>SigE</td></tr></tbody></table>

strain (21). SigK regulates many genes encoding spore structure proteins that participate in the synthesis of the spore coat and spore exosporium. In fact, we found that many of the SigK-regulated genes such as cotJBD, cotA, cotB, cotE, bclA3, and bclA2 as well as the sleC and cdeC genes were significantly underexpressed in the tcdR mutant compared to the WT strain. The downregulation of these genes was confirmed by qRT-PCR analysis (Fig. S3 and S4).

**CDR20291_2121 and CDR20291_2122 (sin operon) were repressed in the tcdR mutant.** Other than genes involved in sporulation morphology, we also found some regulatory genes potentially involved in sporulation to be affected in the tcdR mutant. In fact, the transcript levels of CDR20291_2121 (coding for a SinR-like protein of *Bacillus subtilis*) and CDR20291_2122 (coding for a DNA binding protein) genes were nearly 20-fold lower in the tcdR mutant than in the WT strain (Table 1). This result was confirmed by qRT-PCR (Fig. 3). In *B. subtilis*, SinR is encoded within the *sin* locus carrying both *sinl* and *sinR* genes. In *B. subtilis*, SinR forms tetramers, which repress *spoOA*.

<footnote><sup>a</sup>Genes were considered differentially expressed if the fold change was ≥2.0 and their adjusted *P* value is ≤0.05. Expression levels of genes marked with (*) were not statistically significant. ID, identifier.</footnote>
<figure><img src="image_7.png" /><figcaption>**FIG 3** Decreased expression of key sporulation genes in the *tcdR* mutant. Data represent results of qRT-PCR analysis of *sigE*, *sigF*, *sigG*, *sigK*, *spo0A*, *sinR*, and *R20291_2122* expression after 16 and 24 h of *C. difficile* growth in 70:30 sporulation medium. Error bars correspond to the standard errors of the means of results from at least three biological replicates. *, *P* < 0.05 (by two-tailed Student's *t* test).</figcaption></figure>

transcription, although *Sinl* is an inhibitor of *SinR* (25). If *SinR* functions similarly in *C. difficile*, a decrease in *SinR* activity should lead to an increase of sporulation. However, we observed decreased sporulation in the *tcdR* mutant (Fig. 2), suggesting that the products of the *sin* locus must function differently in *C. difficile*.

**Spores derived from the tcdR mutant have increased heat sensitivity.** To compare the levels of heat sensitivity of spores between the WT and the *tcdR* mutant strains, we incubated purified spores at 70°C for 0.5 h, 4 h, and 8 h. When we monitored cell viability using the heat-treated spores, we found that spores from the *tcdR* mutant lost most of their viability upon 4 h of heat treatment and that they were nearly 10-fold more sensitive to heat than the WT spores (Fig. 4A). This could have been due to the decreased expression of both *sigG* and *sigE* in the *tcdR* mutant as observed in our transcriptional analysis; their activities are known to be involved in the formation of heat resistance of spores (21). In addition, the lower expression of many of the spore structure proteins (including *cdeC*) in the *tcdR* mutant can also explain the heat sensitivity of these spores.

**Increased taurocholate was required by tcdR mutant spores for germination.** To test if the lower transcription of sporulation-associated genes observed in the *tcdR* mutant (Table 1) affects the ability of *C. difficile* spores to germinate, we determined the apparent interaction of spores with taurocholic acid (Fig. 4B). *C. difficile* spores were

<figure><img src="image_8.png" /></figure>

<table><caption>**FIG 4** (A) The *tcdR* mutant affects spore germination. Heat resistance of spores of *C. difficile* strain R20291 and its *tcdR* mutant derivatives was measured by heat-treating aliquots at 70°C for 0.5 h, 4 h, and 8 h. The surviving spores were enumerated as described in Materials and Methods. The data represent the averages of the results of three independent experiments, and error bars represent standard errors of the means. Asterisks (*) indicate statistical difference at a *P* value of <0.05. (B) Apparent affinity of taurocholate for *C. difficile* spores. EC₅₀s were individually calculated from three independent germination experiments and are reported as averages with standard errors of the means. A Student's *t* test was performed, and that asterisk indicates that the calculated *P* value is <0.05.</caption><thead><tr><td>Strain</td><td>Apparent Affinity (EC50 in mM)</td></tr></thead><tbody><tr><td>R20291</td><td>2.1 ± 0.52</td></tr><tr><td>R20291:: tcdR</td><td>6.0 ± 1.2 *</td></tr></tbody></table>
<figure><img src="image_9.png" /><figcaption>**FIG 5** Transmission electron microscopic analysis of *C. difficile* spores. The images show thin sections of spores from the WT R20291 strain (A) and the R20291::tcdR mutant (B). Abbreviations: ex, exosporium; ct, coat; co, core; cx, cortex. Bar, 100 nm.</figcaption></figure>

