c-di-AMP Is Essential for the Virulence of Enterococcus faecalis

ABSTRACT

Second messenger nucleotides are produced by bacteria in response to environmental stimuli and play a major role in the regulation of processes associated with bacterial fitness, including but not limited to osmoregulation, envelope homeostasis, central metabolism, and biofilm formation. In this study, we uncovered the biological significance of c-di-AMP in the opportunistic pathogen Enterococcus faecalis by isolating and characterizing strains lacking genes responsible for c-di-AMP synthesis (cdaA) and degradation (dhhP and gdpP). Using complementary approaches, we demonstrated that either complete loss of c-di-AMP (ΔcdaA strain) or c-di-AMP accumulation (ΔdhhP, ΔgdpP, and ΔdhhP ΔgdpP strains) drastically impaired general cell fitness and virulence of E. faecalis. In particular, the ΔcdaA strain was highly sensitive to envelope-targeting antibiotics, was unable to multiply and quickly lost viability in human serum or urine ex vivo, and was virtually avirulent in an invertebrate (Galleria mellonella) and in two catheter-associated mouse infection models that recapitulate key aspects of enterococcal infections in humans. In addition to evidence linking these phenotypes to altered activity of metabolite and peptide transporters and inability to maintain osmobalance, we found that the attenuated virulence of the ΔcdaA strain also could be attributed to a defect in Ebp pilus production and activity that severely impaired biofilm formation under both in vitro and in vivo conditions. Collectively, these results demonstrate that c-di-AMP signaling is essential for E. faecalis pathogenesis and a desirable target for drug development.

INTRODUCTION

Second messenger nucleotides are synthesized by bacteria in response to internal or external stimuli and are used to reprogram cell physiology through physical interactions with proteins (allosteric regulation), RNA riboswitches, or both (1–3). Despite being a relatively recent discovery, cyclic di-AMP (c-di-AMP) has been shown to control a variety of bacterial processes, including osmoregulation, cell envelope homeostasis, stress and antibiotic tolerance, biofilm formation, central metabolism, DNA repair, genetic competence, and sporulation (4–16). In addition, c-di-AMP plays an important role in host-pathogen interactions, as it can be exported to the bacterial extracellular milieu, promoting a potent STING-mediated type I interferon immune response (17). Two classes of enzymes have been identified as controlling c-di-AMP metabolism. Diadenylate cyclases (DAC) are responsible for c-di-AMP synthesis (from two molecules of ATP or ADP), while phosphodiesterases (PDE) degrade c-di-AMP into 5′-phosphoadenylyl-(3′→5′)-adenosine (5′-pApA) and/or AMP (18). Most bacteria possess a single DAC, with spore-forming bacilli and clostridia being among the few exceptions, encoding three and two DACs, respectively (18). Most bacteria encode at least two types of c-di-AMP PDEs, which have either DHH/DHHA1 (aspartate-histidine-histidine) or HD (histidine-aspartate) catalytic domains (18). These DAC and PDE enzymes are ubiquitous in Gram-positive bacteria but can also be found in Gram-negative bacteria and Archaea (15, 16). Initially, c-di-AMP was thought to be essential as DAC gene deletion strains were not viable, but subsequent studies discovered that c-di-AMP is dispensable if cells are grown in minimal media (10, 19–21). Moreover, intracellular accumulation of c-di-AMP, as in the case of PDE-null mutants or overactive DAC, was also detrimental to cell homeostasis. Thus, c-di-AMP is both essential and toxic and often referred to as an “essential poison” (6). As a result, a growing number of studies have shown that disruption of c-di-AMP homeostasis can greatly diminish the virulence of bacterial pathogens (10, 11, 13, 22–24).

A natural resident of the human gastrointestinal tract, Enterococcus faecalis is also an opportunistic pathogen and leading agent of health care-associated infections, including wound infections, infective endocarditis, catheter-associated urinary tract infection (CAUTI), and central line-associated bloodstream infections (CLABSI) (25, 26). The inherent tolerance of E. faecalis to environmental stresses, including resistance to antibiotics, coupled with the abilities to develop robust biofilms on indwelling medical devices and host tissues and to subvert the immune system make enterococcal infections commonplace and difficult to eradicate (25, 27–30). Previous studies from our group demonstrated that another regulatory nucleotide, the stringent response effector (p)ppGpp, like c-di-AMP, regulates central metabolism, stress tolerance, and biofilm formation in E. faecalis, and that complete lack of (p)ppGpp greatly increases antibiotic sensitivity and attenuates virulence of E. faecalis (31–34). Of interest, evidence indicates that the (p)ppGpp and c-di-AMP signaling networks are intertwined in members of the Firmicutes phylum. Specifically, the activity of two cyclic-di-AMP PDE enzymes, the Staphylococcus aureus GdpP and the Listeria monocytogenes PgpH, is inhibited by (p)ppGpp, such that accumulation of (p)ppGpp leads to c-di-AMP accumulation (35, 36). Two recent studies also showed that c-di-AMP-binding proteins (CbpB in L. monocytogenes and DarB in B. subtilis) are direct activators of the bifunctional (p)ppGpp synthetase/hydrolase Rel enzyme (also known as Rsh or RelA), and that c-di-AMP specifically binds to CbpB/DarB to inhibit its interaction with Rel and, consequently, impairs (p)ppGpp synthesis (37, 38). In addition, the inability of an L. monocytogenes DAC mutant to grow in complex medium was overcome by inactivation of its (p)ppGpp synthetases, thereby linking (p)ppGpp toxicity to c-di-AMP essentiality (19).

In this study, we sought to define the importance of c-di-AMP signaling to E. faecalis pathophysiology. We identified a single DAC (cdaA) and two PDE enzymes (dhhP and gdpP) and showed that genetic manipulations of these genes that lead to either a complete loss of c-di-AMP (ΔcdaA) or c-di-AMP accumulation (ΔdhhP, ΔgdpP, and ΔdhhP ΔgdpP) have profound consequences on the cell fitness and virulence of E. faecalis. Notably, we found that c-di-AMP is critical to E. faecalis pathogenesis, as the ΔcdaA strain was avirulent in multiple animal infection models. In addition to evidence linking c-di-AMP dysregulation to defects in metabolite and peptide transport and the inability to maintain osmotic balance, we discovered that the attenuated virulence of the ΔcdaA strain could also be attributed to a defect in Ebp pilus production and activity, which severely impaired biofilm formation in vitro and in vivo. Collectively, our results underpin that antimicrobial therapies targeting c-di-AMP enzymes and, perhaps, c-di-AMP effector molecules can facilitate the development of new antimicrobial therapies.

