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. 2023 Sep 27;9(10):1878–1888. doi: 10.1021/acsinfecdis.3c00192

Correlating Antibiotic-Induced Dysbiosis to Clostridioides difficile Spore Germination and Host Susceptibility to Infection Using an Ex Vivo Assay

John H Moore †, Armita Salahi †, Carlos Honrado †, Christopher Warburton †, Steven Tate †, Cirle A Warren ‡,*, Nathan S Swami †,§,*
PMCID: PMC10581205  PMID: 37756389

Abstract

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Antibiotic-induced microbiota disruption and its persistence create conditions for dysbiosis and colonization by opportunistic pathogens, such as those causing Clostridioides difficile (C. difficile) infection (CDI), which is the most severe hospital-acquired intestinal infection. Given the wide differences in microbiota across hosts and in their recovery after antibiotic treatments, there is a need for assays to assess the influence of dysbiosis and its recovery dynamics on the susceptibility of the host to CDI. Germination of C. difficile spores is a key virulence trait for the onset of CDI, which is influenced by the level of primary vs secondary bile acids in the intestinal milieu that is regulated by the microbiota composition. Herein, the germination of C. difficile spores in fecal supernatant from mice that are subject to varying degrees of antibiotic treatment is utilized as an ex vivo assay to predict intestinal dysbiosis in the host based on their susceptibility to CDI, as determined by in vivo CDI metrics in the same mouse model. Quantification of spore germination down to lower detection limits than the colony-forming assay is achieved by using impedance cytometry to count single vegetative bacteria that are identified based on their characteristic electrical physiology for distinction vs aggregated spores and cell debris in the media. As a result, germination can be quantified at earlier time points and with fewer spores for correlation to CDI outcomes. This sets the groundwork for a point-of-care tool to gauge the susceptibility of human microbiota to CDI after antibiotic treatments.

Keywords: Microbiota, Bacteria, Microfluidics, Cytometry, Antibiotics

The human gut is colonized by a diverse community of microorganisms that work in symbiosis with their host to resist colonization by opportunistic pathogens.1 Loss of diversity in composition of gut microbiota, due to antibiotic administration and/or aging,2,3 is widely linked to intestinal dysbiosis, degradation of immune functions, and onset of various diseases.4 Hence, tools capable of measuring differential degrees of microbiota disruption and its impact on host susceptibility to infection by pathogenic bacteria are of great interest. Currently, metagenomics is used to systematically characterize alterations in the diversity of microorganisms,5 but it can only be used in specialized lab settings. Also, given the system-level effects of each strain and/or metabolite, their aggregate effects on susceptibility of gut microbiota to colonization by specific opportunistic pathogens is difficult to screen. Furthermore, given the heterogeneity in microbial composition and host factors among individuals, it is challenging to generalize the role of specific strains or metabolites for determining susceptibility to infection.6 Instead, this study focuses on a novel tool for correlation of antibiotic-induced dysbiosis to infection susceptibility in the same host model. Specifically, an assay is developed for correlating dysbiosis in mice after varying degrees of antibiotic treatment to their susceptibility for infection by Clostridioides difficile by using their fecal supernatant as the ex vivo sample for C. difficile spore germination.

C. difficile infection (CDI) occurs due to C. difficile overgrowth upon disruption of the host gut microbiome, causing the secretion of toxins (toxin A, or TcdA, and toxin B, or TcdB) that lead to intestinal disease by inhibiting the Rho family of small GTPases. These disturb signal transduction mechanisms at cell–cell junctions, cause dysfunction of the actin cytoskeleton, and induce apoptosis.7−9 CDI is the most common cause of nosocomial (hospital-acquired) infection in the United States, surpassing methicillin-resistant Staphylococcus aureus (MRSA) infection.10 It is responsible for nearly a half-million annual incidences and 29,000 fatalities in the U.S.11−13 Current therapies include antibiotics—metronidazole, vancomycin, or fidaxomicin.14 However, a significant portion of patients (∼25%) exhibit recurrent CDI (rCDI), with 35–65% experiencing multiple episodes of rCDI.15−17 This recurrence is linked to the germination and outgrowth of C. difficile spores present from the initial infection or to re-colonization from persistence of microbiota alterations.18,19 Strategies to help with the recovery of the host microbiota, such as the introduction of probiotic species20,21 or fecal transplant,22 have shown promise in the prevention of rCDI. However, standardization of these strategies is challenging due to differing microbiota contents between patients, highlighting the need for strategies to measure host recovery from dysbiosis and their susceptibility to CDI during the recovery.