suspended in rich medium alone or supplemented with increasing concentrations of the germinant taurocholate. The kinetics of spore germination were followed by measuring the rate of the decrease in optical density at 600 nm (OD₆₀₀) as the spores germinated (see Materials and Methods). Though not traditional enzyme kinetics, this assay allows us to understand how spores interact with the taurocholate germinant. *C. difficile* R20291 spores display a 50% effective concentration (EC₅₀) of 2.1 mM (similar to what has been previously reported for other strains) (26–28). However, the *tcdR* mutant spores display an EC₅₀ of 6.0 mM, corresponding to a 3-fold reduction (<0.05 *P* value) in TA affinity. These results support the overall observation that spore-associated functions were affected when *tcdR* was inactivated in strain R20291.

**Exosporium assembly was affected in the tcdR mutant.** Spores of the R20291::tcdR mutant were compared to those of the WT using electron microscopy to assess any effect on gross spore morphology. Samples were viewed as embedded thin sections, and the analysis revealed that *tcdR* mutant spores had a defect in their exosporium assembly (Fig. 5B and S5). The spore core of the *tcdR* mutant was stained weakly compared to the core of the WT spores, and darkly stained particulate materials were present over the spore coat and throughout these preparations. Weaker exosporium in the *tcdR* mutant spores could have made them susceptible to structural changes during chemical fixation procedures, resulting in these darker particles around the spores. In contrast, most of the R20291 WT spore had an intact exosporium that fully enclosed the spore coat (Fig. 5A) and was devoid of the darker debris observed in the *tcdR* mutant spores. This observation suggests that the *tcdR* mutation in R20291 affects the spore structure, with a profound effect on its exosporium assembly.

**The effect of tcdR on sporulation is strain specific.** Previous studies have shown that mutations in a specific gene can result in different phenotypes in different *C. difficile* strain backgrounds (29, 30). To understand whether the effect of TcdR is strain dependent, we created a *tcdR* mutant in the 630Δ*erm* strain using a ClosTron system. Toxin production in the 630Δ*erm*::*tcdR* mutant was severely downregulated as observed in strain R20291::*tcdR* (Fig. S6). But unlike the results seen with the R20291::*tcdR* strain, the sporulation efficiency of the 630Δ*erm*::*tcdR* strain was nearly 2-fold greater than that of its WT strain (Fig. 6). The similar opposing phenotype was previously reported for the *spo0A* mutants of strain R20291 versus the 630Δ*erm* mutant, which also affects the toxin production (29, 30). Though the *spo0A* mutation resulted in increased toxin production in the R20291 strain, it resulted in reduced toxin production in the 630Δ*erm* background. Even though the R20291 and 630 strains share 3,247 core genes, their genomes are significantly different from one another (31), whereas there are 47 coding sequences unique in R20291 compared to the 630 strain and 505 coding sequences unique in 630 compared to the R20291 strain (31). Therefore, the difference
<figure><img src="image_10.png" /><figcaption>**FIG 6** Effect of *tcdR* on sporulation is strain specific. (A) Sporulation frequency (CFU per milliliter of ethanol-resistant spores) of 630Δ*erm* and 630Δ*erm::tcdR* strains grown for 24 h in 70:30 sporulation medium. The error bars correspond to standard deviations of results from at least three biological replicates. The asterisk (*) indicates a *P* value of <0.05 (by two-tailed Student's *t* test). (B) Phase-contrast microscopy of paraformaldehyde-fixed 630Δ*erm* and 630Δ*erm::tcdR* strains grown for 48 h in a 70:30 sporulation plate. At least three independent experiments were performed to calculate percent sporulation (± standard errors of the means).</figcaption></figure>

that we observed in these two strains concerning the impact of the *tcdR* mutation on
sporulation might be related to the presence or absence of any of these unique genes.
Even though we do not know the exact reason for these differences, these observations
suggest that the *C. difficile* genome is dynamic and that its regulatory networks are fluid
in nature.