RESULTS

Identification of the enzymes responsible for synthesis and degradation of c-di-AMP in E. faecalis.

Through bioinformatic analysis using the amino acid sequences of DAC and PDE orthologues from closely related streptococci, we identified one DAC-encoding gene (gene ID OG1RF_RS08715) and two PDE-encoding genes (OG1RF_RS06025 and OG1RF_RS00060) in the E. faecalis OG1RF genome (GenBank accession no. CP002621.1). Similar to CdaA orthologues (18), the E. faecalis DAC displays a C-terminal DAC domain and three transmembrane helices at the N-terminal domain. Both PDE proteins contain conserved DHH-DHHA1 catalytic domains, and while OG1RF_RS06025 is predicted to be a standalone DHH-DHHA1 cytoplasmic protein, OG1RF_RS00060 possesses two N-terminal transmembrane domains, followed by a PAS (Per-Arnt-Sim) domain, a degenerate GGDEF domain, and the N-terminally located DHH-DHHA1 domain. Based on the presence of these conserved domains and high similarity with previously characterized enzymes, we adopted the cdaA (OG1RF_RS08715), dhhP (OG1RF_RS06025), and gdpP (OG1RF_RS00060) nomenclature. Here, we note that the dhhP gene is also known as pde or pde2 in closely related bacteria (11, 39).

To confirm the predicted functions of the cdaA, dhhP, and gdpP gene products, single (ΔcdaA, ΔdhhP, ΔgdpP) and double (ΔdhhP ΔgdpP) mutants were generated using a markerless in-frame deletion strategy (40). Based on the prediction that c-di-AMP is essential for growth in complex media, the ΔcdaA strain was isolated in a chemically defined medium (CDM) supplemented with 20 mM glucose (41). Upon confirmation that the mutations occurred as designed by Sanger sequencing, we used liquid chromatography-tandem mass spectrometry (LC-MS/MS) to determine the intracellular levels of c-di-AMP in exponentially grown CDM cultures of parent (OG1RF) and mutant (ΔcdaA, ΔdhhP, ΔgdpP, and ΔdhhP ΔgdpP) strains (Fig. 1A). In agreement with the predicted function, no detectable c-di-AMP was found in the ΔcdaA strain, indicating that CdaA is the only enzymatic source of c-di-AMP in E. faecalis. In addition, high intracellular c-di-AMP pools were detected in the ΔgdpP (∼3-fold) and ΔdhhP (∼5-fold) mutants compared to the parent strain. The increase in c-di-AMP in single PDE mutants was additive, as the ΔdhhP ΔgdpP double mutant displayed an ∼8-fold increase in c-di-AMP pools (Fig. 1A).

FIG 1.

Growth characteristics of DAC and PDE mutants. (A) LC-MS/MS quantification of c-di-AMP in E. faecalis OG1RF (WT) and indicated mutant strains grown to mid-log phase in CDM. Statistical analysis was performed using unpaired t test with Welch’s correction on four biological replicates. Error bars represent the standard error of the margin. *, P ≤ 0.05; **, P ≤ 0.01; ***, P ≤ 0.001. (B to F) Growth curves of E. faecalis WT and mutants in CDM (B), BHI (C), CDM with 0.01% peptone (D), and genetic complementation of the ΔcdaA strains in BHI (E) and CDM with 0.01% peptone (F). Data points represent the average and error bars represent the standard error of the margin for nine biological replicates. Statistical analysis was performed using simple linear regression of exponential growth phase (4 to 12 h), and the slope of each mutant’s growth kinetics was compared to the parent strain.

c-di-AMP is essential for growth in the presence of peptides and osmolytes.

The essentiality of c-di-AMP for growth in complex media or in minimal media supplemented with osmolytes or peptides has been demonstrated in several bacteria, including B. subtilis (20), L. monocytogenes (19), and S. aureus (21). Thus, we determined the ability of the ΔcdaA, ΔdhhP, ΔgdpP, and ΔdhhP ΔgdpP strains to grow in the complex medium BHI or in CDM supplemented with low-molecular-weight (MW) osmolytes or peptides. In low-salt and peptide-free CDM supplemented with 10 mM glucose, the ΔcdaA strain grew slower with a slightly extended lag phase and lower growth yields (Fig. 1B). However, the ΔcdaA strain failed to grow in BHI or CDM supplemented with 0.01% peptone or more (Fig. 1C and D). Under these conditions, the ΔgdpP, ΔdhhP, and ΔdhhP ΔgdpP strains displayed no significant differences in growth rate compared to the parent strain (Fig. 1B to D). We also tested the ability of the mutants to grow in CDM supplemented with the low-MW osmolytes carnitine and glycine betaine. The addition of either one of the two osmolytes abolished growth of the ΔcdaA strain while not having an impact on growth of parent or PDE mutant strains (see Fig. S1A and B in the supplemental material). Genetic (in trans) complementation rescued the growth defect of the ΔcdaA strain in BHI as well as in CDM supplemented with peptone, glycine betaine, or carnitine (Fig. 1E and F, Fig. S1C and D). Next, we tested if exogenous addition of c-di-AMP could rescue the growth defects of the ΔcdaA strain in CDM supplemented with peptone or BHI. Interestingly, the addition of purified c-di-AMP, but not c-di-GMP, rescued the growth defect of the ΔcdaA strain in a dose-dependent manner (Fig. 2A to C). However, exogenous addition of c-di-AMP did not rescue the growth of the ΔcdaA strain in BHI media (Fig. 2D).

FIG 2.

Growth characteristics of OG1Rf and ΔcdaA strains in the presence of exogenous c-di-AMP. Growth of E. faecalis OG1RF (WT) and ΔcdaA strains in CDM with 1% peptone that was supplemented with increasing concentrations of c-di-AMP (A and B) or c-di-GMP (C) or BHI supplemented with c-di-AMP only (D). For panels A and B, data points represent the averages from nine biological replicates. For panels C and D, data points represent the averages from three biological replicates. Statistical analysis was performed using simple linear regression of exponential growth phase (2 to 6 h), and the slope of each strain’s growth kinetics was compared to the parent strain.

