Eleven C. difficile strains were utilized in this study. Strains R20291 (ribotype 027, tcdA⁺, tcdB⁺, tcdC⁺, cdtA^-, ctdB⁺) and 630 (ribotypes 012, tcdA⁺, tcdB⁺, tcdC⁺, ctdA^-, ctdB^-) used in this study are described elsewhere (McEllistrem et al., 2005; Buckley et al., 2011). The other nine isolates, which consisted of three tcdA+tcdB+ isolates (TNHP 29, 59, and 207), three tcdA-tcdB+ isolates (TNHP 79, 82, 403), and three tcdA-tcdB- isolates (1, 3, and 6), were originally isolated from a hospital in Southern Taiwan and have been described by Hung et al. (2016). All C. difficile strains were cultured anaerobically on brain–heart infusion (BHI) agar or in BHI broth (Thermo Fisher Scientific, Waltham, MA, United States) supplemented with 0.05% L-cysteine. Anaerobic experiments were conducted inside a Don Whitley DG250 anaerobic workstation (Don Whitley Scientific Ltd., West Yorkshire, United Kingdom).

Fatty acids (propionic acid, butyric acid, isobutyric acid, valeric acid, isovaleric acid, hexanoic acid, octanoic acid, capric acid, lauric acid, myristic acid, and palmitic acid) were purchased from Sigma–Aldrich (St. Louis, MO, United States). A minimum inhibitory concentration (MIC) assay was conducted according to the guidelines of the Clinical Laboratory and Standards Institute (formerly the National Committee for Clinical Laboratory Standards) for anaerobes. An overnight-grown culture was refreshed 50-fold in fresh brain–heart infusion-supplemented (BHIS) broth, incubated until the optical density at 600 nm (OD₆₀₀) was approximately 0.35, and then diluted 8-fold in 96-well microplates. FAs predissolved in dimethyl sulfoxide (DMSO) were then added to the bacterial suspension to reach a 5% DMSO final concentration. Concentrations of FAs ranged from 0.01 to 5 mg/mL. Bacterial cells were also incubated in BHIS + 1% DMSO only as a control. Plates were incubated anaerobically at 37°C for 24 h. To determine the MBC, bacterial suspensions from each well were streaked out onto BHIS agar plates and incubated for an additional 24 h. The MBC of each FA was defined as the lowest concentration at which no visible colony was observed. To determine the inhibitory effects of different concentrations of lauric acid on the C. difficile strain R20291, the percentage of growth at each concentration was calculated using the following equation: inhibition (%) = [1 - (OD₆₀₀ of growth with lauric acid/OD₆₀₀ of growth in broth only) × 100]. The growth inhibition rate was plotted against the log of the lauric acid concentration, and the IC₅₀ value was defined as the value that caused a 50% reduction in bacterial growth. At least three independent samples were analyzed for each experiment.

To determine the antibacterial activity of lauric acid on C. difficile growth, we employed different concentrations of lauric acid (1×, 4×, and 8× MBC) in the growth assay conducted in an anaerobic chamber. In the assay, 1% DMSO served as a treatment control. The C. difficile strain R20291 was cultured in BHIS broth at 37°C in the anaerobic chamber for 16 h, and the overnight cultures were then refreshed 50-fold in fresh BHIS broth until the late exponential to early stationary phase (OD₆₀₀ = approximately 0.8). Next, bacterial suspensions were added to DMSO or lauric acid, and OD595 was determined using a Libra S2 Colorimeter (Biochrom, Cambridge, United Kingdom). At least three independent samples were analyzed for each experiment.

To determine the effect of lauric acid on biofilm formation, an overnight culture of the C. difficile strain R20291 was refreshed to the late exponential to early stationary phase (OD₆₀₀ = approximately 0.8) in BHIS broth and then diluted 100-fold in fresh medium (BHIS + 0.1 M glucose) in 24-well polystyrene plates. Lauric acid at concentrations ranging from 0.125× to 1× MBC was added, and plates were incubated anaerobically at 37°C for 72 h. To determine the effect of lauric acid on preformed biofilms, biofilms were prepared for 24 h before treatment with lauric acid at concentrations ranging from 1× to 4× MBC for another 24 h. Moreover, 20 μg/mL vancomycin [40× the MBC of strain R20291, (Dapa et al., 2013)] and 1% DMSO were used as controls. To quantify the biofilm mass, supernatants were carefully decanted, and biofilms that formed in all wells were allowed to dry at room temperature. Two percent crystal violet (CV) was added to each well for 30 min and then removed through methanol treatment for an additional 30 min. Extracted dye contents were quantified by measuring the absorbance at 595 nm by using a Multiskan^TM GO Microplate Spectrophotometer (Thermo Fisher Scientific). Experiments were performed at least three times.