CDI is transmitted through ingestion of metabolically dormant spores or their persistence from a prior infection, which then germinate in the colon. Germination of C. difficile spores is induced through a complex process involving exposure to metabolites in the colon, including primary bile salts,23d-glycine,24 and pH.25 Commensal gut microbiota secrete hydrolase enzymes that metabolize primary bile salts into secondary bile salts, such as chenodeoxycholates, which inhibit C. difficile spore germination in the colon.26 Dysbiosis induced by antibiotic exposure prevents the conversion of primary to secondary bile acids, thereby facilitating spore germination27 (Figure 1A). We hypothesize that the in vivo metabolite conditions leading to C. difficile spore germination in the gut are reflected in the sterile-filtered fecal supernatant from the antibiotic-treated mouse model. Hence, incubation of C. difficile spores in this ex vivo sterile-filtered fecal supernatant can simulate the in vivo conditions for spore germination (Figure 1B), thereby enabling the correlation of spore germination under varying degrees of dysbiosis to host susceptibility to CDI.

Figure 1.

Figure 1

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(A) Dysbiosis under antibiotic treatment promotes C. difficile spore germination. (B) Stool sample collection from antibiotic-treated mouse model and preparation of fecal supernatant for ex vivo co-culture with C. difficile spores to assess spore germination by microfluidic impedance cytometry of vegetative cells (4 h) and validation by CFU analysis (24 h). (C) The impedance phase at 10 MHz (ϕZ10 MHz) is used to gate vegetative bacteria vs spore aggregates (details on threshold in Figure 5), to quantify spore germination based on the % bacteria determined from events beyond the threshold. (D) The same mouse model is subjected to a C. difficile challenge, to correlate in vivo metrics of CDI to ex vivo spore germination.

Germination of C. difficile spores to vegetative bacteria involves major alterations to cellular structure, physiology, and function. This alteration needs to be measured at high sensitivity since colonization by the vegetative form can arise from low numbers of C. difficile spores. Measurement of spore germination by colony-forming unit (CFU) counts is labor intensive, time-consuming (24–48 h), and not sensitive due to serial dilution and plating steps on nutrient agar plates, which are difficult to translate to point-of-care application. Single-bacteria cytometry is highly sensitive, especially since C. difficile in vegetative form28 (rod-shape of ∼0.5 μm by ∼3 μm) differs in size and shape from its spore form (∼0.5 μm sphere) (Figure 1A). However, spores form aggregates in the fecal supernatant. Hence, their distinction from the vegetative form requires metrics other than the external size. The cell interior of the vegetative C. difficile form, which includes the cell wall (∼30 nm) and cell membrane (∼8 nm thick) enveloping the conductive cytoplasmic region, differs in electrical physiology from the aggregated spores, which only have insulating endosporium and exosporium layers (Figure S1A). Using impedance-based flow cytometry, we developed a method to probe the interior cellular regions for distinguishing the two forms based on electrical physiology at single-particle sensitivity (Figure S1B).29 Specifically, using electric fields at 10 MHz, wherein the field penetrates through the cell envelope30,31 the conductive interior of vegetative C. difficile causes distinctly higher impedance phase shifts vs that of its spore form, after signal normalization against co-flowing insulating beads. By comparison of C. difficile bacteria to insulating polystyrene beads of equivalent volume that are opaque to high-frequency fields (Figure S1C–E) and to “heat-inactivated” C. difficile cells that exhibit lower membrane capacitance (Figure S1F), an optimized impedance phase threshold (Figure 1C) is deduced (based on details in description of Figure 5) to quantify single vegetative C. difficile cell numbers by impedance cytometry. Our prior work29 utilized this metric to measure progressive enhancement of spore germination with addition of primary bile salts that are known to enhance germination. This enables quantification within just 4 h of spore incubation in fecal supernatant conditioned media, in absence of laborious sample preparation steps, whereas the CFU assay requires >24 h, thereby indicating potential for point-of-care translation.32 Herein, we validate the ex vivo spore germination assay in the fecal supernatant of a mouse model with varying degrees of dysbiosis due to antibiotic treatment for correlation to CDI disease parameters in the same mouse model, so that host susceptibility to C. difficile infection can be predicted.

Figure 5.

Figure 5

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Setting the threshold for impedance phase (ϕZ10 MHz) at 218° based on homogeneous spore sample vs near-complete germinated sample of vegetative bacteria: (A) Scatter plot of Z10 MHz vs ϕZ10 MHz. (B) Histogram of ϕZ10 MHz, with the gate set to capture >95% of spore and vegetative events on either side for the respective samples. Using this ϕZ10 MHz threshold to compare spore vs vegetative events after 4 h co-culture of C. difficile spores with fecal supernatant from (C) untreated and (D) clindamycin-treated mice. (E) ϕZ10 MHz histogram showing reduction in spore events and rise in vegetative events for untreated (364 vegetative events) vs clindamycin-treated (462 vegetative), for 3090 total events in both samples.

