Skip to main content
NCBI home page

This is a preprint.

It has not yet been peer reviewed by a journal.

The National Library of Medicine is running a pilot to include preprints that result from research funded by NIH in PMC and PubMed.

bioRxiv logoLink to bioRxiv
[Preprint]. 2026 Mar 10:2023.05.16.541032. [Version 2] doi: 10.1101/2023.05.16.541032

Acidification-dependent suppression of C. difficile by pathogenic and commensal enterococci

Holly R Neubauer 1, Ibukun M Ogunyemi 1, Alicia K Wood 1, Angus Johnson 1, Avi Z Stern 1, Zainab Sikander 1, Lesly-Hannah Gutierrez 1, Addelis A Agosto 1, Peter T McKenney 1,*
PMCID: PMC13061049  PMID: 41959164

Abstract

Clostridioides difficile and Vancomycin-resistant Enterococcus faecium (VRE) are commonly co-isolated from hospitalized patients. We sought to develop a co-culture biofilm model to characterize interactions between these two opportunistic pathogens. Upon growth in biofilm-promoting media containing added glucose, fructose or trehalose, VRE produces sufficient acid to lower the pH and inhibit growth of C. difficile. We found this effect depended on the carbon source, and that acidification by VRE was necessary and sufficient to suppress C. difficile growth in liquid medium and in cecal content extracts from germ free mice. VRE frequently dominates the intestine of patients administered antibiotics which can predispose to the development of C. difficile infection. We reasoned that it may be possible to suppress C. difficile growth during co-infection with VRE by supplementing the mouse diet with a fermentable sugar. A VRE-dominated gut microbiota may convert the sugar to acid, lower the pH and reestablish colonization resistance to C. difficile. Supplementation of the diet of VRE colonized mice with high levels of fructose neither resulted in a lower pH, nor did it prevent colonization by C. difficile. Taken together, these data suggest that VRE can suppress growth of C. difficile by organic acid production in a carbon source-dependent manner in vitro, however, the mammalian intestine may require sophisticated approaches to lower pH therapeutically.

Introduction

High population diversity in the gut microbiota is generally correlated with health across gastrointestinal diseases. Treatment with antibiotics can disrupt the gut microbiota to the point that a single species can expand and dominate the gut, making up a high percentage of the reads in a 16S microbiome sequencing data set. Treatment with broad-spectrum antibiotics can lead to expansion of VRE (Donskey et al., 2000) and domination of the gut microbiota by the genus Enterococcus. This was observed in humans (Taur et al., 2012) and mice (Ubeda et al., 2010). This domination occurs in hematopoietic stem cell transplant patients where a low diversity microbiota is correlated with mortality (Liao et al., 2021; Taur et al., 2012). Vancomycin-resistant Enterococcus faecium (VRE) is a rising clinical concern that causes difficult-to-treat systemic infections. But it is often forgotten that E. faecium is also a common member of the human gut microbiota, a lactic acid bacterium, and a commonly used probiotic in humans and agricultural animal production (Dubin & Pamer, 2014; Krawczyk et al., 2021; Yahan Wei et al., 2024).

The majority of studies suggest that patients with a low-diversity gut microbiota following antibiotic treatment are also at elevated risk for Clostridioides difficile infection (CDI). Colonization of the gut with VRE and the genus Enterococcus has been correlated with increased risk of CDI in hospitalized human patients (Fujitani et al., 2011; Lee et al., 2017; Smith et al., 2022). Enterococcus species in the gut microbiota were predicted to enhance CDI risk in mathematical modeling that combined data from human and mouse infections (Buffie et al., 2015). In mouse models, genus Enterococcus was also correlated with increased persistence of C. difficile in the gut (Tomkovich et al., 2020) and increased toxin production and virulence under high dietary zinc conditions (Zackular et al., 2016). One other study described correlation of genus Enterococcus abundance with attenuation of C. difficile virulence (De Wolfe et al., 2019). It is possible that strain diversity may contribute to differing results as some commensal enterococci suppress C. difficile growth in vitro (Rolfe et al., 1981).

Two direct tests of the effects of Enterococcus colonization on the CDI mouse model reported higher toxin titer and pathology (Keith et al., 2020; Smith et al., 2022). The mechanism of enhancement of CDI severity in mice was linked to metabolic cross-talk between E. faecalis OG1RF and C. difficile in which arginine supplementation was sufficient to reduce toxin production and pathology, without significantly affecting colonization by either organism (Smith et al., 2022). Furthermore, E. faecalis benefits from the release of host heme caused by C. difficile toxin-mediated damage in mouse models of CDI (Smith et al., 2024).

Both C. difficile and enterococci form biofilms (Ch’ng et al., 2019; Tremblay & Dupuy, 2022). In both species, biofilms contribute to antibiotic tolerance (Dapa & Unnikrishnan, 2013; Holmberg & Rasmussen, 2016) and may be a reservoir for infection recurrence in CDI (Frost et al., 2021; Normington et al., 2021). This study began as an attempt to establish an in vitro co-culture biofilm model of VRE and C. difficile. We quickly noticed that the byproducts of primary metabolism produced by VRE negatively affected C. difficile growth. Here we have established that VRE and commensal enterococci are capable of acidifying unbuffered bacterial growth media below a pH that is growth inhibitory to C. difficile and other commensal and opportunistic clostridia. Under these conditions, we show that acidification of a carbon source is necessary and sufficient for inhibition of C. difficile and clostridia growth. These data suggest that in conditions where lactic acid bacteria such as VRE dominate the local environment, alteration of acid production and pH levels in a carbon source-dependent manner affect the viability of C. difficile in co-culture in vitro. Finally, we modeled the effects of carbon source control in VRE dominated mice by supplementing the diet with high levels of fructose followed by C. difficile infection. Fructose supplementation did not affect C. difficile CFU levels in VRE co-infected mice. In our mouse model of co-infection, fructose supplementation alone was not sufficient to alter C. difficile colonization, suggesting that more complex interventions may be necessary to reproduce the in vitro phenomenon.

Results

Glucose metabolism inhibits C. difficile growth in co-culture with VRE

We began this work as an attempt to establish a dual-species in vitro liquid biofilm model for VRE and C. difficile. Typically for Gram-positive bacteria, glucose is added to the culture media to a final concentration of 0.2–1%, which promotes attachment and production of exopolysaccharide matrix (Dapa & Unnikrishnan, 2013; Donelli et al., 2012; Kristich et al., 2004; Pillai et al., 2004; Toledo-Arana et al., 2001). When we performed a pilot experiment of liquid co-culture growth in Supplemented Brain Heart Infusion (BHI) broth with 0.4% added glucose (total glucose = 0.6%) we found a significant decrease in C. difficile growth at 8 and 24 hours post-inoculation when in co-culture with VRE (Figure 1A). We observed a similar elimination of C. difficile when co-cultured with VRE in Sporulation Media (SM) which lacks an added carbon source, when it was supplemented with 0.4% glucose or greater in co-culture (Figure 1C).

Figure 1. C. difficile is inhibited by VRE during co-culture in excess glucose.

Figure 1.

Open in a new tab

A) Time course CFUs of C. difficile and VRE alone and in co-culture in BHI + 0.4% glucose (0.6% total glucose). Data are combined from 2 independent experiments, n = 3–5 biological replicates per time point, Mann-Whitney test of alone vs. co-culture. B) VRE was grown for 48 hours in BHI + 0, 0.2, 0.4 or 0.6% additional glucose, then filter sterilized to create conditioned media (VRE CM) which was then inoculated with C. difficile. The average pH of the VRE CM is shown above the graph. Data are combined from 2 independent experiments, n = 3–5 biological replicates per time point, Mann-Whitney test. C) Time course CFUs of C. difficile and VRE alone and in co-culture in SM + 0.6% glucose. Data are combined from 2 independent experiments, n = 3–5 biological replicates per time point, Mann-Whitney test of alone vs. co-culture. D) VRE was grown for 48 hours in SM + 0, 0.2, 0.4 or 0.6% additional glucose. CM was filter sterilized to create VRE CM, the average pH of the VRE CM is shown above the graph. It was then inoculated with mid-log C. difficile and grown for 48 hours. Data are combined from 2 independent experiments, n = 5–6 biological replicates per time point, Mann-Whitney test.