DISCUSSION

TcdR-mediated toxin gene regulation is well studied in *C. difficile* (7, 8, 16). The aim of this study was to understand whether TcdR could influence cellular processes other than toxin production. To investigate this issue, we created a *tcdR* mutant in the R20291 strain and performed several phenotypic assays. As expected, the *tcdR* mutant strain either produced no toxins or produced toxins at levels that were not detectable. Surprisingly, we also observed that the level of spores produced by the mutant was significantly reduced compared to the level seen with wild type.

The link between toxin production and sporulation in *C. difficile* has always been
suggested but has not been well studied. For example, in *C. difficile* R20291, a mutation
in *spo0A*, the master regulator of sporulation, resulted in changes in toxin production
(32). More recently, Edwards et al. reported that inactivation of CD3688 (*rstA*) in
*C. difficile* strain 630 affects sporulation, toxin production, and motility (33). Moreover,
it has been shown that the global regulators *CodY* and *CcpA* regulate toxin production
along with sporulation (10, 12, 34). Thus, if the *tcdR* mutation affects *codY*, *ccpA*, or
*spo0A* expression, both toxin production and sporulation could be influenced. When we
measured transcript levels of these genes by qRT-PCR, we found no change in their
levels in the *tcdR* mutant compared to the WT (see Fig. S2 in the supplemental
material). However, the genome-wide transcriptome analyses of the *tcdR* mutant
confirmed that many sporulation genes were affected.

Nearly 50% of the sporulation genes downregulated in the *tcdR* mutant are known (or predicted) to be under the control of SigE and SigK for their transcription (Table 1) (19–21, 35, 36). Among the downregulated SigE-dependent genes, we found *sigK*, whose presence could explain the transcriptional decrease in the levels of several SigK target genes in the *tcdR* mutant. The RNA-seq analyses of the *tcdR* mutant showed that transcription of *spoIIR* and *spoIIID* genes was reduced (Table 1). SpoIIR is essential for the activation of SigE (19, 35), and *spoIIID*, encoding a transcriptional regulator, is involved in the transcription of *sigK* (36). In *C. difficile*, as in *B. subtilis*, SigE is activated by proteolytic cleavage of the SigE precursor form (pro-SigE) (20). In *B. subtilis*, the enzyme SpoIIGA, which is responsible for pro-SigE processing, is coexpressed with *sigE* and is activated only when the mother cell and forespore compartments are formed (37–39). The trigger for SpoIIGA activation is the SpoIIR signal protein that is synthe-
sized in the newly formed forespore and whose presence is communicated to the
mother cell (40, 41). In *B. subtilis*, *spoII*R is regulated by SigF, whereas in *C. difficile*, partial
SigE processing is observed in *sigF* mutants, suggesting a lower level expression of
*spoII*R in the absence of SigF (19, 20). If the expression of *spoII*R in a *sigF* mutant is
influenced by TcdR, this could explain the partial processing of SigE in *sigF* mutants.
Thus, a reduced abundance of *spoII*R in the *tcdR* mutant could lead to low levels of
activated SpoIIGA and part of pro-SigE would remain unprocessed and inactive. If so,
this would also result in a decrease of *spoIIID* levels as observed in the transcriptome
(Table 1); therefore, little or no transcription of *sigK* would occur, resulting in poor spore
maturation.

Most of the genes identified as affected in the *tcdR* mutant code for proteins that are part of the spore proteome (20, 32, 42) and are involved in spore structure and germination. To determine whether *tcdR* mutant spore properties are different from those of the WT spores, we performed heat sensitivity and germination assays using purified spores. TcdR mutant spores were 10 times more heat-sensitive than WT spores (Fig. 4A). Accordingly, transcriptome analysis showed that several exosporium and coat protein coding genes were underexpressed in the *tcdR* mutant. A recent study on the *C. difficile* exosporium protein BclA3 demonstrated its role in spore heat resistance (43). The authors found that BclA3 is glycosylated by a glycosyltransferase encoded by the adjacent gene (CD3350) within the same operon whose mutation resulted in unglycosylated BclA3. They showed that spores from this mutant were highly susceptible to heat treatment compared to the WT spores (43). The same heat susceptibility was observed with the exosporium protein CdeC, which is present only in *C. difficile* and is needed for the assembly of exosporium (44). Also, *C. difficile* *spoVAC* and *dpaAB* mutants produced heat-sensitive spores (45). The dipicolinate synthase enzyme subunits (SpoVFB and DpaA) are responsible for the production of dipicolinic acid (DPA), which protects spores during heat treatment (46–48). Moreover, proteins encoded in the *spoVA* operon are responsible for transporting DPA from the mother cell to forespores during spore development (49).