Dysbiosis through increasing antibiotic exposure results in collateral disruption of the host gut microbiota, leading to greater susceptibility to initial CDI and its recurrence.33−37 Our novel contribution is the ability to correlate the level dysbiosis in the host to its susceptibility to CDI by coupling spore germination in an ex vivo sample to in vivo CDI metrics in the same animal model (Figure 1B–D). Impedance cytometry is used to enumerate vegetative bacteria within small volumes of the fecal supernatant at low detection limits down to 32 vegetative bacteria in a 50 μL sample,29 whereas CFU assays are prone to variability under small numbers of vegetative bacteria due to need for serial dilutions and limitations in spatial extent on the plate to count the colonies. As a result, quantification of C. difficile spore germination by impedance cytometry for prediction of CDI is more sensitive vs the CFU method, so that germination can be assessed at earlier time points and/or with fewer spore numbers, as encountered in vivo. This sets the groundwork for follow-up work to develop a rapid point-of-care tool to gauge the susceptibility of microbiota to CDI after antibiotic treatments.

Results and Discussion

The feasibility for using fecal supernatant from antibiotic-treated mouse models as a surrogate to probe dysbiosis is first studied based on C. difficile spore germination in this ex vivo sample (Figure 2). Next, the timeline for antibiotic-induced dysbiosis in the mouse model prior to the C. difficile challenge to cause CDI is explored (Figure 3), for its utilization to study CDI metrics under varying levels of dysbiosis (Figure 4). The impedance phase threshold to differentiate spores vs vegetative bacteria is then deduced (Figure 5). Finally, the optimized conditions for varying dysbiosis and inducing CDI are then utilized to validate the ability of ex vivo spore germination, as quantified by impedance cytometry, to detect the specific level of dysbiosis that leads to CDI (Figure 6), and to monitor the persistence of dysbiosis (Figure 7).

Figure 2.

Figure 2

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Correlating spore germination in fecal supernatant media to clindamycin-induced dysbiosis. (A) Timeline for antibiotic treatment, with stool collection pre- (D-5) and post-antibiotics (D0). CFU assay of germination with (B) VPI10463 spores and (C) UK1 spores. Optical density (absorbance at 589 nm) for growth determination in liquid media of (D) VPI10463 spores and (E) UK1 spores.

Figure 3.

Figure 3

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Clindamycin-treated single-antibiotic mouse model of CDI shows disease onset after challenge with spores. (A) Experimental timeline: mice (n = 4 per group) were given clindamycin at 25 mg/kg/day via oral gavage for 5 days to disrupt gut microbiota, and then challenged with C. difficile UK1 spores at 105 CFU (VPI10463 vegetative and spore form in Figure S3). Their disease metrics of (B) weight loss and (C) clinical score were monitored for 7 days (D1–D7), while their stool was analyzed for (D) C. difficile toxin A/B based on ELISA and (E) shedding of C. difficile via qPCR.

Figure 4.

Figure 4

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Increasing the duration of clindamycin treatment prior to C. difficile challenge increased CDI severity. (A) Mice (n = 7 per group) were given clindamycin (25 mg/kg/day) for 10, 5, or 1 days with an untreated control group, then challenged with UK1 spores at 105 CFU. Mice were monitored for 10 days to measure disease severity. (B) Weight change from baseline: mice lost increasing amounts of weight based on the duration of clindamycin pre-treatment. Significance levels shown are in comparison to the vehicle control group (****p < 0.0001, ***p < 0.001, **p < 0.01, *p < 0.05) with a mixed-effects model (REML) with Tukey’s multiple comparisons post-test. (C) Clinical scoring: mice had increasing disease severity based on the duration of clindamycin pretreatment. All statistical comparisons are to baseline, with mixed-effects model (REML) with Dunnett’s multiple comparisons post-test. (D) Survival: mortality increased with the duration of clindamycin given pre-infection, although not significantly (Log-rank test). (E) C. difficile shedding in stool, measured via qPCR, found similar levels in all clindamycin-treated groups on days 1 and 3, with animals receiving clindamycin for 1 day prior to infection showing drop-off in shedding on days 5, 7, and 10, in comparison with the 10-day clindamycin-treated group.

Figure 6.

Figure 6

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Germination of UK1 spores in fecal supernatants from the antibiotic-treated mouse model (0–10 day abx) based on the CFU assay (A, C) and % bacteria from impedance cytometry (B, D) for the pre-infection (A, B) and post-infection groups (C, D). Compared with CFU assay, spore germination determined by impedance cytometry correlated with CDI metrics and the duration of antibiotic (abx) treatment.

Figure 7.

Figure 7

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Measuring persistence of dysbiosis after termination of antibiotic treatment using the spore germination in the fecal supernatant from mice with varying durations of clindamycin treatment. (A) Timeline for mice (n = 5 per group) after treatment with clindamycin for 1 or 5 days (with an untreated control) and stool collection over 4 days following clindamycin treatment for the spore germination assay. (B) Germination rate determined by impedance cytometry as % bacteria indicates significantly higher levels over respective baseline levels from untreated mice for D1–D3 after 5-day clindamycin treatment, but only for D1 after 1-day clindamycin treatment. * indicates significant difference (p < 0.05) over baseline (untreated) on each day of stool.