Next to determine if the inhibitory factor produced by VRE is soluble and filterable, we generated conditioned media by growing VRE to exhaustion (OD600 ~ 1.8) and filtered it through a 0.22 μm filter. To determine if the inhibition was dose-dependent, we generated VRE conditioned media with escalating levels of added glucose. The conditioned media was then inoculated with C. difficile and grown for 48-hours. We found that increasing the concentration of glucose results in inhibition of C. difficile growth and that the inhibitory factor is soluble and filterable in both BHI and SM (Figure 1B & 1D). Enterococci are closely related to lactic acid bacteria and are known to produce large amounts of acid when cultured in glucose (Ramsey, Hartke, and Huycke 2014). We tested the pH of the VRE conditioned media and found that pH is inversely correlated with glucose concentration. In BHI conditioned media the starting pH was < 5.3 for all tested glucose concentrations, which did not support C. difficile growth (Figure 1B). BHI contains 0.2% glucose as formulated and is unbuffered, likely accounting for the reduced pH. In conditioned SM with 0.4% added glucose and an average starting pH of 6.4, C. difficile growth was not supported (Figure 1D). These data suggest an inhibitory pH threshold of around 6 for C. difficile in VRE conditioned media. We note that C. difficile itself is capable of lowering medium pH when in monocoulture in both SM (pH 6.5) and BHI (pH 5.7) with added glucose, however, VRE lowers pH by an additional unit in both media (Figure S1). When fructose is added to co-cultures during stationary phase, C. difficile CFUs were significantly reduced along with a corresponding drop in pH (Figure S2). These data suggest that inhibition is not limited exponential growth phase. This effect can be bactericidal as resuspending late log-phase C. difficile in VRE-conditioned high glucose media for 2 hours resulted in a significant reduction in C. difficile (Figure S3). To further test if acidification is necessary for inhibition of C. difficile growth in VRE-conditioned media, we grew VRE to exhaustion in SM +/− 0.6% glucose with 100 mM of HEPES, MOPS or PIPES buffer (Figure S4). Buffering with MOPS resulted in a rise in the pH of the conditioned medium from 5.7 to 6.1, which resulted in a partial rescue of C. difficile growth. Taken together these data suggest that in glucose rich media VRE produces organic acids which lower pH and inhibit C. difficile.

The inhibition of C. difficile growth described above could be the result of an inhibitory factor produced by VRE or it could be due to nutrient limitation. To differentiate between these two mechanisms, we first generated VRE conditioned SM in a range of glucose concentrations. Then we created a dilution series of those conditioned media in sterile PBS. VRE conditioned media inhibits C. difficile growth with 0.4% and 0.6% total glucose concentration (Figure 2). When VRE conditioned media containing 0.4% glucose is diluted 1:2 in PBS, it no longer inhibits C. difficile growth. These data confirm that VRE inhibits C. difficile in a manner consistent with the production of organic acids and not via nutrient limitation.

Figure 2: Inhibition by VRE-conditioned media is not caused by nutrient limitation.

Figure 2:

Open in a new tab

VRE conditioned sporulation media (SM) were generated with increasing concentrations of added glucose, filter sterilized, then kept undiluted (A) diluted 1:1 (B) or 1:2 (C) with reduced PBS before inoculation with C. difficile and plating at 48 hours of growth. Data are combined from 4 independent experiments (n=3–4 biological replicates per condition). Kruskal-Wallace 1-way ANOVA with Dunn’s correction vs. 0% glucose.

To determine if VRE-mediated acidification is necessary for inhibition of C. difficile we neutralized VRE conditioned BHI + 0.4% glucose (pH 5) with sodium hydroxide to pH 7. In the neutralized conditioned media C. difficile grew to similar levels as in naive BHI + 0.4% glucose (Figure 3A), suggesting that acidification is necessary for inhibition of C. difficile under these conditions. To determine if acidification of media is sufficient to inhibit C. difficile growth we acidified naive BHI and SM (pH 7.0) with hydrochloric acid and found that C. difficile growth was inhibited at a pH between 5.0–4.5 in acidified BHI and 6.0–5.5 in acidified SM (Figure 3B). Finally, to determine if acidification has the potential to synergize with other secreted effectors in VRE conditioned media, we used HCl to acidify VRE-conditioned SM, which had a starting pH of 7.6, to a range of pH from 7.0 to 4.5 Here we observed an inhibitory pH between 6.0–5.5, similar to the threshold observed in HCl acidification of naive SM (Figure 3C). These data suggest that acidification is the primary inhibitor of C. difficile under these in vitro conditions.

Figure 3: Acidification is necessary and sufficient for inhibition of C. difficile.

Figure 3:

Open in a new tab

A) VRE-conditioned BHI + 0.4% glucose or neutralized with NaOH to a pH of 7 and inoculated with C. difficile for 48 hours before plating and compared to C. difficile growth in naive medium at a pH of 7.0. Data are combined from 3 independent experiments, n=3 biological replicates, Kruskal-Wallis One-way ANOVA with Dunn’s correction vs. BHI + 0.4% Glu. B) BHI and SM were acidified with reduced HCl in 0.5 unit pH increments before inoculation with C. difficile and growth for 48 hours. Data are combined from 3 independent experiments, n=3 biological replicates, a: p = 0.0351, b: p = 0.0084, Kruskal-Wallis One-way ANOVA with Dunn’s correction versus pH 7.0 CFUs for each medium. C) VRE conditioned SM was filter-sterilized and acidified with reduced HCl before inoculation with C. difficile for 48 hours. Data are combined from 3 independent experiments, n=6 biological replicates, Kruskal-Wallis One-way ANOVA with Dunn’s correction versus pH 7.0 SM CFUs for each condition.

VRE can metabolize specific carbon sources into organic acid to inhibit C. difficile

Next we tested the hypothesis that the acidification of any sugar is sufficient to cause VRE to inhibit the growth of C. difficile in conditioned media. We used a panel of simple sugars that differed in their reported metabolism by enterococci. Glucose and fructose were reported to be acidified, while fucose and xylose were reported not to be acidified (Schleifer & Kilpper-Balz, 1984). Tagatose and arabinose acidification was reported to differentiate between E. faecalis and E. faecium (Schleifer & Kilpper-Balz, 1984). We also included the disaccharide trehalose and the polysaccharide inulin which have been implicated experimentally in mouse models of C. difficile infection (Collins et al., 2018; Hryckowian et al., 2018). We generated VRE conditioned SM with 0.6% of each sugar and grew C. difficile for 48 hours before plating to simulate biofilm culture conditions. We found that only VRE conditioned SM containing glucose, fructose and trehalose inhibited growth of C. difficile when compared to growth in the same naive media (Figure 4A). Glucose, fructose and trehalose lowered pH of the conditioned media significantly with mean pH of 6.1, 6.0 and 5.7, respectively. These pH levels are consistent with the inhibitory pH levels described above.

Figure 4: Inhibition of C. difficile by VRE is carbon source dependent.

Figure 4:

Open in a new tab

A) C. difficile was grown in fresh SM + 0.6% of the carbon source (SM + Sugar) indicated on the x-axis or VRE conditioned SM + 0.6% carbon source (VRE CM + Sugar). Starting mean pH of VRE CM + Sugar for each condition is reported above the graph. Data are combined from 3–5 independent experiments per condition, Mann-Whitney test. B) Cytotoxicity on Vero cells of filtered supernatants from A. Data are representative of 2 independent assays, Kruskal-Wallis One-way ANOVA with Dunn’s correction versus SM + Sugar + CD for each condition.

We performed cytotoxicity assays on Vero cells using filtered supernatants from the carbon source screen described above. We found significant reductions in cytotoxicity of VRE conditioned SM containing the acidified carbon sources, glucose, fructose or trehalose, which reflects the killing of C. difficile by VRE (Figure 4B). It is important to note that although glucose can suppress toxin production via catobolite repression and the transcription factor CcpA during exponential phase (Antunes et al., 2011; Dupuy & Sonenshein, 1998; Hofmann et al., 2021), we did not observe a decrease in cytotoxicity in samples containing C. difficile grown in naive SM + 0.6% glucose compared to SM with no carbon source. Our samples were collected at 48 hours post-inoculation, well beyond the exponential phase and likely reflects the total accumulation of toxin over 48 hours of growth. Additionally, in supernatants collected from VRE monocultures, we observed an unexpected but statistically significant increase in cytotoxicity from VRE conditioned SM + 0.6% inulin. This cytotoxicity was not neutralized by C. difficile anti-toxin.