To date, the essentiality of c-di-AMP has been closely associated with its role in K⁺ homeostasis (20, 42, 43). However, the effect of K⁺ on growth of c-di-AMP mutants varies according to the composition of the growth media and bacterial species. In B. subtilis, a strain lacking c-di-AMP is fully viable under low K⁺ conditions but unable to survive high K⁺ concentrations (20). Paradoxically, increasing osmotic pressure with addition of salts (KCl or NaCl) rescued growth defects of L. monocytogenes and S. aureus c-di-AMP-null strains in complex media (12, 21). Similar to the latter, we found that addition of high salt concentrations (250 to 500 mM KCl or NaCl) partially restored growth of the ΔcdaA strain in CDM-peptone, even though high salt concentrations were mildly inhibitory to cell growth (Fig. S2). To determine the salt tolerance of the ΔcdaA strain, we compared the ability of OG1RF and ΔcdaA strains to grow in (peptone-free) CDM supplemented with 250 or 500 mM either KCl or NaCl. Interestingly, omission of peptone from the growth media enhanced salt sensitivity of both parent and mutant strains, indicating that the protein-peptide-amino acid mixture counteracted the negative effect of salt stress (Fig. S2).

c-di-AMP contributes to E. faecalis antibiotic tolerance.

In other bacteria, alterations in c-di-AMP concentrations lead to defects in cell division, chain length, and aberrant morphology and increased sensitivity to cell envelope-targeting antibiotics (10, 13, 44–46). Using transmission electron microscopy (TEM) and scanning electron microscopy (SEM), we observed an aberrant morphology for the ΔcdaA strain displaying enlarged cells with irregular division patterns and a large number of lysed/dead cells when grown overnight in CDM (Fig. 3); however, the morphologic characteristics of the ΔdhhP ΔgdpP strain were indifferent, largely resembling the parent strain. Next, we tested the susceptibility of DAC and PDE mutants to antibiotics that target cell wall biosynthesis (ampicillin, bacitracin, and vancomycin) or the cell membrane (daptomycin). In general, the ΔdhhP, ΔdhhP ΔgdpP, and ΔcdaA strains showed increased sensitivity to all four antibiotics, although the differences observed in the presence of ampicillin or vancomycin were not significant or were significant but modest (Fig. 4A and B). Interestingly, the ΔdhhP, ΔdhhP ΔgdpP, and ΔcdaA strains were more susceptible to bacitracin or daptomycin, providing the first evidence of a strong phenotypic defect of strains that accumulate c-di-AMP (Fig. 4C and D). Finally, the ΔgdpP strain phenotypically resembled the parent strain with regard to tolerance to all antibiotics tested (Fig. 4).

FIG 3.

Electron microscopy images of E. faecalis OG1RF, Δpde ΔgdpP, and ΔcdaA strains. (Top) Representative transmission electron microscopy (TEM) images. (Bottom) Representative SEM images. Images shown are representative of 20 images that are acquired from one biological sample from each strain, respectively.

FIG 4.

Antibiotic susceptibility of DAC and PDE mutants. Final growth yields of E. faecalis OG1RF (WT) and indicated mutants after 24 h of incubation in CDM supplemented with 2-fold increasing concentrations of ampicillin (A), vancomycin (B), daptomycin (C), and bacitracin (D). Data points represent averages from nine biological replicates. Error bars represent the standard error of the margin. Statistical analysis was performed using unpaired t test with Welch’s correction. *, P ≤ 0.05; **, P ≤ 0.01; ***, P ≤ 0.001; ****, P ≤ 0.0001.

c-di-AMP mediates growth and survival in human serum and urine.

Next, we assessed the ability of parent and mutant strains to grow and survive in pooled human serum or urine ex vivo. The PDE mutants grew and remained viable for 24 h in either serum or urine with modest differences in growth yields observed for ΔdhhA and ΔdhhA ΔgdpP strains in urine. However, the ΔcdaA strain was unable to grow and lost viability (∼1-log CFU reduction in serum and ∼3.5-log CFU reduction in urine) after 8 h of incubation (Fig. 5A and B). The growth/survival defect of the ΔcdaA strain was restored in the genetically complemented strain (Fig. 5C and D). Of interest, we recently showed that transcription of dhhP and gdpP was strongly induced when a mid-log-phase-grown culture of E. faecalis OG1RF was transferred to human urine (32). Here, we found that transcription of the gdpP and dhhP genes was also induced after a 30-min incubation of cells in human serum (∼2.5- and 6-fold, respectively), whereas cdaA mRNA was repressed by ∼2-fold (Fig. 5E). In concurrence with the transcriptional patterns, we detected ∼6-fold reduction in c-di-AMP when comparing intracellular c-di-AMP pools of exponentially growing cell in CDM to cells incubated in serum or urine for 30 min (Fig. 5F).

FIG 5.

Growth and survival of DAC and PDE mutants in serum or urine. (A and B) Number of CFU of E. faecalis OG1RF (WT) and indicated mutants incubated in pooled human serum (A) or pooled human urine (B). (C and D) CFU counts of WT, ΔcdaA, and complemented ΔcdaA (ΔcdaA pCIE::cdaA) strains after 24 h of incubation in pooled serum (C) or pooled urine (D). Data points represent averages from nine biological replicates, and error bars represent the standard errors of the margin. (E) Comparison of reversed transcribed cDNA copy number of cdaA, gdpP, and dhhP in E. faecalis OG1RF grown to mid-log phase in CDM and 30 min after transfer to human serum. (F) LC-MS/MS quantification of c-di-AMP in E. faecalis OG1RF (WT) grown to mid-log phase in CDM or 30 min after transfer to serum or urine. Data points in panels E and F represent averages from nine and five biological replicates, respectively. Dashed lines in panels C and D indicate limit of detection (100 CFU). Statistical analysis was performed using one-way analysis of variance (ANOVA) with Dunnett’s multiple-comparison test. *, P ≤ 0.05; **, P ≤ 0.01; ***, P ≤ 0.001; ****, P ≤ 0.0001.

c-di-AMP is critical for E. faecalis virulence in multiple animal infection models.

Presently, a direct assessment of the significance of c-di-AMP in bacterial pathogenesis is limited to a few bacteria, with most investigations conducted only with strains that accumulate c-di-AMP (10, 11, 45, 47, 48). Here, we used three well-established animal infection models to uncover the significance of c-di-AMP to E. faecalis pathogenesis. In the Galleria mellonella invertebrate model (Fig. 6A), virulence of ΔdhhP and ΔdhhP ΔgdpP strains was attenuated with ΔgdpP strain showing a similar but not statistically significant trend. Most notably, the ΔcdaA strain was nonpathogenic to G. mellonella (Fig. 6A). Next, we used two foreign body-associated mouse infection models that recapitulate some of the environmental and immunological conditions that promote enterococcal infections in humans. In a catheter-associated peritonitis mouse model (49), all three mutants tested (ΔcdaA, ΔdhhP, and ΔdhhP ΔgdpP) were recovered in significantly lower numbers from peritoneal washes, implants, or spleens (Fig. 6B). The total bacteria recovered from animals infected with ΔdhhP and ΔdhhP ΔgdpP mutants was nearly identical (∼1-log₁₀ CFU reduction) and, as expected, the most significant differences were observed in animals infected with the ΔcdaA strain (≥2-log₁₀ CFU reduction), with no ΔcdaA colonies isolated from implanted catheters (Fig. 6B). Following the same pattern, the ΔcdaA strain was virtually nonpathogenic in a CAUTI mouse model, whereas ΔdhhP and, even more so, ΔdhhP ΔgdpP strains showed intermediate phenotypes (Fig. 6C).