Dysbiosis Enhances Germination of Spores

Antibiotic-induced dysbiosis disrupts microbiota, leading to increased levels of primary vs secondary bile acid, which promotes germination of C. difficile spores.25−27 To quantify this correlation, we studied spore germination in an ex vivo filtered fecal supernatant stool sample that is collected from mice, pre- vs post-antibiotic treatment. For this purpose, dysbiosis was induced by an antibiotic cocktail that we have previously used to induce susceptibility to severe CDI,34−37 which includes vancomycin, colistin, gentamicin, and metronidazole for 3 days, followed by 2 days with no antibiotics to allow mice to pass any remaining antibiotics out of their system, and followed by a clindamycin injection. Per Figure 2A, stool was taken from mice just prior to antibiotic treatment (pre-antibiotics) and 1 day following clindamycin treatment (post-antibiotics) for spore germination analysis, with the respective analysis in PBS media serving as the control. For spore germination, two C. difficile strains—VPI10463, a non-epidemic, laboratory strain, and UK1, a hypervirulent, epidemic clinical strain—were examined using the CFU assay in solid media (Figure 2B,C) and growth assay in liquid BHI media (Figure 2D,E). Based on the CFU assay, germination of UK1 spores was significantly higher (two-tailed student t test: ****p < 0.0001) post-antibiotics vs pre-antibiotics (Figure 2C).

On the other hand, VPI10463 spores showed less significant increases when cultured in stools, post-antibiotics vs pre-antibiotics, but exhibited similar trends (Figure 2B). Growth differences in liquid media showed a similar trend, but significant differences in growth (one-way ANOVA with Sidak’s multiple comparisons post-test) are apparent only after 20.6 h for VPI10463 spores (Figure 2D) in pre- vs post-antibiotic-treated supernatant, while significant differences are not apparent following growth with the UK1 spores (Figure 2E). These results indicate the feasibility of utilizing sterile filtered fecal supernatant from the mouse model for ex vivo spore germination studies with C. difficile spores, as a surrogate for antibiotic-induced dysbiosis in the mouse model. However, there are differences between the CFU assay performed in solid media and the growth assay performed in liquid media. Since significant differences occur for growth assay in liquid media only after 20.6 h, which is well after the onset of logarithmic bacterial growth, we perform all future validation of the germination of UK1 spores using the CFU assay in solid media, since it follows spore germination more closely than the growth assay.

Clindamycin-Treated Mouse Model of CDI

To establish the timeline for antibiotic-induced dysbiosis followed by the C. difficile challenge for rendering the mouse model susceptible to CDI, a single-antibiotic model based on clindamycin was used. The duration of treatment was altered to modify the degree of dysbiosis and host susceptibility to CDI. Clindamycin is the antibiotic that presents the highest risk for development of CDI, with an odds ratio of 16.8.38 While previous models of CDI using clindamycin as the antibiotic to induce susceptibility to infection have used VPI10463 spores,39 we use UK1 spores (a hypervirulent, epidemic strain) to induce the C. difficile challenge at day 0 following antibiotic treatment until recovery at day 7 (Figure 3A), due to its higher germination levels after dysbiosis (Figure 2C vs 2B). The in vivo CDI metrics of the mouse model over 7 days show disease onset based on % weight change (Figure 3B), clinical score (Figure 3C), C. difficile toxin levels by ELISA (Figure 3D), and C. difficile shedding in stool by qPCR (Figure 3E). Interestingly, symptomatic infection was only apparent with the UK1 spore infected mice, with significant weight loss by day 2, followed by gradual recovery (Figure 3B) and differences in clinical scoring onward from day 2 (Figure 3C). For the UK1-infected mice, C. difficile toxins A/B in stool measured by toxin ELISA (Figure 3D) show high toxin levels on day 1 post-infection and continue to be high on day 3, with some drop-off on day 7 post-infection. C. difficile shedding in stool tracks with the toxin (Figure 3E). After challenge with the VPI10463 vegetative and spore-infected groups (Figure S2A), the respective differences in weight loss (Figure S2B) and clinical score (Figure S2C) were not apparent, while toxin secretion (Figure S2D) and C. difficile shedding (Figure S2E) were only apparent for the VPI10463 spore-infected groups. These results highlight the ability of C. difficile UK1 spores to colonize and cause disease (CDI) following clindamycin-induced dysbiosis, making it a suitable model for the ex vivo germination assay.

Modulating CDI Susceptibility by Promoting Dysbiosis

To increase host susceptibility to CDI, the clindamycin dose (Figure S3) and duration (Figure 4) prior to the C. difficile challenge were altered.