To test the effects of carbon source availability in a more physiologically relevant in vitro system, we utilized extracts from cecal content of germ free mice and compared growth of C. difficile in naive and VRE conditioned filter-sterilized cecal extracts. In cecal content conditioned by VRE, we observed a slight, but not statistically significant (p = 0.13531) increase in C. difficile growth after 48 hours (Figure 5A) which agrees with recent findings that E. faecalis can promote C. difficile growth in vivo (Smith et al., 2022). However, when we added 0.6% glucose as a carbon source to the cecal extract before VRE conditioning, growth of C. difficile was not supported and pH was reduced to 4.01. Addition of 0.6% fucose, which is not acidified by VRE, as a carbon source did not result in a significant difference in C. difficile growth or pH (6.28) between naive and VRE conditioned cecal extract. These data suggest that the byproducts of carbon source metabolism by VRE can suppress C. difficile growth in cecal extracts. In these conditions we did not observe a significant reduction in cytotoxicity of VRE conditioned cecal content extract containing glucose, despite suppression of C. difficile growth below the level of detection (Figure 5B).

Figure 5. Acidification of glucose inhibits C. difficile growth in germ-free cecal content extracts.

Figure 5

Open in a new tab

A) Sterile filtered extracts of cecal content from germ-free mice were prepared with reduced PBS and the indicated added carbon source or equivalent volume PBS and inoculated with VRE and grown for 24 hours. Conditioned cecal content (VRE CCC) were sterile filtered then inoculated with C. difficile and grown for 48 hours followed by plating on selective media. Data are combined from 3 independent experiments, n = 3 biological replicates. Unpaired Welch’s t-test. B) Vero cell toxin tire assay of filtered supernatants following C. difficile culture. Data are representative of two independent experiments, Unpaired Welch’s t-test.

We then tested if acidification-mediated suppression of C. difficile is conserved among enterococci. We generated conditioned SM + 0.6% glucose from a panel of enterococci including E. hirae, and vancomycin sensitive isolates of E. faecuim and E. faecalis. All of the tested strains completely suppressed C. difficile growth and produced conditioned media with a pH below 6 (Figure 6A). We then tested a small panel of Clostridioides species for growth in VRE conditioned media containing glucose (Figure 6B). Growth of C. scindens, C. inoccuum, C. citrone, and C. difficile ribotype 027 strain R20291 were inhibited below the limit of detection, while C. bifermentans showed significantly reduced growth. Acidification was necessary for growth inhibition of all strains as growth was restored when VRE conditioned media containing glucose was neutralized with NaOH. These data suggest that acidification is conserved among enterococci and is necessary and sufficient to inhibit growth of diverse Clostridioides species under these conditions.

Figure 6: Acidification mediated inhibition is conserved among enterococci and clostridia.

Figure 6:

Open in a new tab

A) SM + 0.6 % glucose was conditioned by incubation with the indicated strains on the x-axis followed by inoculation with C. difficile and growth for 48 hours. Mean pH of the conditioned media are displayed below the x-axis (n=3). CFU data are combined from 3 independent experiments, n=5 biological replicates, Kruskal-Wallis One-way ANOVA with Dunn’s correction vs. Naïve SM + Glu B) VRE conditioned SM + 0.6% glucose was created by 48 hours of VRE growth followed by filter sterilization or was neutralized with NaOH to pH7 before filter sterilization and inoculation with C. difficile for 48 hours. Data are combined from 3 independent experiments, n=3 biological replicates, Kruskal-Wallis One-way ANOVA with Dunn’s correction vs. Naive SM + Glu.

Fructose supplementation is not sufficient to restore colonization resistance in mice

To model the effects of supplementation with a fermentable carbon source, we adapted an existing mouse by supplementing with 15% w/v fructose in drinking water during C. difficile and VRE co-infection (Keith et al., 2020). Excess dietary fructose accumulates in the colon and alters the metabolism of VRE (Isaac et al., 2022; Jang et al., 2018). Mice were sensitized to infection through vancomycin and ampicillin in drinking water, followed by colonization with VRE which is resistant to both antibiotics (Keith et al., 2020). Fructose supplementation was added 24 hours before VRE challenge and maintained to the conclusion of the experiment (Figure 7A). Twenty-four hours after colonization with VRE, antibiotics were withdrawn for 48 hours followed by C. difficile infection for 24 hours followed by sampling of cecal content. Contrary to our hypothesis, supplementation with fructose in VRE colonized mice did not affect C. difficile colonization or toxin production (Figure 7B-C). There was a not significant trend towards higher fructose in fructose supplemented control mice (p = 0.07) when compared to naïve mice. However, we did not detect a significant change in fructose levels between CD + VRE mice and CD + VRE + Fructose. We also did not detect a significant change in pH of cecal content between dual colonized mice and dual colonized mice supplemented with fructose. We did, however, detect a rise in the mean pH between Naïve and antibiotics treated mice of 6.30 ± 0.02 to 6.85 ± 0.14 as has been reported in previously (Sorbara et al., 2019). These data suggest that the dietary supplementation attempted here was not sufficient to alter the pH of the cecum.

Figure 7: C. difficile – VRE dual-infection with fructose supplementation.

Figure 7:

Open in a new tab

A) Mice were treated with Ampicillin and Vancomycin in drinking water and 15% Fructose was added to the drinking water of one arm at day −4, followed by VRE colonization on day −3. Antibiotics were withdrawn for 48 hours followed by C. difficile infection on day 0. Data in B-E are combined from 3 independent experiments. B) C. difficile and VRE CFU levels in cecal content measured by selective plating. C) Vero Cell cytotoxicity of filtered cecal content extracts. D) Fructose levels in cecal content measured by an enzymatic assay. E) pH of cecal content. Data was collected with 4 −5 mice per condition combined from 3 independent experiments. Statistics: A and B, t-test with Welch’s correction. C-E, One-way ANOVA with Dunn’s correction.

Discussion

We initially set out to develop an in vitro co-culture biofilm model of VRE and C. difficile using excess glucose to promote adherence of the bacteria to plastic plates (Ðapa et al., 2013). Growth of both species in excess glucose was not supported in co-culture (Figure 1). We found that acidification of glucose is necessary, sufficient and appears to be the primary mechanism by which VRE affects C. difficile growth in the presence of excess glucose in vitro (Figure 3). To test if VRE conditioned media inhibits C. difficile under all growth conditions we altered the available carbon source and found that growth inhibition is limited to contexts in which the carbon source is acidified by VRE. We observed a pH-mediated inhibition of growth in the presence of glucose, fructose and trehalose (Figure 4A). Added glucose also led to pH-mediated growth inhibition when the growth medium was germ-free mouse cecal content extract (Figure 5A). These data suggest that control of the available carbon source is critical to developing in vitro co-culture assays with lactic acid bacteria. We also measured the toxin titre of C. difficile in the carbon source screen and did not find significant changes in toxin production without an underlying inhibition of growth (Figure 4B). This is likely due to the 48-hour time point chosen for analysis, which is beyond the exponential phase in which toxin production is affected by carbon source in single species culture (Dupuy & Sonenshein, 1998). Given the importance of growth phase to toxin production, it is likely that competition for nutrients in dual-species culture may affect the induction of toxin at earlier time points (Smith et al., 2022).

We also detected what is potentially a novel toxicity of VRE towards cultured Vero cells. This cytotoxic activity was present when conditioned medium was prepared from SM containing 0.6% inulin, a fructan dietary fiber. The samples used in the assay were centrifuged and filtered with a 0.2 μm filter, suggesting that the cytotoxic substance is soluble and filterable. Inulin has generally enhanced protection in human fecal chemostat (Hopkins & Macfarlane, 2003) and mouse models of C. difficile infection (Hryckowian et al., 2018). If inulin induces cytotoxicity by VRE, it is possible that supplementation would result in increased virulence in co-infected mice.

In mouse models of infection, C. difficile first begins to accumulate in the caecum 6–12 hours post infection (Koenigsknecht et al., 2015). The colon, particularly the cecum, is densely colonized and is a site of high degrees of population diversity and countless metabolic products. A general feature of the mouse (Shimizu et al., 2021; Sorbara et al., 2019) and human cecum (Koziolek et al., 2015; Nugent et al., 2001) is an acidic pH of 5.0–6.5 at steady state (Brinck et al., 2025). During in vitro culture C. difficile growth and sporulation is limited following as little as a half-unit shift in pH (Wetzel & McBride, 2020). Growth, sporulation and germination are almost completely inhibited at pH of less than 6 (Kochan et al., 2018; Wetzel & McBride, 2020; Woo et al., 2011). Treatment with antibiotics in mice raises colon pH from acidic to slightly alkaline conditions, suggesting that the gut microbiota are necessary for maintaining acidic pH (Shimizu et al., 2021; Sorbara et al., 2019). These data suggest that the pH of the gut lumen, unlike the pH of blood, is not tightly buffered and may be among the many factors that contribute to C. difficile colonization resistance mediated by the gut microbiota. Most bacteria are capable of living in a pH range of 3–4 units and pH may act as a constraint on growth of commensals and opportunistic pathogens (Jin & Kirk, 2018). We found that pH-mediated inibition of C. difficle is at least partly bactericidal (Figure S3). Given that ingested C. difficile spores germinate well in the pH neutral small intestine (Koenigsknecht et al., 2015), it is possible that transit into the acidic colon (pH ~ 6) is a natural restraint on growth and contributes to colonization resistance. In humans, a small study found a significant association between an alkaline fecal pH and C. difficile infection (Gupta et al., 2016).