FIG 6.

Virulence of DAC and PDE mutants in different animal models. (A) Percent survival of G. mellonella 96 h postinfection with E. faecalis OG1RF (WT) or the indicated mutants. Each curve represents a group of 15 larvae injected with ∼1 × 10⁵ CFU of E. faecalis. Data points represent averages from 3 biological replicates. Statistical analysis was performed using log rank (Mantel-Cox) test. (B) Total CFU recovered after 48 h from peritoneal wash, spleen, or catheter of mice infected with 2 × 10⁸ CFU of WT or the indicated mutants. (C) Total CFU recovered after 24 h from bladder, kidney, or catheter of mice infected with 1 × 10⁷ CFU of the WT or the indicated mutants. For panels B and C, data points represent nine mice infected with three biological replicates. Black line represents the median. Statistical analysis was performed using Mann-Whitney test. *, P ≤ 0.05; **, P ≤ 0.01; ***, P ≤ 0.001; ****, P ≤ 0.0001. The dashed line represents the limit of detection.

c-di-AMP mediates biofilm formation in an Ebp-dependent manner.

In both mouse models, the inability of the ΔcdaA strain to colonize the catheters suggests that c-di-AMP modulates expression and/or activity of virulence factors associated with biofilm formation or surface adhesion. Using a microtiter plate-based assay to quantify surface-adhered biomass, we confirmed this prediction as the ΔcdaA strain formed very poor biofilms that were restored to parent strain levels upon genetic complementation (Fig. 7A). In E. faecalis, robust biofilm formation on host tissue surfaces and urinary catheters is directly associated with expression of the EbpABC pilus; strains lacking ebp genes form poor biofilms and display attenuated virulence in CAUTI and infective endocarditis models (50, 51). To evaluate if the in vitro and in vivo biofilm defects of the ΔcdaA strain was linked to the Ebp pilus, we performed an in vitro adhesion assay incubating normalized mid-log-phase-grown cells in phosphate-buffered saline (PBS) with 50 mM bicarbonate, a condition that stimulates Ebp expression (52). As expected, bicarbonate increased surface adhesion by the parent strain; however, the ability of the ΔcdaA strain to adhere to the plate surface was unchanged under the Ebp-inducing condition (Fig. 7B). Moreover, inactivation of dhhP, gdpP, or both did not affect E. faecalis biofilm formation or surface adhesion (Fig. 7A and B).

FIG 7.

c-di-AMP mediates biofilm formation by modulating Ebp expression and biogenesis. (A) Adherence biofilm biomass quantification of E. faecalis OG1RF (WT) and indicated mutants grown in CDM for 24 h. (B) Adherent cells of WT and indicated mutants under Ebp-inducing conditions (CDM plus 50 mM sodium bicarbonate). Data points represent nine biological replicates assessed in three independent experiments. For panels A and B, statistical analysis was performed using one-way ANOVA with Welch’s correction. (C) Comparison of reversed-transcribed cDNA copy number of ebpA in E. faecalis WT and ΔcdaA strains grown to mid-log phase in CDM. Data points represent six biological replicates. Statistical analysis was performed using unpaired t test with Welch’s correction. **, P ≤ 0.01; ***, P ≤ 0.001; ****, P ≤ 0.0001. Error bars represent the standard error of the margin. (D) EbpA binding to EbpA-specific antibody using ELISA. Data points represent six biological replicates, and statistical analysis was performed using one-way ANOVA with Dunnett’s multiple-comparison test. ****, P ≤ 0.0001. (E) Immunoblot showing Ebp expression in whole-cell lysates, cell wall fraction, and protoplasts of E. faecalis WT and ΔcdaA cells harvested during mid-log growth phase. Immunoblot using a polyclonal anti-DnaK antibody was used as a protein loading control. The images shown are representative of three experiments with three biological replicates. (F) Densitometry analysis of immunoblot band density presented as a ΔcdaA mutant/WT ratio for whole-cell lysate, cell wall fraction, and protoplast fraction. Statistical analysis was performed using unpaired t test. *, P ≤ 0.05; ****, P ≤ 0.0001. (G) Immunoelectron microscopy of WT and ΔcdaA strains using EbpA-specific antibody (1:1,000).

Through quantitative real-time PCR (qRT-PCR) and enzyme-linked immunosorbent assay (ELISA), we found that ebpA transcription was significantly reduced (∼5-fold) in the ΔcdaA strain and, surprisingly, that EbpA levels were below the detection limit in the ELISA, mirroring the ΔebpA strain (Fig. 7C and D). On the other hand, immunoblot analysis revealed that EbpA was produced in the ΔcdaA strain, albeit in smaller amounts than OG1RF (Fig. 7E). Densitometry analysis of the immunoblot revealed significant reduction in Ebp expressed on the cell wall but not the protoplast fraction; however, Ebp expressed in ΔcdaA whole-cell lysate is significantly reduced compared to the parent strain (Fig. 7F). Here, we speculate that the ELISA result was caused by the combination of reduced Ebp production and impaired function, possibly due to defects in protein translocation, pilus assembly, or surface anchoring. To explore some of these possibilities, we used TEM to visualize the spatial organization and abundance of the Ebp pilus on the cell surface. As shown in previous studies (53, 54), the Ebp pilus was detected as extracellular filamentous fibers extending outwards from the cell surface of OG1RF (Fig. 7G). However, these fiber-like structures were completely absent from the ΔcdaA strain, with EbpA evenly distributed around the cell surface (Fig. 7G).

The impact of c-di-AMP on global gene expression.