Spores were prepared by plating a 1:100 dilution of overnight culture onto BHIS agar plates and then incubating the plates for 10 days at 37°C under anaerobic conditions. Spores were harvested with ice-cold sterile distilled water and purified with 50% Nicodenz (Axis Shield, Oslo, Norway) using a previously described method (Sorg and Sonenshein, 2008). Spores were purified to >99% purity as determined using phase contrast microscopy, and the number of spores per milliliter was quantified through visual enumeration using a Neubauer Chamber (Sigma–Aldrich) prior to use.

To monitor germination efficiency, purified spores were first heat activated by incubating them for 30 min at 60°C and were then adjusted to an OD₆₀₀ of 1.0 in sterile water; 75-μL aliquots of spore suspensions were mixed with equal volumes of lauric acid or sterile water supplemented with 5% DMSO for 20 min at room temperature. Next, 10 mM taurocholic acid (Fisher Scientific) was added, and OD₆₀₀ was measured at 2-min intervals (Multiskan^TM GO Microplate Spectrophotometer, Thermo Fisher Scientific). The ratio of the OD₆₀₀ at time X to the OD₆₀₀ at time zero (t0) was plotted against time. The level of dipicolinic acid (DPA) release was monitored in real-time through terbium fluorescence, as described in previous studies (Bhattacharjee et al., 2015; Francis et al., 2015). Briefly, 75 μL of purified spores previously adjusted to the OD₆₀₀ of 1.0 were resuspended in germination buffer (10 mM Tris-Cl, 150 mM NaCl, 100 mM glycine, pH 7.5) and were then treated with equal volumes of various concentrations of lauric acid for 20 min. Next, 10 mM taurocholic acid was added, and DPA release was monitored using a FlexStation^® 3 Multi-Mode Microplate Reader (Molecular Devices, Sunnyvale, CA, United States) at an excitation wavelength of 270 nm and an emission wavelength of 545 nm. Spores boiled at 100°C for 30 min served as a positive control for total DPA release. Statistical analysis was performed using GraphPad Prism version 6.0. Three independent experiments were conducted. DPA release was calculated using the following equation:

To measure spore outgrowth, from the aforementioned germination assay, a 100-μL aliquot of spores from each reaction was serially diluted (10^-1–10^-7) with sterile phosphate-buffered saline (PBS) and then spread onto BHIS agar plates supplemented with 0.l% sodium taurocholate (TA) and incubated anaerobically at 37°C overnight. Spore outgrowth was calculated using the following equation:

An overnight culture of C. difficile R20291 was diluted at 1:50 in fresh BHIS broth and then grown to an OD₆₀₀ of approximately 0.8. Bacterial cells were then treated with various concentrations of lauric acid (1×–8× MBC) anaerobically at 37°C. Moreover, 1% DMSO in PBS served as the negative control, and 100 μg/mL nisin (Sigma–Aldrich) served as the positive control. At various time points, supernatants were collected, and the absorbances at 260 nm were recorded using a BioPhotometer UV/Visible Spectrophotometer (Eppendorf, Hamburger, Germany). Three independent experiments were performed.

To measure cell viability, an overnight culture of C. difficile R20291 was diluted at 1:50 in fresh BHIS broth and then grown to an OD₆₀₀ of approximately 0.8. Bacterial cells were then treated with lauric acid at 0.25× MBC or 1% DMSO in PBS for 20 min. Bacterial pellets were collected and resuspended in sterile PBS. Suspensions were then mixed with the LIVE/DEAD BacLight staining reagent mixture (Molecular Probes, Invitrogen) according to the manufacturer’s instructions. Samples were visualized under a FluoView^TM FV1000 confocal microscope (Olympus), and fluorescence was detected at an excitation wavelength of 488 nm and an emission wavelength of 500 nm (SYTO9) and 635 nm (propidium iodide). Cell viability is expressed as the ratio of SYTO-9-stained cells to the total number of cells. At least three independent trials were performed for each experiment.