The pH of the gut lumen is modulated and constrained by inputs from the host diet, and the metabolism of the gut microbiota and the colonic epithelium. The colonic epithelium produces and secretes bicarbonate, which increases the alkalinity of the gut lumen (Alka & Casey, 2014), which could potentially buffer acid production by VRE in our model. However, several studies have noted decreased levels of the bicarbonate transporter DRA (SLC26A3) in the colon of C. difficile infected mice (Coffing et al., 2018; Peritore-Galve et al., 2023; J. Wang et al., 2020). In addition to DRA, toxin-dependent damage also reduced the expression of the host glucose transporter SGLT1 in mice and resulted in an increase in the level of glucose in the stool (Peritore-Galve et al., 2023). The colonic epithelium also produces lactic acid as a byproduct of glycolysis which can be converted into short chain fatty acids by the gut microbiota (Louis et al., 2022). On the bacteria side of this interaction, in cells of C. difficile during active infection, toxin-mediated damage causes the upregulation of carbohydrate PTS importers (Fletcher et al., 2021). Carbohydrate metabolism transcripts as a class, including genes involved in fructose metabolism were increased by both C. difficile and E. faecalis OG1RF during co-culture in vitro. Taken together, these data suggest that C. difficile and VRE may import and metabolize excess fructose in the colon. Primary metabolism by the gut microbiota produces weak organic acids such as lactic acid and short chain fatty acids when grown in a fermentable carbon source (Gregory et al., 2021). The short chain fatty acid butyrate is growth suppressive to C. difficile in vitro and is inversely correlated with colonization in mice and humans (Pensinger et al., 2024, 2023). All these factors together have the capacity to influence the standing pH of the large intestine.

In certain cases, such as in critically ill patients treated with antibiotics who become dominated by VRE (Liao et al., 2021), it may be desirable to enhance the acidity of the gut lumen through administration of a diet or prebiotics that results in acid production and suppression of C. difficile. Our data suggest that supplementation with fructose in drinking water alone was not sufficient to affect C. difficile colonization in VRE co-infected mice, nor was it sufficient to affect the pH of the contents of the cecal lumen (Figure 7). It is possible that more complex formulations of dietary input (Hryckowian et al., 2018; Mefferd et al., 2020) may be necessary to restore C. difficile colonization resistance in a low diversity gut microbiota.

Clinical trials have tested the efficacy of probiotic lactic acid bacteria on C. difficile infections with mixed results (Goldstein et al., 2017; Mills et al., 2018). It is clear, however, that strains from genus Bifidobacterium and Lactobacillus can inhibit C. difficile in vitro by decreasing pH (Fredua-Agyeman et al., 2017) and in some cases similar probiotics have reduced C. difficile virulence in animal models of infection (Wei et al., 2018). For example, lactic acid production by Streptococcus thermophilus lowered the pH of conditioned media and was inversely correlated with C. difficile growth and toxin production in vitro and in a mouse model of infection (Kolling et al., 2012). These data suggest that acidification-mediated effects on growth of C. difficile are not limited to enterococci and may be a general feature of lactic acid bacteria and any bacteria capable of acidifying a carbon source.

Multiple groups have reported that changes in pH affect growth and survival of core taxa of the gut microbiota. For example, during in vitro culture at pH 5.5 multiple human gut isolates from genus Bacteroides failed to grow (Duncan et al., 2009). This pH sensitivity of Bacteroides has been replicated in several batch culture fermenter studies seeded with donor human feces (Firrman et al., 2022; Ilhan et al., 2017; S. P. Wang et al., 2020). These studies also consistently reported reductions in levels of genus Clostridioides (or Clostridium) under low pH conditions. In keeping with the batch fermentation studies mentioned above, we also found that commensal Clostridioides species are subject to a similar pH-mediated sensitivity to VRE-conditioned media as C. difficile (Figure 6). Therefore, efforts to remediate dysbiosis in recurrent C. difficile infection by reintroducing commensal spore-forming anaerobes (Feuerstadt et al., 2022; Louie et al., 2023) may benefit from a different strategy. Here, temporarily limiting acid production by enterococci and other lactic acid bacteria through dietary intervention may favor engraftment of the bacteriotherapy. The conundrum here is similar to that faced by chemists since the beginning of antibiotics development: How do you specifically target pathogenic bacteria, while sparing or promoting the closely related commensal strains necessary for steady-state health? A solution will require a more complete knowledge of the metabolism of pathogenic and commensal clostridia.

Methods

Strains and Growth Conditions

Strains are listed in Table 1. All experiments were conducted in an anaerobic chamber (Coy Laboratory Products), with an atmosphere of 90% N2, 5% CO2 and 5% H2. C. difficile and other clostridia strains were routinely cultured on BHI plates and liquid media (Brain Heart Infusion + 0.5% yeast extract) supplemented with 0.1% taurocholate and 0.3% L-cysteine. VRE and other enterococci were routinely cultured in BHI liquid medium and Enterococcosel (BD Biosciences) plates. For differential selection during co-culture, VRE and enterococci were plated on Enterococcosel (containing 8 μg / mL vancomycin and 100 μg / mL ampicillin for VRE) while C. difficile was plated on BHI plates supplemented with 0.1% taurocholate, 0.3% L-cysteine, 250 μg/mL D-cycloserine, 16 μg/mL cefoxitin. Commensal Clostridia were enumerated by plating on Columbia Blood Agar and counting by distinguishing between colony morphology of VRE.

Media

BHI was made with 36g/L Bacto BHI powder with 5g/L yeast extract, bacterial, autoclaved, then 3 mL filter sterilized 10% L-cysteine was added to the media before aliquoting and reducing in the anaerobic chamber. Sporulation medium was made with 90g Bacto peptone, 5 g Bacto protease peptone, 1 g ammonium sulfate and 1.5 g tris base, then 3 mL 10% L-cystine was added after autoclaving. All liquid and plate media were pH adjusted to 7.0 with HCl unless otherwise specified before autoclaving. Glucose and other sugars used were dissolved into water at a 6% weight per volume ratio before being filter sterilized. Sugars were then added into the media before being placed in the anaerobic chamber or allowed to reduce in the chamber before being added to the media. Inulin was added as described with heating to allow full solubility before being filtered. All plates were poured in 18mL volumes and kept at 4°C until needed. 8–24 hours before using, plates were placed in the anaerobic chamber to reduce fully.

C. difficile growth in VRE and enterococci conditioned media

Mid-log VRE overnight culture was OD-normalized to 0.5 before being inoculated (1:20 dilution) into SM + 0.6% sugar. The culture was grown for at least 24 hours anaerobically at 37°C until the OD600 was > 2 to ensure growth to saturation. Cultures were spun at 1500g x 15 minutes before being filter sterilized with a 0.22 μm filter syringe into 3 mL aliquots into glass tubes. 1.5 mL aliquots were also frozen at −80C for further use in cytotoxicity assays. C. difficile was inoculated 1:60 from mid-log into the VRE-conditioned media and incubated at 37°C for 48 hours before being serially diluted and drop plated for CFU/mL enumeration.

Glucose titration in BHI and SM

BHI and SM were used to create media containing the following glucose concentrations: 0.0%, 0.2%, 0.4% and 0.6%. Media was placed in the anaerobic chamber to reduce before using. VRE was inoculated 1:20 into each glucose concentration of both BHI and SM. Cultures were incubated for 24 hours before being filtered sterilized as described above. C. difficile was inoculated 1:60 into VRE-conditioned media and fresh BHI and SM conditions and incubated for 24 hours before serial dilutions and drop plating for enumeration.