To evaluate the impact of c-di-AMP dysregulation at a global level, we used RNA sequencing (RNA-seq) to compare the transcriptome of OG1RF, ΔcdaA, and ΔdhhP ΔgdpP strains. The complete absence of c-di-AMP (ΔcdaA) resulted in the differential expression of 302 genes (2-fold cutoff, 147 upregulated and 155 downregulated; Table S1), and c-di-AMP accumulation (ΔdhhP ΔgdpP) resulted in 163 differentially expressed genes (2-fold cutoff, 62 upregulated and 101 downregulated; Table S2). Notably, the ebpABC operon was among the most strongly repressed genes (4.5- to 11-fold downregulation) in the ΔcdaA strain, validating the qRT-PCR quantifications. Among the differently expressed genes, 77 were common for both mutants, with 12 genes showing opposite patterns of expression. Among the genes with opposing expression patterns were genes from a transcriptional unit coding for a putative compatible solute ABC-type transporter (OG1RF_RS10300-RS10305) that was downregulated in the ΔcdaA strain and upregulated in the ΔdhhP ΔgdpP strain. Given the strong association of c-di-AMP with osmoregulation, it was interesting to also observe that transcriptional units coding for polyamine/osmolyte transporters (OG1RF_RS10340-RS10355 and OG1RF_RS05150-RS05165) were downregulated in the ΔcdaA strain. Conversely, a glutamate transport operon (OG1RF_RS06355-65) and an annotated oligopeptide transporter (OG1RF_RS00290) were among the most highly upregulated genes in the ΔcdaA strains. Among genes showing the same expression trend in ΔcdaA and ΔdhhA ΔgdpP strains were a large transcriptional unit (OG1RF_RS07320- RS07370) coding for genes involved in pyrimidine metabolism, a cluster of surface-anchored WxL domain proteins with unknown function (OG1RF_RS02570-RS02595), the murein hydrolase regulator lrgA-lrgB genes (OG1RF_RS12565-RS12570), and the arginine deiminase (ADI) operon (OG1RF_RS490-RS515).

DISCUSSION

To date, c-di-AMP has emerged as one of the most important second messenger nucleotides, with key roles in a variety of cellular processes (15, 16, 43, 55). Furthermore, accumulating evidence showing that c-di-AMP mediates bacterial virulence and the realization that the enzymes that synthesize and degrade c-di-AMP are absent in eukaryote makes c-di-AMP metabolism and, possibly, c-di-AMP-binding proteins desirable targets for drug development. In this study, we expanded the catalog of bacterial species whose general fitness, antibiotic tolerance, and virulence is strongly tied to c-di-AMP signaling by adding E. faecalis, an opportunistic pathogen of great medical concern due to a high prevalence in nosocomial infections and limited treatment options.

Interestingly, we found that exogenous addition of c-di-AMP reverts the growth defect of the ΔcdaA strain in CDM supplement with peptone but not in BHI; we suspect that other metabolites present in BHI somehow inhibit or interfere with c-di-AMP uptake. Because c-di-AMP is negatively charged and, in theory, should not diffuse through the cytoplasmic membrane, this result seems to indicate that c-di-AMP is being actively imported by E. faecalis cells, either via a promiscuous transporter or by a dedicated c-di-AMP transporter. While several investigations have shown that bacteria respond to exogenous c-di-GMP, to our knowledge, only one previous report demonstrated the effect of exogenous c-di-AMP on expression of c-di-AMP-dependent phenotypes. Specifically, exogenous c-di-AMP, mixed with cationic polyamines to facilitate internalization, was shown to promote B. subtilis sporulation (14). With the understanding that working with physiologically relevant concentrations will be crucial, this result indicates that exogenous c-di-AMP can be used as a tool to further explore the scope and physiological consequences of c-di-AMP regulation and to identify new c-di-AMP targets.

A novel aspect of c-di-AMP signaling that also emerges from the present study is that E. faecalis reduces intracellular c-di-AMP pools when exposed to serum or urine, which led us to speculate that this is part of an adaptive bacterial response during infection. Of interest, low c-di-AMP levels have been associated with salt tolerance (56), whereas high c-di-AMP is associated with salt hypersensitivity (57, 58). It follows that salt concentrations in healthy human urine are relatively high (average osmolarity between 300 and 900 mosM/kg H₂O), and the composition and osmolarity of urine in the bladder are constantly undergoing major fluctuations due to cycles of urine concentration and dilution. On the other hand, serum osmolarity does not undergo large fluctuations, and normal osmolarity, ranging between 275 and 295 mosM/kg H₂O, is considerably lower than that in urine. Thus, the environmental cues leading to the observed low levels of c-di-AMP in E. faecalis cells grown in urine or serum and the significance of this observation are unclear and await further investigation, particularly using in vivo infection models. In addition to changes in osmolality or salt concentrations, another possibility is that specific environmental cues present in serum and urine trigger c-di-AMP secretion, which could further deplete intracellular c-di-AMP. In L. monocytogenes, two multidrug efflux pumps were shown to export c-di-AMP from the cytosol during macrophage infection (59) such that a precedent of host-pathogen interactions triggering c-di-AMP secretion seems to already exist.

In bacteria featuring the DhhP-GdpP pair, such as streptococci and staphylococci, the cytosolic DhhP is thought to play a dominant role in c-di-AMP degradation (10). While both DhhP and GdpP degrade c-di-AMP, DhhP cleaves c-di-AMP into the pApA intermediate and then further into the final hydrolysis product, AMP (16, 18). GdpP, on the other hand, cleaves c-di-AMP into pApA only. Because DhhP converts pApA into AMP, it has been proposed that this activity is involved in feedback inhibition of GdpP-dependent c-di-AMP hydrolysis (60). Other than the cumulative effect of DhhP and GdpP on controlling intracellular c-di-AMP pools, loss of GdpP alone, for the most part, did not yield important phenotypes. In fact, the daptomycin tolerance of the ΔgdpP strain was identical to the parent strain, which is in apparent contradiction with a report associating the emergence of a daptomycin-resistant E. faecalis isolate to a GdpP^I440S single-nucleotide polymorphism (SNP) that led to intracellular accumulation of c-di-AMP due to impaired GdpP activity (61). However, the GdpP^I440S strain harbored an additional point mutation in liaR, the response regulator of the LiaFSR system that is known for its critical regulatory role in cell membrane remodeling that is also important for daptomycin tolerance (62).