Transmission electron microscopy (TEM) was employed to visualize the cells damaged by lauric acid treatment. C. difficile R20291 cells grown to the exponential phase were concentrated through centrifugation and treated with 0.25× MBC of lauric acid anaerobically for 15 min. Samples were embedded in Embed-812 (Electron Microscopy Sciences) and cut with an EM UC6 ultramicrotome (Leica, Wetzlar, Germany). Sections with a thickness of 90 nm were placed on copper grids (Electron Microscopy Sciences) and then stained with 2% uranyl acetate and lead citrate. Ultrathin sections were examined under a JEM-1400 transmission electron microscope (JEOL) with 120 kV acceleration and a 4k × 4k CCD Camera System Model 895 (Gatan, Inc.). The results are representative of three independent experiments.

Reactive oxygen species was measured using the carboxy derivative of fluorescein, CM-H2DCFDA (Life Technologies), according to the protocol provided by the manufacturer with the following modification: briefly, an overnight-grown C. difficile culture was refreshed to OD₆₀₀ of approximately 0.8 in BHIS broth. Moreover, 198 μL of the bacterial suspension was incubated with 2 μL of stock CM-H2DCFDA anaerobically at 37°C for 30 min. Cells were then treated with 1× MBC of lauric acid for 10 min, and fluorescence was then measured using the FlexStation^® 3 Multi-Mode Microplate Reader (Molecular Devices) at an excitation wavelength of 488 nm and an emission wavelength of 535 nm. The following controls were included: bacterial suspensions in BHIS broth containing 1% DMSO as the negative control; bacterial suspensions in BHIS broth containing 0.0035% hydrogen peroxide solution (H₂O₂; Sigma–Aldrich) and 10 mM tert-butyl hydroperoxide solution (TBHP; Sigma–Aldrich) as the positive control. The results are representative of three independent experiments.

An overnight culture of C. difficile R20291 was refreshed in Tryptone Yeast or BHIS broth and was grown anaerobically at 37°C until OD₆₀₀ was approximately 0.8. Bacterial cells were treated with various concentrations of lauric acid or 1% DMSO (control group) for 30 min anaerobically at 37°C. Cells were harvested through centrifugation, and total RNA was isolated using the RNAprotect Bacteria Reagent (QIAGEN, Venlo, Netherlands) in accordance with the manufacturer’s instructions. Genomic DNA was removed using RQ1 RNase-free DNase (Promega). RNA was reverse transcribed into complementary DNA (cDNA) by using SuperScript^TM II Reverse Transcriptase (Invitrogen) and random primers (Thermo Fisher Scientific) according to the manufacturer’s instructions. The relative transcriptional level of putative ROS-related genes between the control group and the lauric acid treatment group was measured through real-time quantitative reverse transcription polymerase chain reaction (qRT-PCR) using the 2x qPCRBIO SyGreen Mix Hi-Rox (PCR Biosystems) and gene specific primers (Table 1), according to the manufacturer’s instructions. A StepOnePlus^TM Real-Time PCR System (Applied Biosystems) was used. The data were analyzed using the 2^-ΔΔCt method, with normalization to the reference gene 16s and the stated reference condition. Samples were analyzed in at least three independent trials. Statistical analyses were performed using GraphPad Prism 6.0.