Dilution of inhibition by VRE conditioned Media

VRE conditioned media was prepared by inoculating VRE 1:20 into BHI with 0.0%, 0.2%, 0.4% and 0.6% added glucose. Cultures were grown for 24–36 hours to an OD600 >2 before being spun down and filter sterilized. A subset of the supernatant was taken out of the chamber for pH recording. The VRE-conditioned media was then diluted 0, 1:1, 1:2, 1:4 and 1:8 with sterile PBS. After dilution, 3 mL aliquots were divided into test tubes per each biological replicate. C. difficile mid-log cultures were diluted 1:10 in sporulation media, then added to the aliquots in a 1:60 dilution and grown between 24 hour before drop plating for enumeration.

Acidification and neutralization of SM and BHI

Acidification of SM and BHI was created by titrating HCl into the media to a pH of 7.0, 6.5, 6.0, 5.5, 5.0 and 4.5 before being autoclaved and aliquoted. C. difficile was inoculated into the reduced media 1:60 and incubated at 37°C for 48 hours before being serial diluted and drop plated for enumeration.

Neutralization was performed by growing VRE in SM + 0.6% glucose or BHI + 0.4% glucose to exhaustion before being spun down and filter sterilized. The filtered media pH was recorded to ensure full acidification at or below 5.5, then neutralized to a pH of 7.0 with NaOH. Neutralized media was filter sterilized again and aliquoted into 3 mL test tubes then placed in an anaerobic incubator for 24 hours to ensure full media reduction. 3 mL aliquots of the acidified media were collected to be used as a negative control. After being reduced, C. difficile was inoculated 1:60 into the media conditions. C. difficile was also inoculated 1:60 into fresh media at a pH of 7.0 as a positive control. All media conditions were incubated at 37°C for 48 hours before drop plating and enumeration.

Cecal extract ex-vivo culture

Cecal content from adult germ-free C57BL/6 mice, 12–16 weeks old, were harvested under sterile conditions and frozen at −80C until use. Mice were maintained at an AAALAC accredited facility under an animal protocol approved by the Institutional Animal Care and Use Committee of Boehringer-Ingelheim Pharmaceuticals Inc. Cecal content extracts were prepared at 0.1g/mL of wet weight in reduced PBS in an anaerobic chamber. Cecal content extracts were prepared at 0.1g/mL of wet weight in reduced PBS in an anaerobic chamber. Contents were vortexed and then centrifuged at 1500g for 15 minute before filter sterilization with a 0.2 μm filter. The extract was split into three conditions with the following supplementation: 0.6% of PBS, glucose or fucose. VRE was inoculated 1:20 into half of the conditions described, grown for 24 hours, spun down and filter sterilized to create VRE conditioned extract. Mid-log C. difficile was inoculated 1:60 from an inoculum into naïve extract or VRE conditioned extract and grown for 24 hours at 37°C before serial dilution and drop plating.

Cytotoxicity assays

Vero cells were grown at 10,000 cells per well in a 96-well plate in Eagle’s minimum essential media (EMEM) + 10% Fetal Bovine Serum and 1x Penicillin & Streptomycin overnight. Culture supernatants and mice samples were frozen at −80C until assay. All samples were spun down for 15 minutes at 15,000g before being filter sterilized into new tubes. Each sample was serial diluted down to 10−6 in a fresh 96-well plate with PBS. C. difficile purified toxin (TechLab) was diluted into 1 mL of sterile water as a positive toxin control. 96 μL of each sample was placed in the top row of a 96-well plate and 4 μL of the anti-toxin (TechLab) was added and incubated for at least 20 minutes to ensure full toxin neutralization from the anti-toxin. Samples with antitoxin were serially diluted in the remaining rows down to 10−6 in PBS. Once all toxin, aliquoted samples and antitoxin samples were completed, 100uL of each sample and the dilutions were placed onto their respective Vero-cell wells. Vero cells were incubated at 37°C with 5.0% CO2 overnight. Cytotoxicity was then analyzed with microscopy. Cells that showed rounding were positive for cytotoxicity. To calculate toxin titre, dilutions with less than 80% cell rounding were considered negative, and the previous dilution in the series was considered positive. This dilution number was then used to calculate the Log10 reciprocal toxin titre.

Conservation of enterococci inhibition of clostridia through acidification

We first grew each enterococcal in BHI+0.6% glucose overnight, filter sterilized and inoculated C. difficile to grow for 48 hours at 37°C before serial dilution and drop plating on C. difficile plates for enumeration. We then used VRE to condition SM + 0.6% glucose, spun down and filter sterilized as previously described. We then inoculated mid-log cultures of the remaining clostridia panel into the VRE-conditioned media in a 1:60 dilution and grew C. difficile for 48 hours at 37°C before drop plating. We also performed neutralization and acidification as described above with this clostridia panel.

Buffering of VRE CM in SM + 0.6% Glucose

VRE was grown overnight to an OD of >1.8 in SM or SM + 0.6% glucose with the following buffers; 100mM PIPES, 100mM HEPES, 100mM MOPS. After VRE growth, the samples were plated on enterococcus selective media, the samples were spun down and filter sterilized. An aliquot from each condition was taken to record pH. C. difficile was inoculated 1:10 into each VRE conditioned media and a naive set of conditions. C. difficile was grown for 48 hours before selectively plating.

Mouse co-infection

C57BL6 mice, 7 weeks old, were purchased from Jackson Laboratories and were maintained under an approved protocol of the Binghamton University Institutional Animal Care and Use Committee (21–852). Mice were screened for C. difficile upon receipt by enrichment culture in CC-BHIS-TA medium (Maslanka et al. 2020). Mice were acclimatized for 1 week prior to antibiotics treatment and were maintained in sterile disposable individually ventilated caging with sterile bedding and irradiated food (LabDiet Rodent Diet 20). Mice were treated with 500 mg/L Ampicillin + 250 mg/L vancomycin in drinking water for 4 days, switched to water without antibiotics for 48 hours followed by infection with 2500 CFU of C. difficile VPI-10463 by oral gavage. Mice receiving VRE were gavaged with VRE after 3 days of antibiotics treatment. For mice receiving dietary supplementation, 15% fructose was added to drinking water after 3 days of antibiotics treatment. Samples were collected and transferred immediately into a pre-reduced anaerobic jar (AnareoPack-Anaero) to minimize oxidative damage to vegetative C. difficile during sampling and transport.

Enumerating bacteria of in vivo models

Fecal pellets were collected from mice into sterile, pre-weighed microcentrifuge tubes on the day of acclimation, the day of VRE challenge, the day of C. difficile challenge and 24 hours after. VRE and C. difficile screening during acclimation was done using selective plating. Commensal enterococci was plated on non-selective enterococci plates and was maintained through the duration of the experiments. 24 hours after C. difficile challenge, mice were euthanized using an Euthanix lid for 10 minutes before cervical dislocation. Cecal content was collected into two tubes, one for CFU plating and one for pH measurements. To calculate CFU’s per 1g sample, tubes were weighed before and after sample collections and the raw CFU’s were divided by the sample weight. During C. difficile infection, tubes were placed in an anaerobic box before collecting. Samples were quickly collected from each mouse and put back into the anaerobic box before processing. All samples were resuspended in 1.0 mL pre-reduced PBS before plating. To enumerate spores, samples were diluted 1:1 in PBS and heat treated at 60°C for 30 minutes and vortexed at 15 minutes. Samples were frozen at −80°C until further use after initial CFU plating. Cecal pH was collected by dissolving cecal contents in 200uL nanopure water before pH measurement using an Accumet microelectrode (Fisher).

Fructose Assay

Cecal contents used for CFU plating and cytotoxicity assays were used for fructose concentration analysis by a colorimetric assay kit (NovusBio). The protocol was followed with the following adaptions: a standard curve was created using 0, 500, 800, 1000, 1600, 1800 and 2000 μg/mL fructose and the linear equation was calculated of y= 0.0002x + 0.0831. 25 uL of sample was combined with 1.5mL assay kit solution and boiled for 8 minutes at 100C. A Nanodrop (Thermo-Fisher) was blanked with nanopure water and measured at 285nm to analyze standard values and all sample values.

Supplementary Material

Supplement 1
media-1.docx (287.9KB, docx)

Acknowledgements

Work in this study was supported by startup funds from Binghamton University, the SUNY Research Foundation and NIAID R21AI171634. PTM is an inventor on US Patents 10,646,520, 11,207,374 and 11,471,495 owned by Memorial Sloan Kettering Cancer Center and receives licensing royalties originating from Seres Therapeutics, Inc. and Nestle Health Sciences. We thank Eric Pamer, Alexander Rudensky, John Hambor and Su-Ellen Brown for bacterial strains. Germ free mouse cecal content was acquired using funds from Boehringer Ingelheim Pharmaceuticals, Inc. for an unrelated project.