For this purpose, the mouse model was pre-treated with clindamycin via oral gavage (Figure 4A), allowed a 2-day break to get any residual clindamycin out of their system due to possible interferences with colonization, and then subjected to the C. difficile challenge (UK1 spores). For the pre- and post-antibiotics groups of mice, stool was collected prior to C. difficile infection to validate the ability of the spore germination assay to predict CDI susceptibility. Stool was collected regularly from these mice groups after the C. difficile challenge to follow C. difficile shedding and assess the ability of the ex vivo spore germination assay to follow recovery from antibiotic-induced dysbiosis. Since a clindamycin dose of 25 mg/kg causes substantial alterations in weight, clinical score, survival, and C. difficile shedding (Figure S3), we used this dose and varied its duration to 1, 5, or 10 days. With progressively higher clindamycin exposure times, the mice exhibited greater weight loss (Figure 4B), higher clinical scores (Figure 4C), and greater mortality (Figure 4D). This suggests that the dysbiosis, which was enhanced with clindamycin exposure time led to more severe CDI metrics. While the C. difficile shedding at day 1 (D1) after the challenge was similar for all the clindamycin receiving groups (1, 5, and 10 days of antibiotic exposure (or “abx”) in Figure 4E), the shedding drops off by day 5 (D5) for the 1-day antibiotic exposure group but remains steadily high for the 5- and 10-day antibiotic exposure groups. This suggests that the microbiota alterations that led to susceptibility to C. difficile colonization were more severe in the groups receiving the higher antibiotic durations vs those receiving higher antibiotic doses (Figure S3). Mice that do not receive clindamycin did not show detectable levels of C. difficile in their stool throughout the experiment, indicating that their microbiota was able to resist colonization.

Impedance-based distinction of Spore vs Vegetative Events

Using a homogeneous sample of C. difficile spores, we investigated their distinction vs a near-complete germinated sample of vegetative bacteria (1% taurocholate-treated). While spores differ in size vs vegetative bacteria, aggregation of spores in fecal supernatant led to inability for distinction based on impedance magnitude (Z10 MHz), but differences are apparent in impedance phase (ϕZ10 MHz) in Figure 5A. The higher ϕZ10 MHz level for vegetative bacteria vs spores (after normalization against co-flowing 2 μm polystyrene beads at ϕZ10 MHz = 0) is attributed to their higher interior conductivity that arises from the cytoplasm in vegetative bacteria, which is absent in spores. Using ϕZ10 MHz histograms of the respective samples (Figure 5B), the threshold for distinction of spores vs vegetative bacteria is set at ϕZ10 MHz= 218°, based on >95% of spore events on the lower side and vegetative on the higher side of this gate. Using this gate, we compare the impedance cytometry events from a set of spore samples (data pooled from triplicate runs) incubated with the fecal supernatant from untreated (Figure 5C) vs clindamycin-treated mice (Figure 5D). The respective ϕZ10 MHz histograms (Figure 5E) show a reduction in spore events and a rise in vegetative events for untreated (364 vegetative events) vs clindamycin-treated (462 vegetative events), for 3090 total events in both samples. Based on this, the ϕZ10 MHz threshold was set at >218° to count the germinated events from vegetative bacteria in each spore sample incubated with the fecal supernatant of various mouse models studied in this work.

Correlating Ex Vivo Spore Germination to In Vivo CDI

The germination of C. difficile spores in the ex vivo fecal milieu from the antibiotic-treated mouse model was studied to assess its correlation with antibiotic-induced dysbiosis conditions and its ability to predict susceptibility to CDI. Based on an experimental timeline analogous to Figure 4A, the correlation of spore germination to dysbiosis was performed using stool samples from the mouse model under pre-infection conditions (i.e., before C. difficile challenge). For the pre-infection group, stool was collected at day 0 (D0) from mice after antibiotic treatment over 0-, 1-, 5-, and 10-day durations. Following C. difficile challenge at day 0, samples for the post-infection group were collected on day 5. After incubation of UK1 spores in the sterile filtered fecal supernatant from these stool samples, the level of spore germination was quantified by impedance cytometry after 4 h, while the CFU assay of serially diluted spores was performed by streaking on BHI agar plate to count colonies after 24 h. For impedance cytometry, we found that quantifying germination as a normalized percentage of the total number of bacteria events was more accurate than the raw numbers of bacteria due to variations in the total number of measured events between samples for the same starting spore numbers. This variation is due to the small volumes used for spore inoculation (∼10 μL), the adsorption of spores and bacteria to plastic microcentrifuge tubes, and sample settling in the chip reservoirs during impedance cytometry.

Since impedance cytometry for counting bacterial cells was performed at the 4 h time point post-inoculation, which is well before the 16 h time point for onset of log phase bacterial growth (Figure 2), the determined % bacterial cells can be attributed solely to spore germination. On the other hand, bacterial cell quantification by other methods (e.g., absorbance in liquid media or CFU analysis) requires >16 h for sufficient detection sensitivity, thereby reducing its accuracy for attribution solely to spore germination, due to additional contributions from log phase bacterial growth. Interestingly, the CFU assay of the pre-infection group did not show significant differences between the groups receiving different durations of the antibiotic (0–10 day abx in Figure 6A). On the other hand, based on the count of % bacteria in the corresponding impedance phase gate (>218°), spore germination was significantly higher in ex vivo samples from the 5- and 10-day antibiotic-treated mice vs those not receiving antibiotics (Figure 6B). Following the C. difficile challenge, these 5- and 10-day antibiotic-treated mice had shown more severe in vivo metrics of CDI (Figure 4B–D) and C. difficile burden based on fecal shedding (Figure 4E) than observed with the 1-day antibiotic-treated and with the untreated mice. This highlights the ability of the ex vivo spore germination assay using impedance cytometry to assess dysbiosis in the antibiotic-treated mouse model for predicting its susceptibility to CDI. Considering the post-infection groups, spore germination as determined by the CFU assay (Figure 6C) and impedance cytometry (Figure 6D) was significantly higher for the 10-day antibiotic-treated mice vs other treatment groups. The germination levels were closer for the 5- and 1-day antibiotic-treated mice and for untreated mice. Hence, antibiotic-induced dysbiosis in the pre- and post-infection groups assessed by the ex vivo germination assay correlates to in vivo CDI metrics (Figure 4).