All these results are consistent with the transcriptome analysis of the *tcdR* mutant,
which showed decreased expression of *bclA3*, *cdeC*, *spoVAC*, and the DPA synthase
coding operon. This probably results in the production of spores with weaker exospor-
ium that must be more sensitive to heat treatment than the WT strain (Fig. 4A).
Transmission electron microscopy (TEM) analysis of the *tcdR* mutant spores confirmed
this speculation, where the exosporium was found to be defective and weakly assem-
bled (Fig. 5).

Germination of bacterial spores is induced when the germinant receptors (GR) sense
germinants and subsequently trigger the release of spore core DPA (46). The release of
DPA from the spore core leads to the activation of cortex hydrolases that degrade the
peptidoglycan (PG) cortex layer, which then allows core hydration. In *C. difficile*, *CspC*
is the bile salt-sensing germinant receptor and is necessary for the release of DPA from
spores (26). *SleC* is the spore cortex lytic enzyme, and its activation depends on *CspC*
(through *CspB*-mediated cleavage of the prodomain to generate active *SleC*) (26,
50–52). A mutation in *sleC* was previously reported to affect germination in *C. difficile*
(51, 53). Thus, lower transcription of *sleC* in *tcdR* mutant (Table 1) suggested that *tcdR*
mutant spores could have inefficient germination. In agreement, we have shown that
the TA affinity of *C. difficile* *tcdR* spores is low compared to that of the WT spores
(Fig. 4B), indicating that germination is significantly reduced.

Several studies have previously identified the TTTACA sequence as the -35 region
of the TcdR-dependent promoters (7, 8). To test whether some of the downregulated
sporulation genes in the tcdR mutant can be directly controlled by TcdR, we looked for
the presence of this consensus sequence in the promoter regions of these genes
(Table 1). In fact, we found 9 genes/operons carrying the sequence in the -35 region
of the TcdR-dependent promoters. These genes include *bclA2*, *bclA3*, *cotJBD*, *spoVFB*,
*cotA*, *cotB*, *cotE*, *dpaA*, and *sin*. To test if any of these genes are directly controlled by
TcdR, we constructed transcriptional fusions between the promoter of the *bclA2* and
bclA3 genes and the *Escherichia coli* β-glucuronidase (*gusA*) gene that we introduced in a *gus*-negative *E. coli* strain expressing or not expressing TcdR as we did previously (54). Compared to the control strains, we did not see any TcdR-mediated transcription of bclA2 or bclA3 promoters, indicating that TcdR is not a direct regulator of these genes (Fig. S7). However, we cannot exclude for these genes the possibility that TcdR may act together with a specific regulator present in the R20291 strain.

Finally, the effect of TcdR on sporulation could be indirect. TcdR is an alternate sigma factor, and its presence or absence could influence the availability of the RNA polymerase core enzyme for other sigma factors in the cell, which in turn can influence the gene expression pattern. Thus, the absence of TcdR in the R20291::tcdR strain, increasing the availability of RNA polymerase core enzyme to other sigma factors, could indirectly affect those involved in the sporulation process. On the other hand, there may be common regulators that connect toxin gene regulation with the sporulation pathway in C. difficile that could be affected by the tcdR mutation. Previous studies have identified several regulators in C. difficile regulating toxin production along with sporulation, which strongly suggests that these two pathways were linked (10, 12, 33, 34, 55).