References

  1. Alka K., & Casey J. R. (2014). Bicarbonate transport in health and disease. IUBMB Life, 66(9), 596–615. [DOI] [PubMed] [Google Scholar]
  2. Antunes A., Martin-Verstraete I., & Dupuy B. (2011). CcpA-mediated repression of Clostridium difficile toxin gene expression. Molecular Microbiology, 79(4), 882–899. [DOI] [PubMed] [Google Scholar]
  3. Brinck J. E., Sinha A. K., Laursen M. F., Dragsted L. O., Raes J., Uribe R. V., Walter J., Roager H. M., & Licht T. R. (2025). Intestinal pH: a major driver of human gut microbiota composition and metabolism. Nature Reviews. Gastroenterology & Hepatology, 22(9), 639–656. [DOI] [PubMed] [Google Scholar]
  4. Buffie C. G., Bucci V., Stein R. R., McKenney P. T., Ling L., Gobourne A., No D., Liu H., Kinnebrew M., Viale A., Littmann E., van den Brink M. R. M., Jenq R. R., Taur Y., Sander C., Cross J. R., Toussaint N. C., Xavier J. B., & Pamer E. G. (2015). Precision microbiome reconstitution restores bile acid mediated resistance to Clostridium difficile. Nature, 517(7533), 205–208. [DOI] [PMC free article] [PubMed] [Google Scholar]
  5. Ch’ng J.-H., Chong K. K. L., Lam L. N., Wong J. J., & Kline K. A. (2019). Biofilm-associated infection by enterococci. Nature Reviews. Microbiology, 17(2), 82–94. [DOI] [PubMed] [Google Scholar]
  6. Coffing H., Priyamvada S., Anbazhagan A. N., Salibay C., Engevik M., Versalovic J., Yacyshyn M. B., Yacyshyn B., Tyagi S., Saksena S., Gill R. K., Alrefai W. A., & Dudeja P. K. (2018). Clostridium difficile toxins A and B decrease intestinal SLC26A3 protein expression. American Journal of Physiology. Gastrointestinal and Liver Physiology, 315(1), G43–G52. [DOI] [PMC free article] [PubMed] [Google Scholar]
  7. Collins J., Robinson C., Danhof H., Knetsch C. W., van Leeuwen H. C., Lawley T. D., Auchtung J. M., & Britton R. A. (2018). Dietary trehalose enhances virulence of epidemic Clostridium difficile. Nature, 553(7688), 291–294. [DOI] [PMC free article] [PubMed] [Google Scholar]
  8. Ðapa T., Leuzzi R., Ng Y. K., Baban S. T., Adamo R., Kuehne S. A., Scarselli M., Minton N. P., Serruto D., & Unnikrishnan M. (2013). Multiple factors modulate biofilm formation by the anaerobic pathogen Clostridium difficile. Journal of Bacteriology, 195(3), 545–555. [DOI] [PMC free article] [PubMed] [Google Scholar]
  9. Dapa T., & Unnikrishnan M. (2013). Biofilm formation by Clostridium difficile [Review of Biofilm formation by Clostridium difficile]. Gut Microbes, 4(5), 397–402. [DOI] [PMC free article] [PubMed] [Google Scholar]
  10. De Wolfe T. J., Kates A. E., Barko L., Darien B. J., & Safdar N. (2019). Modified Mouse Model of Clostridioides difficile Infection as a Platform for Probiotic Efficacy Studies. Antimicrobial Agents and Chemotherapy, 63(7). 10.1128/AAC.00111-19 [DOI] [Google Scholar]
  11. Donelli G., Vuotto C., Cardines R., & Mastrantonio P. (2012). Biofilm-growing intestinal anaerobic bacteria. FEMS Immunology and Medical Microbiology, 65(2), 318–325. [DOI] [PubMed] [Google Scholar]
  12. Donskey C. J., Chowdhry T. K., Hecker M. T., Hoyen C. K., Hanrahan J. A., Hujer A. M., Hutton-Thomas R. A., Whalen C. C., Bonomo R. A., & Rice L. B. (2000). Effect of antibiotic therapy on the density of vancomycin-resistant enterococci in the stool of colonized patients. The New England Journal of Medicine, 343(26), 1925–1932. [DOI] [PMC free article] [PubMed] [Google Scholar]
  13. Dubin K., & Pamer E. G. (2014). Enterococci and Their Interactions with the Intestinal Microbiome. Microbiology Spectrum, 5(6). 10.1128/microbiolspec.BAD-0014-2016 [DOI] [Google Scholar]
  14. Duncan S. H., Louis P., Thomson J. M., & Flint H. J. (2009). The role of pH in determining the species composition of the human colonic microbiota. Environmental Microbiology, 11(8), 2112–2122. [DOI] [PubMed] [Google Scholar]
  15. Dupuy B., & Sonenshein A. L. (1998). Regulated transcription of Clostridium difficile toxin genes. Molecular Microbiology, 27(1), 107–120. [DOI] [PubMed] [Google Scholar]
  16. Feuerstadt P., Louie T. J., Lashner B., Wang E. E. L., Diao L., Bryant J. A., Sims M., Kraft C. S., Cohen S. H., Berenson C. S., Korman L. Y., Ford C. B., Litcofsky K. D., Lombardo M.-J., Wortman J. R., Wu H., Auniņš J. G., McChalicher C. W. J., Winkler J. A., … von Moltke L. (2022). SER-109, an Oral Microbiome Therapy for Recurrent Clostridioides difficile Infection. The New England Journal of Medicine, 386(3), 220–229. [DOI] [PubMed] [Google Scholar]
  17. Firrman J., Liu L., Mahalak K., Tanes C., Bittinger K., Tu V., Bobokalonov J., Mattei L., Zhang H., & Van den Abbeele P. (2022). The impact of environmental pH on the gut microbiota community structure and short chain fatty acid production. FEMS Microbiology Ecology, 98(5). 10.1093/femsec/fiac038 [DOI] [Google Scholar]
  18. Fletcher J. R., Pike C. M., Parsons R. J., Rivera A. J., Foley M. H., McLaren M. R., Montgomery S. A., & Theriot C. M. (2021). Clostridioides difficile exploits toxin-mediated inflammation to alter the host nutritional landscape and exclude competitors from the gut microbiota. Nature Communications, 12(1), 462. [Google Scholar]
  19. Fredua-Agyeman M., Stapleton P., Basit A. W., Beezer A. E., & Gaisford S. (2017). In vitro inhibition of Clostridium difficile by commercial probiotics: A microcalorimetric study. International Journal of Pharmaceutics, 517(1–2), 96–103. [DOI] [PubMed] [Google Scholar]
  20. Frost L. R., Cheng J. K. J., & Unnikrishnan M. (2021). Clostridioides difficile biofilms: A mechanism of persistence in the gut? PLoS Pathogens, 17(3), e1009348. [Google Scholar]
  21. Fujitani S., George W. L., Morgan M. A., Nichols S., & Murthy A. R. (2011). Implications for vancomycin-resistant Enterococcus colonization associated with Clostridium difficile infections. American Journal of Infection Control, 39(3), 188–193. [DOI] [PubMed] [Google Scholar]
  22. Goldstein E. J. C., Johnson S. J., Maziade P.-J., Evans C. T., Sniffen J. C., Millette M., & McFarland L. V. (2017). Probiotics and prevention of Clostridium difficile infection. Anaerobe, 45, 114–119. [DOI] [PubMed] [Google Scholar]
  23. Gregory A. L., Pensinger D. A., & Hryckowian A. J. (2021). A short chain fatty acid-centric view of Clostridioides difficile pathogenesis. PLoS Pathogens, 17(10), e1009959. [DOI] [PMC free article] [PubMed] [Google Scholar]
  24. Gupta P., Yakubov S., Tin K., Zea D., Garankina O., Ghitan M., Chapnick E. K., Homel P., Lin Y. S., & Koegel M. M. (2016). Does Alkaline Colonic pH Predispose to Clostridium difficile Infection? Southern Medical Journal, 109(2), 91–96. [DOI] [PubMed] [Google Scholar]
  25. Hofmann J. D., Biedendieck R., Michel A.-M., Schomburg D., Jahn D., & Neumann-Schaal M. (2021). Influence of L-lactate and low glucose concentrations on the metabolism and the toxin formation of Clostridioides difficile. PloS One, 16(1), e0244988. [Google Scholar]