In the literature, there are several examples whereby c-di-AMP signaling modulates biofilm development and maturation, although the direction of the association of c-di-AMP with biofilm formation differs among bacterial species. For example, high levels of c-di-AMP due to inactivation of PDE-encoding genes reduced biofilms in B. subtilis and Streptococcus gallolyticus (63, 64), while more robust biofilm formation was seen in S. mutans and S. pyogenes (10, 47). Here, we showed that the ΔcdaA strain formed poor biofilms under in vitro conditions (Fig. 7), which is in line with a recent report showing that a DAC inhibitor impairs biofilm formation and exopolysaccharide synthesis of E. faecalis (65). We also found that expression of ebpABC, coding for the biofilm-associated and major virulence factor Ebp pilus, was strongly downregulated in the absence of c-di-AMP. Subsequent analyses revealed that, in addition to reduced mRNA/protein production, Ebp activity was severely compromised in the absence of c-di-AMP (Fig. 7). These results indicate that the adhesion/biofilm defect and attenuated virulence of the ΔcdaA strain can be attributed to defects in Ebp production, subunit translocation, and pilus assembly. At this time, we propose that perturbations in cell envelope homeostasis hinders EbpABC biogenesis, surface anchoring, or both, which would explain why EbpA was undetected in the ELISA and the almost complete absence of pilus-like structures in the ΔcdaA strain. It remains to be discovered how c-di-AMP regulates ebpABC transcription, but initial investigations should focus on probing possible interactions of c-di-AMP with known ebpABC transcriptional regulators such as EbpR, AhrC (also annotated as ArgR3), ArgR2, the Fsr quorum-sensing system, and the RNA-processing enzyme RNase J2 (66–69).

To evaluate the impact of c-di-AMP dysregulation at a global level, we used RNA-seq to compare the transcriptome of E. faecalis OG1RF, ΔdhhP ΔgdpP, and ΔcdaA strains. In B. subtilis, c-di-AMP regulates transcription by interacting with a c-di-AMP-specific riboswitch (2), with several c-di-AMP riboswitch-regulated genes involved in K⁺ and osmolyte transport and cell wall metabolism (70). While enterococcal genomes do not possess sequences that resemble the c-di-AMP riboswitch (2), several genes associated with osmolyte and oligopeptide transport were dysregulated (down- or upregulated) in the ΔcdaA strain. At this time, it is unclear how c-di-AMP affects transcription of E. faecalis genes. One possibility is that an additional (yet-to-be-identified) c-di-AMP riboswitch exist, as in the case of the structurally related c-di-GMP, which has been shown to bind to two distinct c-di-GMP riboswitches (71). A more plausible possibility that has been shown in other bacteria (7, 72–74) but does not invalidate the multiple riboswitch possibility is that c-di-AMP allosterically controls the activity of regulators that control transcription of transport-associated genes. Another important observation that emerges from the transcriptional analysis was the strong downregulation of genes from the arginine deiminase (ADI) system on both ΔcdaA and Δpde ΔgdpP transcriptomes. While the implication of this observation is unclear, it has been proposed that increases in arginine catabolism protects E. faecalis from oxidative stress and is linked to increased antibiotic tolerance (75). The ADI operon also is under the control of two regulators of the ArgR family of transcriptional factors. Based on evidence that ebpABC transcription is also regulated by ArgR-type regulators (AhrC/Arg3 and Arg2) (67), future studies to explore a connection between c-di-AMP, arginine catabolism, and Ebp expression are warranted.

In this report, we have added E. faecalis to the growing list of bacterial pathogens in which fitness, antibiotic tolerance, and virulence are under c-di-AMP control. Using the Galleria mellonella invertebrate model and two foreign body-associated mouse models, we showed that c-di-AMP is critical for the establishment of E. faecalis infections, as the ability of strains that accumulated c-di-AMP to colonize and multiply in the host is severely compromised, whereas the strain lacking only the DAC enzyme (ΔcdaA) is nonpathogenic. We also discovered that expression and biogenesis of the Ebp pilus was markedly impaired in the ΔcdaA strain, linking c-di-AMP signaling with expression of a major enterococcal virulence factor. Collectively, our results provide compelling evidence that approaches to interfere with c-di-AMP signaling might be highly effective for the treatment of E. faecalis infections, a pathogen of great medical concern due to its clinical prevalence and limited therapeutic options.

MATERIALS AND METHODS

Bacterial strains and growth conditions.

The bacterial strains used in this study are listed in Table 1. Strains were grown in brain heart infusion broth (BHI) or in chemically defined medium (CDM) (76) supplemented with 20 mM glucose at 37°C under static conditions. The CDM formulation used for this study is listed in Table S3 in the supplemental material. Strains harboring the pheromone-inducible pCIE plasmid (77) were grown under antibiotic pressure (10 μg ml⁻¹ chloramphenicol) with or without the addition of 5 ng ml⁻¹ cCF10 peptide (Mimotopes, USA). For growth kinetics assays, overnight cultures grown in CDM were adjusted to an optical density at 600 nm (OD₆₀₀) of 0.25 (∼10⁸ CFU ml⁻¹) and inoculated into fresh CDM or BHI at a ratio of 1:50. Growth was monitored at the OD₆₀₀ using an automated growth reader (Oy Growth Curves AB Ltd., Finland). NaCl, KCl, peptone, glycine betaine hydrochloride, carnitine, c-di-AMP, or c-di-GMP (all purchased from Sigma-Aldrich, USA) was added to CDM at the concentrations indicated in the figure legends.

TABLE 1.

Strains and plasmids used in this study

Strain	Relevant characteristic(s)	Source
E. faecalis
OG1RF wild type (WT)	Laboratory strain, Rifr, Fusr	Lab stock
OG1RF Δgdpp	gdpP (OG1RF_RS00060) deletion	This study
OG1RF ΔdhhP	dhhP (OG1RF_RS06025) deletion	This study
OG1RF Δgdpp ΔdhhP	gdpP and dhhP double deletion	This study
OG1RF ΔcdaA	cdaA (OG1RF_RS08715) deletion	This study
CK111	OG1Sp upp4::P23repA4, Specr	Lab stock
OG1RF pCIE	pCIE empty vector, ccF10 pheromone inducible, Cmr	This study
OG1RF Δpde pCIE	pCIE empty vector, ccF10 pheromone inducible, Cmr	This study
OG1RF Δpde pCIE::dhhP	ccF10 pheromone inducible, Cmr	This study
OG1RF ΔcdaA pCIE	pCIE empty vector, ccF10 pheromone inducible, Cmr	This study
OG1RF ΔcdaA pCIE::cdaA	ccF10 pheromone inducible, Cmr	This study
E. coli
EC1000	Host for cloning RepA-dependent plasmids	Lab stock
DH10b	Cloning host	Lab stock

General cloning techniques.