In the past decade, large C. difficile outbreaks, with higher relapse rates and increased mortality rates, were reported throughout the world and were attributed to C. difficile strains belonging to ribotype 027. Strain R20291 used in this study is a ribotype 027 isolate (56). Genetic and phenotypic features of this ribotype hint that the strains grouped as 027 ribotypes are different from other C. difficile strains (31). Recently, Lyon et al. reported that CdtR, a regulator in the binary toxin locus CdtLoc, could regulate toxin production only in 027 ribotypes and not in others (57). The authors of that study proposed that CdtR could be regulating toxin production by regulating the TcdR through a yet-to-be-identified intermediary regulator in the 027 ribotype. It has been previously proposed that the ability to regulate toxin production in response to various environmental cues with various regulatory responses may be different for 027 ribotypes in comparison to other C. difficile ribotypes (31). Results from subsequent studies are in agreement with this proposal. For example, a mutation in the highly conserved codY gene results in different phenotypes from 027 ribotypes and other ribotypes. The codY mutation results in a hypersporulation phenotype in a 027 ribotype (UK1 strain) and produces only a moderate effect on the sporulation in an 012 ribotype (630 strain) (34). It is also worth noting that sin locus expression levels were different in codY mutants in these two different C. difficile backgrounds (34). Similarly, a mutation in spo0A resulted in increased toxin production only in the 027 ribotype and not in the 012 ribotype (29, 30). In the current study, we observed the positive influence of TcdR on sporulation only in R20291 of the 027 ribotype and not in strain 630 of ribotype 012. Even though those previous studies, along with our observations, suggested that ribotype 027 has unique gene regulatory networks that differ from those of other C. difficile strains, variations may be present in strains within the 027 ribotype. Detailed study is needed to check whether the gene regulatory networks of the toxin synthesis and sporulation pathway are connected in all known ribotype 027 strains. In such a case, the ability to synchronize the toxin production and the sporulation can provide the selective advantage to ribotype 027 isolates to enable them to be more successful, with increased virulence and high transmission abilities. Deciphering the connections between toxins and the sporulation regulatory network could lead to the discovery of other novel regulators and pathways that can be targeted for the development of new therapeutics to manage C. difficile infections. Any treatment that leads to inhibition of toxin production and spore formation in patients with C. difficile infection can potentially lower the severity of the disease in addition to the transmission and recurrence of infection through dissemination of the spores.

## MATERIALS AND METHODS

**Bacterial strains and growth conditions.** *Clostridium difficile* strains (Table 2) were grown anaerobically in TY agar (tryptose, yeast extract) or 70:30 medium (58) as described previously (15, 54). Cefoxitin
<table><caption>TABLE 2 Bacterial strains and plasmids used in this study</caption><thead><tr><th>Strain or plasmid used</th><th>Description</th><th>Reference or source</th></tr></thead><tbody><tr><td><strong>Strains</strong></td><td></td><td></td></tr><tr><td>C. difficile R20291</td><td>NAP1/027 ribotype</td><td>31</td></tr><tr><td>C. difficile R20291::tcdR</td><td>R20291 with intron insertion in tcdR gene</td><td>This study</td></tr><tr><td>C. difficile 630Δerm</td><td>Erm' derivative of strain 630</td><td>63</td></tr><tr><td>C. difficile 630Δerm::tcdR</td><td>630Δerm with intron insertion in tcdR gene</td><td>This study</td></tr><tr><td>E. coli DH5α</td><td>endA1 recA1 deoR hsdR17 (rK<sup>-</sup> mK<sup>+</sup>)</td><td>NEB laboratories</td></tr><tr><td>E. coli S17-1</td><td>Strain with integrated RP4 conjugation transfer function for conjugation between E. coli and C. difficile</td><td>59</td></tr><tr><td>E. coli GM241(DE3)</td><td>gusA mutant lysogenized with DE3 phage and host for gusA reporter plasmids</td><td>54</td></tr><tr><td><strong>Plasmids</strong></td><td></td><td></td></tr><tr><td>pMTL007-CE5</td><td>ClosTron plasmid</td><td>18</td></tr><tr><td>pMTL007-CE5::tcdR-141</td><td>pMTL007-CE5 carrying tcdR-specific intron</td><td>This study</td></tr><tr><td>pRPF185</td><td>C. difficile shuttle vector</td><td>64</td></tr><tr><td>pRGL294</td><td>pRPF185 with tcdR expressed from its own promoter</td><td>This study</td></tr><tr><td>pACYC184</td><td>E. coli cloning vector; compatible with pET16B</td><td>Neb</td></tr><tr><td>pACYC515</td><td>pACYC184 vector carrying gusA gene under the control of the tcdR promoter</td><td>54</td></tr><tr><td>pET16b</td><td>E. coli expression vector</td><td>Novagen</td></tr><tr><td>pRGL312</td><td>pET16B with tcdR</td><td>This study</td></tr><tr><td>pRGL320</td><td>pACYC184 vector carrying gusA gene under the control of the bclA2 promoter</td><td>This study</td></tr><tr><td>pRGL321</td><td>pACYC184 vector carrying gusA gene under the control of the bclA3 promoter</td><td>This study</td></tr><tr><td>C. difficile R20291::tcdR + pRGL294</td><td>R20291::tcdR complemented with tcdR</td><td>This study</td></tr><tr><td>C. difficile R20291::tcdR + pRPF185</td><td>R20291::tcdR with vector control</td><td>This study</td></tr></tbody></table>