Spore Germination for Monitoring Persistence of Dysbiosis

CDI has been shown to cause persistent dysbiosis in mice through the induction of intestinal inflammation.40 We studied the ability of the ex vivo spore germination assay in fecal supernatant from mouse models to monitor the persistence of dysbiosis, which may serve as a potential biomarker for CDI recurrence following treatment. Persistent dysbiosis could leave the host vulnerable to subsequent infections through ingestion of new spores or germination of leftover spores in the gut microenvironment.41

Per the timeline in Figure 7A, mice were treated with clindamycin for 1- and 5-day durations and then given a day to recover from the antibiotic treatment, and their stool was collected for the spore germination assay. In Figure 7B, we set the baseline level of % bacteria from the spore germination assay based on the group of mice that received no clindamycin. The mice receiving clindamycin for 5 days had significantly greater spore germination levels above this baseline level for days 1–3 after cessation of antibiotic treatment, but reached the baseline level by day 4. On the other hand, mice receiving clindamycin for 1 day had significantly greater spore germination above this baseline level, only on day 1 after cessation of antibiotic treatment, with a return to baseline levels on subsequent days. This suggests that functional recovery of the microbiota after dysbiosis to resist colonization to C. difficile can take longer at the higher durations of the initial exposure to clindamycin. The recovery of microbiota for the 5- and 1-day clindamycin-treated mice is also apparent based on the in vivo CDI metrics after C. difficile infection of the mice, which show a return to baseline levels after 7 days of antibiotics (Figure 4B,C).

Conclusions and Outlook

Antibiotic-induced intestinal dysbiosis is widely associated with susceptibility of the host to C. difficile infection (CDI), and persistence of dysbiosis is associated with recurrence of CDI. However, given the wide person-to-person differences in host microbiota, there is a need for probes that can assess the dynamics of dysbiosis and its recovery in individuals, as well as its influence on susceptibility of the host to cause initial or recurrent CDI. Since the germination of C. difficile spores is a virulence trait for the onset of CDI, we utilize spore germination within sterile-filtered fecal supernatant samples from the host after antibiotic treatment as a rapid means to probe dysbiosis and its effect on their susceptibility to CDI. This fecal supernatant after each antibiotic treatment condition represents the temporal profile of the intestinal milieu from the host. The degree of intestinal dysbiosis due to antibiotic treatment of the mouse model likely alters the metabolite milieu in the gut, as reflected in their stool sample, thereby promoting the germination of C. difficile spores to cause colonization and CDI. Specifically, we infer that challenge with UK1 C. difficile spores after antibiotic treatment causes effective infection in the mouse model. The UK1 C. difficile spores also exhibit high levels of ex vivo spore germination in the metabolite milieu of stool samples from the antibiotic-treated mouse model. Hence, this system can serve as an effective vehicle to systematically probe dysbiosis under antibiotic treatment to assess host susceptibility to CDI, as validated by in vivo outcomes. Also, spore germination in the ex vivo samples from the model can enable prediction of the dysbiosis conditions leading to CDI. Using clindamycin at a dose of 25 mg/kg/day prior to C. difficile challenge with UK1 spores, our results indicate that the in vivo CDI metrics show greater severity with increasing duration of the antibiotic exposure over the 1- to 10-day period (i.e., greater clostridial shedding, % weight loss, clinical score, and fatality). The resulting C. difficile burden based on shedding of the bacteria in stool was greatest for the 10- and 5-day antibiotic exposures, with shedding continuing through until day 10 after the C. difficile challenge. For prediction of CDI susceptibility after antibiotic-induced dysbiosis based on the ex vivo spore germination assay prior to the C. difficile challenge (i.e., pre-infection group), single-cell impedance cytometry is shown to be essential for high sensitivity quantification of % bacteria in the co-cultured sample. This method was able to determine significantly higher germination levels for mice receiving 10- and 5-day antibiotic exposure vs those receiving 1-day and no antibiotic exposure. In this manner, the susceptibility of antibiotic-treated mice (after 10- and 5-day exposure) to C. difficile infection after the C. difficile challenge was correlated to the spore germination in the respective ex vivo stool samples obtained from the pre-infected mice. This suggests that spore germination in fecal supernatant from an antibiotic-treated mouse model can detect intestinal dysbiosis and susceptibility to CDI. Similar distinction was not possible in the pre-infection group based on the CFU assay for spore germination, due to its high variability in quantifying small numbers of vegetative cells, which arises from its need for serial dilutions and from spatial extent limitations on the plate to count the colonies. On the other hand, microfluidic impedance cytometry is designed to detect single vegetative cells within small volumes (down to 32 vegetative cells in 50 μL samples29). Furthermore, we infer that persistence of dysbiosis in the mouse model due to clindamycin treatment at a dose of 25 mg/kg/day can be quantified based on spore germination using impedance cytometry. For exposure at this dose over a 5-day duration, the dysbiosis persists up to day 3 following the treatment. For exposure over a 1-day duration, the model recovers from dysbiosis by day 2 following the treatment. Recovery from dysbiosis after antibiotic treatment was validated by showing its ability to resist CDI. Microfluidic detection of single bacterial cells29,32,42−46 and on-chip culture systems47,48 have emerged in several recent works. Specifically, we show that since impedance cytometry is designed to quantify limited numbers of vegetative bacteria within small sample volumes, it is well suited for high-sensitivity quantification of spore germination. This will allow us to probe germination at earlier time points and with smaller starting spore levels. The minimal sample preparation needs are significant for point-of-care translation, while low detection limits are important, since the onset of CDI can occur after exposure to just 100 spores. Future work will consider antibiotics with a wider spectrum of activity, dose, and duration ranges. We will also couple GC-MS based quantification of proportions of secondary to primary bile acids49 within the ex vivo stool samples from antibiotic-treated mice for directly correlating C. difficile spore germination to the composition of the metabolite milieu. For clinical translation, further work is also needed to carry out the assay in human fecal specimens to identify patients with an increased risk of CDI and to correlate the degree of spore germination with disease severity and recurrence.