  26. Holmberg A., & Rasmussen M. (2016). Mature biofilms of Enterococcus faecalis and Enterococcus faecium are highly resistant to antibiotics. Diagnostic Microbiology and Infectious Disease, 84(1), 19–21. [DOI] [PubMed] [Google Scholar]
  27. Hopkins M. J., & Macfarlane G. T. (2003). Nondigestible oligosaccharides enhance bacterial colonization resistance against Clostridium difficile in vitro. Applied and Environmental Microbiology, 69(4), 1920–1927. [DOI] [PMC free article] [PubMed] [Google Scholar]
  28. Hryckowian A. J., Van Treuren W., Smits S. A., Davis N. M., Gardner J. O., Bouley D. M., & Sonnenburg J. L. (2018). Microbiota-accessible carbohydrates suppress Clostridium difficile infection in a murine model. Nature Microbiology, 3(6), 662–669. [Google Scholar]
  29. Ilhan Z. E., Marcus A. K., Kang D.-W., Rittmann B. E., & Krajmalnik-Brown R. (2017). pH-Mediated Microbial and Metabolic Interactions in Fecal Enrichment Cultures. MSphere, 2(3). 10.1128/mSphere.00047-17 [DOI] [Google Scholar]
  30. Isaac S., Flor-Duro A., Carruana G., Puchades-Carrasco L., Quirant A., Lopez-Nogueroles M., Pineda-Lucena A., Garcia-Garcera M., & Ubeda C. (2022). Microbiome-mediated fructose depletion restricts murine gut colonization by vancomycin-resistant Enterococcus. Nature Communications, 13(1), 7718. [Google Scholar]
  31. Jang C., Hui S., Lu W., Cowan A. J., Morscher R. J., Lee G., Liu W., Tesz G. J., Birnbaum M. J., & Rabinowitz J. D. (2018). The Small Intestine Converts Dietary Fructose into Glucose and Organic Acids. Cell Metabolism, 27(2), 351–361.e3. [DOI] [PMC free article] [PubMed] [Google Scholar]
  32. Jin Q., & Kirk M. F. (2018). pH as a Primary Control in Environmental Microbiology: 1. Thermodynamic Perspective. Frontiers of Environmental Science & Engineering in China, 6. 10.3389/fenvs.2018.00021 [DOI] [Google Scholar]
  33. Keith J., Dong Q., Sorbara M. T., Becattini S., Sia J. K., Gjonbalaj M., Seok R., Leiner I., Littmann E., & Pamer E. G. (2020). Impact of antibiotic-resistant bacteria in the mouse intestine on immune activation and Clostridioides difficile infection. Infection and Immunity. 10.1128/IAI.00362-19 [DOI] [Google Scholar]
  34. Kochan T. J., Shoshiev M. S., Hastie J. L., Somers M. J., Plotnick Y. M., Gutierrez-Munoz D. F., Foss E. D., Schubert A. M., Smith A. D., Zimmerman S. K., Carlson P. E. Jr, & Hanna P. C. (2018). Germinant Synergy Facilitates Clostridium difficile Spore Germination under Physiological Conditions. MSphere, 3(5). 10.1128/mSphere.00335-18 [DOI] [Google Scholar]
  35. Koenigsknecht M. J., Theriot C. M., Bergin I. L., Schumacher C. A., Schloss P. D., & Young V. B. (2015). Dynamics and establishment of Clostridium difficile infection in the murine gastrointestinal tract. Infection and Immunity, 83(3), 934–941. [DOI] [PMC free article] [PubMed] [Google Scholar]
  36. Kolling G. L., Wu M., Warren C. A., Durmaz E., Klaenhammer T. R., Timko M. P., & Guerrant R. L. (2012). Lactic acid production by Streptococcus thermophilus alters Clostridium difficile infection and in vitro Toxin A production. Gut Microbes, 3(6), 523–529. [DOI] [PMC free article] [PubMed] [Google Scholar]
  37. Koziolek M., Grimm M., Becker D., Iordanov V., Zou H., Shimizu J., Wanke C., Garbacz G., & Weitschies W. (2015). Investigation of pH and Temperature Profiles in the GI Tract of Fasted Human Subjects Using the Intellicap(®) System. Journal of Pharmaceutical Sciences, 104(9), 2855–2863. [DOI] [PubMed] [Google Scholar]
  38. Krawczyk B., Wityk P., Gałęcka M., & Michalik M. (2021). The Many Faces of Enterococcus spp.—Commensal, Probiotic and Opportunistic Pathogen. Microorganisms, 9(9), 1900. [DOI] [PMC free article] [PubMed] [Google Scholar]
  39. Kristich C. J., Li Y.-H., Cvitkovitch D. G., & Dunny G. M. (2004). Esp-independent biofilm formation by Enterococcus faecalis. Journal of Bacteriology, 186(1), 154–163. [DOI] [PMC free article] [PubMed] [Google Scholar]
  40. Lee Y. J., Arguello E. S., Jenq R. R., Littmann E., Kim G. J., Miller L. C., Ling L., Figueroa C., Robilotti E., Perales M.-A., Barker J. N., Giralt S., van den Brink M. R. M., Pamer E. G., & Taur Y. (2017). Protective Factors in the Intestinal Microbiome Against Clostridium difficile Infection in Recipients of Allogeneic Hematopoietic Stem Cell Transplantation. The Journal of Infectious Diseases, 215(7), 1117–1123. [DOI] [PMC free article] [PubMed] [Google Scholar]
  41. Liao C., Taylor B. P., Ceccarani C., Fontana E., Amoretti L. A., Wright R. J., Gomes A. L. C., Peled J. U., Taur Y., Perales M.-A., van den Brink M. R. M., Littmann E., Pamer E. G., Schluter J., & Xavier J. B. (2021). Compilation of longitudinal microbiota data and hospitalome from hematopoietic cell transplantation patients. Scientific Data, 8(1), 71. [DOI] [PMC free article] [PubMed] [Google Scholar]
  42. Louie T., Golan Y., Khanna S., Bobilev D., Erpelding N., Fratazzi C., Carini M., Menon R., Ruisi M., Norman J. M., Faith J. J., Olle B., Li M., Silber J. L., & Pardi D. S. (2023). VE303, a Defined Bacterial Consortium, for Prevention of Recurrent Clostridioides difficile Infection: A Randomized Clinical Trial. JAMA: The Journal of the American Medical Association, 329(16), 1356–1366. [DOI] [PMC free article] [PubMed] [Google Scholar]
  43. Louis P., Duncan S. H., Sheridan P. O., Walker A. W., & Flint H. J. (2022). Microbial lactate utilisation and the stability of the gut microbiome. Gut Microbiome, 3, e3. [DOI] [PMC free article] [PubMed] [Google Scholar]
  44. Mefferd C. C., Bhute S. S., Phan J. R., Villarama J. V., Do D. M., Alarcia S., Abel-Santos E., & Hedlund B. P. (2020). A high-fat/high-protein, Atkins-type diet exacerbates Clostridioides (Clostridium) difficile infection in mice, whereas a high-carbohydrate diet protects. MSystems, 5(1). 10.1128/mSystems.00765-19 [DOI] [Google Scholar]
  45. Mills J. P., Rao K., & Young V. B. (2018). Probiotics for prevention of Clostridium difficile infection. Current Opinion in Gastroenterology, 34(1), 3–10. [DOI] [PMC free article] [PubMed] [Google Scholar]
  46. Normington C., Moura I. B., Bryant J. A., Ewin D. J., Clark E. V., Kettle M. J., Harris H. C., Spittal W., Davis G., Henn M. R., Ford C. B., Wilcox M. H., & Buckley A. M. (2021). Biofilms harbour Clostridioides difficile, serving as a reservoir for recurrent infection. NPJ Biofilms and Microbiomes, 7(1), 16. [DOI] [PMC free article] [PubMed] [Google Scholar]
  47. Nugent S. G., Kumar D., Rampton D. S., & Evans D. F. (2001). Intestinal luminal pH in inflammatory bowel disease: possible determinants and implications for therapy with aminosalicylates and other drugs. Gut, 48(4), 571–577. [DOI] [PMC free article] [PubMed] [Google Scholar]
  48. Pensinger D. A., Dobrila H. A., Stevenson D. M., Hryckowian N. D., Amador-Noguez D., & Hryckowian A. J. (2024). Exogenous butyrate inhibits butyrogenic metabolism and alters virulence phenotypes in Clostridioides difficile. MBio, 15(3), e0253523. [Google Scholar]