(Cef; 25 µg/ml), thiamphenicol (Thio; 15 µg/ml), and lincomycin (Lin; 15 µg/ml) were added to C. difficile cultures whenever necessary. *Escherichia coli* strains were grown in (LB) broth. E. coli strain S17-1 (S9), used for conjugation, was supplemented with ampicillin (100 µg/ml) or chloramphenicol (25 µg/ml) when indicated and cultured aerobically in LB broth.

**Construction of a tcdR mutant.** A tcdR mutation was constructed in a C. difficile strain using a ClosTron gene knockout system (18). The group II intron insertion site in the antisense orientation between nucleotides 141 and 142 of the tcdR ORF was selected using the Perutka algorithm, a Web-based design tool available at http://www.clostron.com. The designed retargeted intron was cloned into pMTL007-CE5, and the resulting plasmid, pMTL007-CE5::Cdi-tcdR-141a, was transferred into R20291 by conjugation as described previously (15, 22). The selection of thiamphenicol-resistant transconjugants in 15 µg·ml⁻¹ lincomycin plates confers potential *Lactococcus lactis* ltrB (LI.ltrB) insertions within the target tcdR gene in the chromosome of R20291. The presence of a putative tcdR mutant was identified by PCR using tcdR-specific primers (Table S1) in combination with the EBSu universal and ERM primers. Specific single integration of the group II intron into the genome was verified by Southern blotting using a (³²P)dATP-radiolabeled probe specific for the tcdR gene as described previously (15, 22). Complementation of the C. difficile R20291::tcdR mutant is described in Text S1 in the supplemental materials.

**Toxin assays.** Cultures of R20291 and the R20291::tcdR mutant were centrifuged after 10 h in TY medium, and toxin ELISAs were performed as described previously (15). Details are presented in Text S1.

**Sporulation assay (microscopic analysis).** C. difficile cultures were grown overnight in TY medium supplemented with 0.1% taurocholate to induce germination of any spores that were present. Cells were then diluted in TY medium to an OD₅₀₀ of 0.5, and then 100 µl was spread on 70:30 sporulation agar (S8). Plates were incubated at 37°C and monitored for the production of spores. Cells were harvested from the plates after 24 h and were suspended in TY medium for phase-contrast microscopy as described previously (S8). At least four fields per strain were obtained, and the numbers of spores and vegetative cells were counted to calculate the percentage of spores based on the total numbers of spores and vegetative cells. Experiments were performed at least three independent times.

**Sporulation assay (ethanol resistance method).** C. difficile strains were inoculated into and grown on 70:30 sporulation agar as described above. After 24 h of growth, cells were scraped from the plates and suspended in 70:30 sporulation liquid medium to an OD₅₀₀ of 1.0. Cells were immediately serially diluted and plated onto TY agar-0.1% taurocholate to enumerate viable vegetative cells and spores. A 0.5-ml aliquot of the culture was removed from the chamber, mixed with 0.5 ml of 95% ethanol, subjected to vortex mixing, and incubated at room temperature for 15 min. Ethanol-treated cells were serially diluted in 1× phosphate-buffered saline (PBS), returned to the anaerobic chamber, and plated onto TY agar-0.1% taurocholate plates to enumerate spores. After 24 h of growth, CFU were enumerated, and percent sporulation was calculated as the number of ethanol-resistant spores divided by the total number of viable cells (vegetative cells and spores).

**Spore preparation.** Spores were generated and purified as previously described (26, 27). Details are presented in Text S1.

**RNA-seq analysis and quantitative reverse transcription-PCR (qRT-PCR).** RNA-seq analysis was performed at the DNA Core Facility at the University of Missouri, and the data were analyzed using
<figure><img src="image_11.png" /></figure>

methods described previously (60–62). Details of the RNA-seq analysis and the qRT-PCR (19, 55) are
provided in Text S1.