The nucleotide sequences of cdaA, dhhP, and gdpP were obtained from the E. faecalis OG1RF genome via BioCyc (78). The Wizard genomic DNA purification kit (Promega, Madison, WI) was used for isolation of bacterial genomic DNA (gDNA), and the Monarch plasmid miniprep kit (New England BioLabs, Ipswich, MA, USA) was used for plasmid purification. The Monarch DNA gel extraction kit (New England BioLabs, USA) was used to isolate PCR products. Colony PCR was performed using PCR 2× master mix (Promega, USA) with primers listed in Table S4.

Construction of gene deletions and genetically complemented strains.

Deletion of cdaA, dhhP, and gdpP from E. faecalis OG1RF was carried out using the pCJK47 markerless genetic exchange system (40) as previously described. The upstream and downstream sequences of cdaA, dhhP, and gdpP were amplified using the primers listed in Table S4. Introduction of amplicons into the pCJK47 vector, followed by conjugation into E. faecalis OG1RF and isolation of deletion mutants, were carried out as described elsewhere (40) but using CDM to grow the ΔcdaA strain. The ΔdhhP ΔgdpP double mutant was obtained by conjugating the pCJK-gdpP plasmid with the ΔdhhP mutant. All gene deletions were confirmed by PCR sequencing of the insertion site and flanking sequences. The cdaA gene was amplified by PCR using the primers listed in Table S4, digested with the appropriate restriction enzymes, and ligated into pCIE vector to yield plasmid pCIE::cdaA. The plasmid was propagated in E. coli DH10b, verified by sequencing, and electroporated into the E. faecalis ΔcdaA strain as described elsewhere (79).

Antibiotic susceptibility assay.

Strains were grown overnight in CDM, normalized to an OD₆₀₀ of 0.5, and diluted 1:1,000. The diluted cultures were inoculated into fresh medium containing increasing concentrations of antibiotics (ampicillin, bacitracin, daptomycin, or vancomycin; all purchased from Sigma-Aldrich) at a ratio of 1:20 and incubated at 37°C for 24 h. The absorbance at OD₆₀₀ was measured in a Synergy H1 microplate reader (Molecular Devices, USA).

Ex vivo survival in serum and urine.

Strains were grown overnight in CDM, normalized to an OD₆₀₀ of 0.5, and inoculated at a 1:1,000 ratio into pooled human serum or pooled human urine (both purchased from Lee BioSolutions, USA). At selected time points, aliquots were serially diluted in PBS and the dilutions plated on CDM agar for CFU determination.

c-di-AMP quantifications.

Overnight cultures grown in CDM were diluted 1:20 in fresh CDM and grown until the cultures reached an OD₆₀₀ of ∼0.5. One-tenth of the volume of cell cultures was set aside for total protein quantification using a Bradford assay kit (Pierce, USA) according to the manufacturer’s protocol. The cultures were filtered through a 0.45-μm filter (Millipore, USA), and the cells collected on the filter membrane were transferred into a tube containing ice-cold PBS using a cell scraper (Biologix, USA). Cell pellets were obtained by centrifugation at 4,000 × g for 5 min and resuspended into cold extraction buffer (acetonitrile-MeOH-H₂O, 50:40:10; LC-MS grade), adding purified c-di-GMP (Sigma-Aldrich) as an internal standard. Cell suspensions were frozen with liquid nitrogen for 15 min and then incubated at 95°C for 10 min. Samples were mixed with 0.5 ml of 1-mm glass beads and lysed in a bead beater thrice at 45-s intervals. Glass beads and cell debris were pelleted by centrifugation at 20,000 × g for 20 min at 4°C and the supernatants stored overnight at −20°C. The next day, the nucleotide extracts were centrifuged at 10,000 × g for 30 min to remove protein precipitate, and the samples were dried at 40°C using nitrogen gas and then stored at 4°C. The dried samples were shipped to PhenoSwitch Bioscience, Inc. (QC, Canada), for LC-MS/MS (LCMS-8060; triple quadrupole system) analysis. Briefly, the dried samples were resuspended into 300 μl extraction buffer and passed through a solid-phase extraction (SPE) column using a c-di-AMP internal standard. The values obtained were normalized by total protein that was determined using the Bradford assay.

Biofilm assay.

Overnight cultures grown in CDM were normalized to an OD₆₀₀ of 0.5 and further diluted 1:25 in fresh CDM supplemented with 0.175% glucose and added to the wells of 96-well polystyrene plates (Grenier CellSTAR, USA) that were then incubated at 37°C under static conditions for 24 h. After incubation, medium containing planktonic cells was discarded from the wells, which were gently washed twice with PBS. Adherent cells were stained with 0.1% crystal violet for 15 min, and the bound dye eluted in a 33% acetic acid solution. Absorbance was measured at the OD₅₉₅ using the Synergy H1 microplate reader (Biotek, USA).

Adhesion assay.

Pilus-mediated adhesion assay was performed as previously described (80), with minor modifications. Briefly, 96-well polystyrene plates (Grenier CellSTAR) were washed twice with 200 μl PBS and blot dried. Cultures were grown overnight at 37°C in 10 ml CDM, washed with PBS, and normalized to an OD₆₀₀ of 0.5. Next, 100 μl of normalized bacterial culture was added to wells of the microtiter plate, and 50 mM Na₂CO₃ suspended in PBS was added to induce pilus expression as previously described (52). The cells were incubated at 37°C for 2 h, washed twice with 200 μl PBS, and stained with 0.1% (wt/vol) crystal violet as described previously (80). Absorbance was measured at the optical density at 595 nm using the Synergy H1 microplate reader.

G. mellonella infection.

Larvae of G. mellonella were used as a model to assess virulence of E. faecalis strains as described previously (34), with minor modifications. Briefly, groups of 15 larvae (200 to 300 mg in weight) were injected with 5 μl of bacterial inoculum containing ∼1 × 10⁵ CFU that have been normalized from overnight static cultures. Larvae injected with heat-inactivated E. faecalis (30 min at 100°C) or PBS were used as negative and vehicle controls, respectively. Postinjection, larvae were kept at 37°C and survival recorded at selected intervals for up to 96 h.

Mouse CAUTI model.

The methods for the CAUTI experiments have been published (80). Thus, only a general overview and minor modifications are presented here. Female C57BL/6Ncr 6-week-old mice (Charles River Laboratories, USA) were anesthetized and subjected to transurethral implantation of a 5-mm length platinum-cured silicone catheter. Immediately after catheter implantation, mice were infected with ∼1 × 10⁷ CFU of bacteria in PBS. Twenty-four hours postinfection, mice were euthanized and the catheter and organs collected. CFU enumeration was performed by plating catheter and organ homogenates on CDM agar plates selective for E. faecalis OG1RF and derivatives (10 μg ml⁻¹ fusidic acid and 200 μg ml⁻¹ rifampin). This procedure was approved by and performed in compliance with the University of Notre Dame Institutional Animal Care and Use Committee (IACUC).