  49. Pensinger D. A., Fisher A. T., Dobrila H. A., Van Treuren W., Gardner J. O., Higginbottom S. K., Carter M. M., Schumann B., Bertozzi C. R., Anikst V., Martin C., Robilotti E. V., Chow J. M., Buck R. H., Tompkins L. S., Sonnenburg J. L., & Hryckowian A. J. (2023). Butyrate Differentiates Permissiveness to Clostridioides difficile Infection and Influences Growth of Diverse C. difficile Isolates. Infection and Immunity, 91(2), e0057022. [Google Scholar]
  50. Peritore-Galve F. C., Kaji I., Smith A., Walker L. M., Shupe J. A., Washington M. K., Algood H. M. S., Dudeja P. K., Goldenring J. R., & Lacy D. B. (2023). Increased intestinal permeability and downregulation of absorptive ion transporters Nhe3, Dra, and Sglt1 contribute to diarrhea during Clostridioides difficile infection. Gut Microbes, 15(1), 2225841. [Google Scholar]
  51. Pillai S. K., Sakoulas G., Eliopoulos G. M., Moellering R. C., Jr, Murray, B. E., & Inouye, R. T. (2004). Effects of glucose on fsr-mediated biofilm formation in Enterococcus faecalis. The Journal of Infectious Diseases, 190(5), 967–970. [DOI] [PubMed] [Google Scholar]
  52. Rolfe R. D., Helebian S., & Finegold S. M. (1981). Bacterial interference between Clostridium difficile and normal fecal flora. The Journal of Infectious Diseases, 143(3), 470–475. [DOI] [PubMed] [Google Scholar]
  53. Schleifer K. H., & Kilpper-Balz R. (1984). Transfer of Streptococcus faecalis and Streptococcus faecium to the Genus Enterococcus nom. rev. as Enterococcus faecalis comb. nov. and Enterococcus faecium comb. nov. International Journal of Systematic Bacteriology, 34(1), 31–34. [Google Scholar]
  54. Shimizu K., Seiki I., Goto Y., & Murata T. (2021). Measurement of the Intestinal pH in Mice under Various Conditions Reveals Alkalization Induced by Antibiotics. Antibiotics (Basel, Switzerland), 10(2). 10.3390/antibiotics10020180 [DOI] [Google Scholar]
  55. Smith A. B., Jenior M. L., Keenan O., Hart J. L., Specker J., Abbas A., Rangel P. C., Di C., Green J., Bustin K. A., Gaddy J. A., Nicholson M. R., Laut C., Kelly B. J., Matthews M. L., Evans D. R., Van Tyne D., Furth E. E., Papin J. A., … Zackular J. P. (2022). Enterococci enhance Clostridioides difficile pathogenesis. Nature, 611(7937), 780–786. [DOI] [PMC free article] [PubMed] [Google Scholar]
  56. Smith A. B., Specker J. T., Hewlett K. K., Scoggins T. R. 4th, Knight M., Lustig A. M., Li Y., Evans K. M., Guo Y., She Q., Christopher M. W., Garrett T. J., Moustafa A. M., Van Tyne D., Prentice B. M., & Zackular J. P. (2024). Liberation of host heme by Clostridioides difficile-mediated damage enhances Enterococcus faecalis fitness during infection. MBio, 15(1), e0165623. [Google Scholar]
  57. Sorbara M. T., Dubin K., Littmann E. R., Moody T. U., Fontana E., Seok R., Leiner I. M., Taur Y., Peled J. U., van den Brink M. R. M., Litvak Y., Bäumler A. J., Chaubard J.-L., Pickard A. J., Cross J. R., & Pamer E. G. (2019). Inhibiting antibiotic-resistant Enterobacteriaceae by microbiota-mediated intracellular acidification. The Journal of Experimental Medicine, 216(1), 84–98. [DOI] [PMC free article] [PubMed] [Google Scholar]
  58. Taur Y., Xavier J. B., Lipuma L., Ubeda C., Goldberg J., Gobourne A., Lee Y. J., Dubin K. A., Socci N. D., Viale A., Perales M.-A., Jenq R. R., van den Brink M. R. M., & Pamer E. G. (2012). Intestinal domination and the risk of bacteremia in patients undergoing allogeneic hematopoietic stem cell transplantation. Clinical Infectious Diseases: An Official Publication of the Infectious Diseases Society of America, 55(7), 905–914. [DOI] [PMC free article] [PubMed] [Google Scholar]
  59. Toledo-Arana A., Valle J., Solano C., Arrizubieta M. J., Cucarella C., Lamata M., Amorena B., Leiva J., Penadés J. R., & Lasa I. (2001). The enterococcal surface protein, Esp, is involved in Enterococcus faecalis biofilm formation. Applied and Environmental Microbiology, 67(10), 4538–4545. [DOI] [PMC free article] [PubMed] [Google Scholar]
  60. Tomkovich S., Stough J. M. A., Bishop L., & Schloss P. D. (2020). The Initial Gut Microbiota and Response to Antibiotic Perturbation Influence Clostridioides difficile Clearance in Mice. MSphere, 5(5). 10.1128/mSphere.00869-20 [DOI] [Google Scholar]
  61. Tremblay Y. D., & Dupuy B. (2022). The blueprint for building a biofilm the Clostridioides difficile way. Current Opinion in Microbiology, 66, 39–45. [DOI] [PubMed] [Google Scholar]
  62. Ubeda C., Taur Y., Jenq R. R., Equinda M. J., Son T., Samstein M., Viale A., Socci N. D., van den Brink M. R. M., Kamboj M., & Pamer E. G. (2010). Vancomycin-resistant Enterococcus domination of intestinal microbiota is enabled by antibiotic treatment in mice and precedes bloodstream invasion in humans. The Journal of Clinical Investigation, 120(12), 4332–4341. [DOI] [PMC free article] [PubMed] [Google Scholar]
  63. Wang J., Ortiz C., Fontenot L., Mukhopadhyay R., Xie Y., Chen X., Feng H., Pothoulakis C., & Koon H. W. (2020). Therapeutic mechanism of macrophage inflammatory protein 1 α neutralizing antibody (CCL3) in Clostridium difficile infection in mice. The Journal of Infectious Diseases, 221(10), 1623–1635. [DOI] [PMC free article] [PubMed] [Google Scholar]
  64. Wang S. P., Rubio L. A., Duncan S. H., Donachie G. E., Holtrop G., Lo G., Farquharson F. M., Wagner J., Parkhill J., Louis P., Walker A. W., & Flint H. J. (2020). Pivotal Roles for pH, Lactate, and Lactate-Utilizing Bacteria in the Stability of a Human Colonic Microbial Ecosystem. MSystems, 5(5). 10.1128/mSystems.00645-20 [DOI] [Google Scholar]
  65. Wei Yahan, Palacios Araya D., & Palmer K. L. (2024). Enterococcus faecium: evolution, adaptation, pathogenesis and emerging therapeutics. Nature Reviews. Microbiology. 10.1038/s41579-024-01058-6 [DOI] [Google Scholar]
  66. Wei Yanxia, Yang F., Wu Q., Gao J., Liu W., Liu C., Guo X., Suwal S., Kou Y., Zhang B., Wang Y., Zheng K., & Tang R. (2018). Protective Effects of Bifidobacterial Strains Against Toxigenic Clostridium difficile. Frontiers in Microbiology, 9, 888. [DOI] [PMC free article] [PubMed] [Google Scholar]
  67. Wetzel D., & McBride S. M. (2020). The Impact of pH on Clostridioides difficile Sporulation and Physiology. Applied and Environmental Microbiology, 86(4). 10.1128/AEM.02706-19 [DOI] [Google Scholar]
  68. Woo T. D. H., Oka K., Takahashi M., Hojo F., Osaki T., Hanawa T., Kurata S., Yonezawa H., & Kamiya S. (2011). Inhibition of the cytotoxic effect of Clostridium difficile in vitro by Clostridium butyricum MIYAIRI 588 strain. Journal of Medical Microbiology, 60(Pt 11), 1617–1625. [DOI] [PubMed] [Google Scholar]
  69. Zackular J. P., Moore J. L., Jordan A. T., Juttukonda L. J., Noto M. J., Nicholson M. R., Crews J. D., Semler M. W., Zhang Y., Ware L. B., Washington M. K., Chazin W. J., Caprioli R. M., & Skaar E. P. (2016). Dietary zinc alters the microbiota and decreases resistance to Clostridium difficile infection. Nature Medicine, 22(11), 1330–1334. [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Supplement 1
media-1.docx (287.9KB, docx)

Articles from bioRxiv are provided here courtesy of Cold Spring Harbor Laboratory Preprints

RESOURCES