**Germination.** Purified *C. difficile* spores were heat activated at 65°C for 30 min and then placed on ice. Ten microliters of the heat-activated spores was added to reach a final optical density at 600 nm (OD~600~) of 0.5 in 990 µl of BHIS medium (brain heart infusion [Difco] supplemented with 5 g/liter yeast extract and 0.1% L-cysteine) alone or supplemented with a 2, 5, 10, 20, or 50 mM concentration of taurocholic acid (TA). Germination was monitored at 600 nm for 30 min in a PerkinElmer (Waltham, MA) Lambda25 UV/Vis spectrophotometer. The data points at OD~600~ (T~q~) were normalized to the starting OD~600~ value (70). The germination rates and the 50% effective concentration (EC~50~) were calculated using the slopes of the linear portions of the germination plots as described previously (26, 28). The EC~50~ is the concentration of germinant needed to reach 50% of the maximum germination rate. EC~50~s were individually calculated from each germination experiment and are reported as averages with standard errors of the means.

**Spore heat resistance.** Purified spores (nearly 1 × 10⁵) prepared as described above were resus-
pended in 500 µl of water and incubated at 70°C. Samples were removed at 0.5 h, 4 h, and 8 h, serially
diluted in PBS, plated onto TY agar plates with 0.1% taurocholate, and grown anaerobically for 48 h
before counting was performed (44, 45). As a control for non-heat-treated spores, an aliquot was plated
onto TY agar-0.1% taurocholate plates prior to the experiment and colonies were counted as described
above.

**Transmission electron microscopy.** All steps in sample preparation were performed at room temperature using pelleted spores in a 1.5-ml microcentifuge tube, and solutions were prepared in 1× PBS unless indicated otherwise. For transmission electron microscopy, spores (10¹⁰) were fixed for 2 h in a solution of 2% glutaraldehyde–2% paraformaldehyde. The spores were thoroughly rinsed three times in 1× PBS (for 5 min each time) and postfixed with 1% osmium tetroxide with constant rotation for 1 to 2 h. The samples were then washed thrice with 1× PBS (for 5 min each time) and stained *en bloc* with 2% aqueous uranyl acetate for 1 h under light-protected conditions and then washed three times (for 5 min each time) with distilled water. The spores were further dehydrated in a graded 50% (vol/vol)-to-95% (vol/vol) acetone series for 5 min and left in 100% acetone overnight. Infiltration was carried out in graded acetone/EMBED 812/araldite resin (Electron Microscopy Sciences) at ratios of 1:1 and 1:2 for 10 min each time at room temperature with constant rotation and incubated in 100% resin overnight. The resin was cured at 60°C for 24 to 48 h, and thin sections (silver to gold color) were cut and absorbed onto on 200-mesh copper grids. Sections were examined with a transmission electron microscope (CM100; FEI Company) at 100 kV, and images were captured using a side-mounted Hamamatsu digital camera (model C8484) with AMT image capture software version 602.591n.

**Accession number(s).** Sequence data have been deposited in the NCBI GEO database under accession number GSE85395.

SUPPLEMENTAL MATERIAL

Supplemental material for this article may be found at https://doi.org/10.1128/
mSphere.00383-16.

TEXT S1, DOCX file, 0.2 MB.

FIG S1, PDF file, 0.2 MB.

FIG S2, PDF file, 0.1 MB.

FIG S3, PDF file, 0.1 MB.

FIG S4, PDF file, 0.1 MB.

FIG S5, PDF file, 1.5 MB.

FIG S6, PDF file, 0.04 MB.

FIG S7, PDF file, 0.1 MB.

TABLE S1, DOCX file, 0.1 MB.

ACKNOWLEDGMENTS

We thank Nigel Minton, University of Nottingham, for sharing the plasmid pMTL007C-E5 and Robert Fagan for providing plasmid pRPF185. We also thank Jose E. Lopez for technical assistance throughout the study.

R.G. is supported by 1R15AI122173 from NIAID. Funds from the Johnson Cancer Center-KSU and a pilot project to R.G. from CBID-KU (1P20GM113117-01) also supported this work. J.A.S. is supported by award 5R01AI116895 from the National Institute of Allergy and Infectious Diseases.

The content is solely our responsibility and does not necessarily represent the
official views of the funding agencies or the National Institutes of Health.
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