Methods

Bacterial Strains and Spore Isolation

Clostridioides difficile strain UK1 (ribotype 027) and VPI10463 were used. Vegetative cells were generated by inoculating 10 mL of liquid BHIS media (brain heart infusion supplemented; Thermo Fisher Scientific, MA, USA). This was followed by growth for 24 h in an anaerobic chamber (BACTRON, Sheldon Manufacturing, OR, USA). To generate an enriched spore sample, vegetative C. difficile cultures were streaked on BHIS plates agar plates and allowed to grow for 7 days. The agar plates were washed with sterile ice cold water to enumerate spores, and the spores were centrifuged at 15000g for 15 min, followed by resuspension in 500 μL of 20% Histodenz (Sigma-Aldrich, USA). This was followed by addition of 500 μL of 50% Histodenz to purify spores through gradient centrifugation at 15000g for 15 min. Spores were centrifuged and washed 3 times with sterile DI water to remove residual Histodenz. Spores were then heated to 70 °C for 20 min to kill any remaining vegetative cells and stored for up to 6 months at 4 °C. To enumerate spores by the CFU assay, 10 μL of culture was streaked on BHIS agar plates supplemented with 0.1% taurocholate to promote germination. Bacteria were grown on these plates for 7 days before washing with ice-cold sterile DI water to count colonies.

Animal Work

Animal experiments were performed under a protocol approved by the Animal Institutional Care and Use Committee at the Center for Comparative Medicine at the University of Virginia, in line with national standards. C57BL/6J male mice, 8 weeks of age (Jackson Laboratories, Farmington, CT, USA) were either untreated or given clindamycin at varying doses and durations—either 5, 25, and 50 mg/kg/day for 5 days or 25 mg/kg/day for 1, 5, or 10 days—via oral gavage (n = 7 animals per group). An initial experiment was also performed using an antibiotic cocktail to investigate whether measuring spore germination in stool was feasible, and stool was collected before and after antibiotic administration in drinking water containing vancomycin (0.0045 mg/g body weight), colistin (0.0042 mg/g), gentamicin (0.0035 mg/g), and metronidazole (0.0215 mg/g) for 3 days. Mice were also administered a single dose of clindamycin (0.032 mg/g) via intraperitoneal injection. Mice were then challenged with C. difficile strain UK1 spores, VPI10463 spores, or VPI10463 vegetative cells following antibiotic administration. Mice were singly housed after infection to prevent cross-contamination and re-infection. Mice were monitored for up to 10 days post-infection and checked daily for weight and disease severity scores. Disease severity scoring was based on a 0–20 point scale,38 which takes into account weight change from day 0 (0–4 points), posture (0–3), coat appearance (0–3), diarrhea (0–3), eye appearance, squint and discharge (0–3), and other activity (0–4). Moribund animals, i.e., animals that had lost more than 25% of their baseline body weight post-infection and animals that had a combined clinical score of 14 or higher, were euthanized according to protocol.