Foreign body-associated peritonitis mouse model.

The methods for the foreign body-associated peritonitis model have also been published (49), such that only a general overview and minor modifications are described here. The day before surgery, 1-cm-long segments of a silicone catheter tubing (Qosina, USA) were cut and coated in 100 μg ml⁻¹ human fibrinogen solution (Sigma-Aldrich) at 4°C overnight. The following day, female C57BL/6J 8-week-old mice (Jackson Laboratories, USA) were anesthetized and one 1-cm-long catheter segment inserted into the peritoneum using an 18-gauge BD spinal needle/stylette (Becton, Dickinson and Company, USA). Four hours postcatheter implantation, mice were injected via intraperitoneally (i.p.) with ∼2 × 10⁸ CFU of mid-log-grown bacteria in PBS. Animals were euthanized 48 h postinfection and peritoneal wash, catheter, and spleen collected for CFU enumeration by plating serial dilutions on CDM agar plates selective for E. faecalis OG1RF and derivatives. This procedure was approved and performed in compliance with the University of Florida Institutional Animal Care and Use Committee (IACUC).

Electron microscopy analyses.

Transmission electron microscopy (TEM) and scanning electron microscopy (SEM) were performed at the Interdisciplinary Center for Biotechnology Research (ICBR) Electron Microscopy core lab of the University of Florida. SEM and TEM analyses were performed using standard preparation and visualization procedures. For immunogold-labeled TEM analysis, nickel grids (400 Ni-UB) were coated with poly-l-lysine solution at a dilution of 1:10 for 15 min and air dried. Stationary-phase E. faecalis was diluted at 1:1,000, and 5 μl of the diluent was pipetted on the surface of the grid and incubated for 5 min. The bacterial cells were fixed with 4% (wt/vol) paraformaldehyde for 20 min, washed with PBS, and blocked with skim milk for 30 min. After blocking, the grids were washed with PBS thrice and incubated with primary mouse anti-EbpA^Full antibody (80) diluted 1:1,000 for 1 h at room temperature. Postincubation, the grids were washed with PBS thrice and incubated with 12-nm gold donkey anti-goat antibody (1:20 dilution) for 1 h at room temperature. The grids were washed with PBS thrice and then washed with sterile water before being negatively stained with 0.02% uranyl acetate for 10 s and air dried before imaging.

ELISA and immunoblot analyses.

Surface expression of EbpA by E. faecalis OG1RF and mutant strains was determined by ELISA as previously described (81). A duplicate set of samples used for ELISA was used for CFU enumeration, and the absorbance readouts were normalized against the CFU obtained. Samples were prepared for immunoblotting (51, 53), with minor modifications. Briefly, cultures were grown overnight in CDM, diluted 1:20 into fresh CDM, and grown to an OD₆₀₀ of 0.5. Cell pellets were collected by centrifugation, washed with PBS, and incubated with 50 μl of a 30-mg ml⁻¹ lysozyme solution (Sigma-Aldrich) for 20 min to separate cell wall and protoplast fractions. Lysozyme-treated cells were centrifuged at 4,000 × g for 10 min at 4°C. The supernatant collected represents the cell wall fraction, while the cell pellets collected represent the protoplast fraction. Protein extracts were resuspended in 2 × SDS buffer with 2.5% beta-mercaptoethanol and 0.1 M dithiothreitol (DTT) and then boiled at 100°C for 15 min prior to loading onto 6% separating Tris-glycine gel. Immunoblotting was performed using wet transfer to polyvinylidene difluoride (PVDF) membranes in Tris-glycine transfer buffer at 100 V for 90 min at 4°C. After the transfer was complete, the PVDF membrane was soaked overnight in blocking buffer (PBS, 0.05% Tween 20, 3% bovine serum albumin [BSA]) at 4°C with constant agitation. The next day, the membrane was washed thrice with washing buffer (PBS, 0.05% Tween 20), followed by 1-h incubations with anti-EbpA^FULL antibody (1:500) in blocking buffer and secondary goat anti-mouse antibody (Invitrogen, USA) (1:4,000) with washes before and after each incubation. Proteins were visualized using the ECL detection kit (Amersham, USA) in a ChemiDoc imaging system (Bio-Rad, USA). As a loading control, a separate blot was prepared in a similar manner using a rabbit polyclonal anti-S. pyogenes DnaK (1:1,000) antibody (82). Densitometry analysis of the immunoblot was performed using ImageJ to acquire mean region of interest (ROI) for each selected band and the background. Calculation was performed by inverting the ROI values and subtracting the background ROI and then displayed as a ratio to the control band.

RNA analysis.

Total RNA was isolated from E. faecalis cells grown to an OD₆₀₀ of 0.35 by acid-phenol-chloroform extractions, and preparation of cDNA libraries was performed as previously described (83). Sample quality and quantity were assessed on an Agilent 2100 Bioanalyzer at the University of Florida ICBR. RNA deep sequencing (RNA-seq) was performed at UF-ICBR using the Illumina NextSeq 500 platform. Read mapping was performed on a Galaxy server hosted by the University of Florida Research Computer using Map with Bowtie for Illumina and the E. faecalis OG1RF genome (GenBank accession no. CP002621.1) used as a reference. The reads per open reading frame were tabulated with htseq-count. Final comparisons between parent and mutant strains were performed with Degust (http://degust.erc.monash.edu/), with a false discovery rate (FDR) of 0.05 and a 2-fold change in cutoff. Quantifications of cdaA, dhhP, gdpP, and ebpA mRNA were obtained by quantitative real-time PCR (qRT-PCR) as previously described (83).

Statistical analysis.

Data were analyzed using GraphPad Prism 9.0 software (GraphPad Software, San Diego, CA, USA). Data from multiple experiments were pooled and appropriate statistical tests were applied, as indicated in the figure legends. An adjusted P value of ≤0.05 was considered statistically significant (*, P ≤ 0.05; **, P ≤ 0.01; ***, P ≤ 0.001; ****, P ≤ 0.0001).

Data availability.

Gene expression data have been deposited in the NCBI Gene Expression Omnibus (GEO) database (https://www.ncbi.nlm.nih.gov/geo) under GEO Series accession number GSE174381.