Sample Preparation

Mouse stool collected from each treatment group was pooled, homogenized, and diluted 1/10 w/v in PBS. Stools were weighed, combined with PBS in a 1:10 weight/volume ratio, and centrifuged to pellet solids, and the supernatant was sterile filtered using a 0.2 μm syringe filter (EMD Millipore, Darmstadt, Germany). 10 μL of UK1 or VPI10463 spores (∼106 spores) were incubated in fecal supernatant for 24 h then diluted 1/10 for use in impedance cytometry or CFU assays.

Colony Forming Units (CFU) and Growth Assay

After incubation in fecal supernatant for 24 h, the spores were serially diluted and then streaked on BHI agar plates. Colonies were then counted after 24 h. For select experiments, spores were placed in liquid BHIS and allowed to grow for 24 h, with growth being measured (via optical density at 589 nm) every 20 min by a Cerillo Optoreader (Cerillo Inc., Charlottesville, VA, USA).

Impedance Cytometry for Spore Germination

Spores incubated in fecal supernatant were diluted in a 1:1 ratio in BHI media and allowed to germinate in growth media for 4 h prior to loading into a microfluidic chip (Figure 1B) using a syringe pump (Pump 11 Pico Plus Elite, Harvard Apparatus, Holliston, MA, USA) for pulse-free flow at 10 μL/min. The chip was fabricated with channels of 30 μm depth by 30 μm width and two sets of top-bottom facing electrodes (10 μm width and spacing) for multifrequency impedance analysis using the Amphasys Z32 impedance analyzer (Amphasys AG, Switzerland). Impedance cytometry measurements were performed at 500 kHz, 2 MHz, and 10 MHz in 0.5× PBS media (Thermo Fisher Scientific, MA, USA) of 0.8 S/m conductivity to optimize frequency for distinction of spore aggregates vs vegetative cells. Following this, we used only the optimized frequency for quantification of vegetative bacterial cells in fecal supernatant samples, to reduce event rejection rate based on bipolar Gaussian signal shape fit, so that we could maintain sufficient event numbers for statistically significant distinctions to enable detection of small numbers of vegetative bacteria at the earliest time points. Each cytometry run was performed in ∼2 min using ∼50 μL of the sample to quantify spore germination based on % of events in the “live” C. difficile gate vs the total number of events (Figure 1C and Figure 5). The events outside this “live” gate represent spore aggregates and dead bacteria, arising from spore preparation. All runs were performed in triplicate and pooled together in some cases.

Statistical Analysis

All statistical analysis was performed using Prism version 8 (GraphPad Software, San Diego, CA). Significance was defined as p < 0.05 in all cases. Comparisons between two groups were done using a Student’s two-tailed t test, while comparisons between multiple groups were performed using one-way ANOVA with a Tukey’s multiple comparisons post-test or two-way ANOVA with Dunnett’s multiple comparisons post-test where appropriate.

Quantitative Real-Time PCR

Genomic DNA was extracted from stool using a QIAamp DNA Stool mini kit (Qiagen, Redwood City, CA, USA) according to the manufacturer’s protocol. Quantitative real-time PCR experiments were performed using the CFX96TM Real-Time PCR Detection System (Bio-Rad Laboratories, Inc., Hercules, CA, USA). A total reaction volume of 20 μL per sample was prepared by mixing 10 μL of Kapa SYBR Fast quantitative PCR Master Mix (Kapa Biosystems, Wilmington, MA, USA), 1 μL of each forward and reverse primers (Eurofins MWG Operon, Huntsville, AL, USA) at 10 μM (final concentration 0.4 μM), 4 μL of template DNA, and 4 μL of DEPC-treated nuclease-free sterile water (Fisher Scientific, Pittsburgh, PA, USA). tcdB forward primer (5′-GGTATTACCTAATGCTCCAAATAG-3′) and tcdB reverse primer (5′-TTTGTGCCATCATTTTCTAAGC-3′) were used to quantify C. difficile in stool.

Supporting Information Available

The Supporting Information is available free of charge at https://pubs.acs.org/doi/10.1021/acsinfecdis.3c00192.

  • Gating live vegetative C. difficile using impedance cytometry data (Figure S1) and effect of clindamycin dose on in vivo CDI metrics (Figures S2 and S3) (PDF)

Author Contributions

John H. Moore: Conceptualization, investigation, formal analysis. Armita Salahi: Investigation, software, formal analysis. Carlos Honrado: Investigation, software, formal analysis. Christopher Warburton: Investigation. Steven Tate: Investigation. Cirle A. Warren: Supervision, conceptualization, resources, funding acquisition. Nathan S. Swami: Project administration, supervision, conceptualization, resources, funding acquisition.

Funding for the work was from NIH Grant 1R21AI130902-01, U.S. AFOSR contract FA2386-21-1-4070, and University of Virginia’s Global Institute for Infectious Diseases (GIDI). C.W. is partially supported by NIH Grant R01AI145322.

The authors declare no competing financial interest.

Supplementary Material

id3c00192_si_001.pdf (725.8KB, pdf)

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