Specific-pathogen-free 8-week-old male C57BL/6 mice were housed in the Laboratory Animal Center of (NCKU). All mice were maintained and handled according to the guidelines of the Institutional Animal Care and Use Committee (IACUC) of NCKU. All animal studies were performed following the protocol approved by the IACUC of NCKU (approval NCKU-IACUC-102-149) and the Biosafety and Radiation Safety Management Division of NCKU. The animal model of CDI was established as previously described (Chen et al., 2008; Liu et al., 2017; Pizarro-Guajardo et al., 2017). Five animals were administered 12 mg/kg (low dose) and 24 mg/kg (high dose) of lauric acid dissolved in PBS orogastrically once per day for 7 days prior to infection with C. difficile and once more 1 day following infection. To condition the animals for CDI, mice were fed drinking water containing an antibiotic mixture, which included 0.4 mg/mL vancomycin, 0.215 mg/mL metronidazole, 0.4 mg/mL kanamycin, 0.035 mg/mL gentamicin, and 850 U/mL colistin, for a total of 5 days before the challenge. All antibiotics were purchased from Sigma–Aldrich. On the day before the challenge, mice were fed the antibiotic mixture without vancomycin and metronidazole, which were excluded to avoid disrupting C. difficile colonization. Esomeprazole dissolved in PBS was given to all mice through oral gavage 12 h prior to infection (18.55 mg/kg) and immediately before infection (4.82 mg/kg). On the day of infection, 1 × 10⁶ CFU of C. difficile R20291 spores were administered through oral gavage, and 4 mg/kg of clindamycin was injected intraperitoneally. Two days after infection, all animals were euthanized through CO₂ asphyxiation. The severity of diarrhea was scored in accordance with mice stool consistency, as follows: (0) well-formed pellets; (1) semiformed stools that did not adhere to the anus; (2) semiformed stools that adhered to the anus; and (3) liquid stools. Organs and gastrointestinal lavage (GAL) fluids were extracted for downstream analysis. The entire animal experiment was performed for a total of three independent sets, and representative results were obtained.

Fecal samples (premixed in PBS) were collected from the animals, immediately heat treated at 65°C for 20 min, and then serially diluted onto BHI agar containing 0.1% TA. Plates were incubated anaerobically at 37°C for 48 h, and colonies were counted for CFU determination.

Colon samples were extracted using RNeasy^® Plus Mini kits (QIAGEN). RNA yield and quality were examined using a NanoDrop Spectrophotometer (Thermo Fisher Scientific). Reverse transcription was performed with SuperScript^TM II Reverse Transcriptase (Invitrogen, Waltham, MA, United States). The expression levels of proinflammatory cytokines and chemokines were measured through qRT-PCR using RealQ Plus 2X Master Mix Green (Ampliqon, Denmark), with β-actin as the reference gene in each reaction (Table 1). The data were analyzed using the ΔΔCt method and expressed as the fold change in the transcription level under the test condition compared with the average for the indicated control and were then normalized to the reference gene β-actin. Statistical analyses were performed using GraphPad Prism 6.0.

The concentrations of GAL cytokines and chemokines were measured using a DuoSet^® enzyme-linked immunosorbent assay (ELISA) development system (R&D Systems, Minneapolis, MN, United States) according to the manufacturer’s instructions. Absorbance was measured at 450 nm using an iMark^TM microplate reader (Bio-Rad, Hercules, CA, United States). Samples were measured in triplicate, and statistical analyses were performed using GraphPad Prism version 6.0.

All data are expressed as the mean ± standard deviations of at least three independent experiments. Statistical comparisons among the groups were made using Student’s t-test. Multiple intergroup comparisons were made using one-way analysis of variance (ANOVA), followed by a post hoc Tukey’s test with GraphPad Prism version 6.0. Statistical significance was set at P < 0.05.

To identify the free FAs with potent C. difficile inhibitory effects, FAs C3–C16 were coincubated with log-phase-grown C. difficile cells, and their MBCs were determined (Table 2). For the test strain R20291, butyric acid (MBC = 50 mg/mL) showed the least inhibitory effects of all FAs tested, whereas lauric acid (MBC = 0.3125 mg/mL) showed the most potent effects. The length of the carbon chain did not appear to be an influencing factor contributing to the antibacterial activity of the FAs, although remarkably MCFAs appeared to exhibit lower MBCs. Specifically, the MBCs of both capric acid (C10) (1.25 mg/mL) and lauric acid (C12) (0.3125 mg/mL) were the second lowest and the lowest of all FAs tested. Moreover, the resulting pH variation in BHIS medium did not appear to contribute to the antibacterial activity of the various FAs (Table 2). As the MBC of lauric acid was significantly lower (fourfold, P < 0.0001) than that of capric acid, lauric acid was chosen as a potential candidate for subsequent experiments.

To further confirm the inhibitory effect of lauric acid on C. difficile growth, 10 additional clinical isolates of different toxinotypes were subjected to the same experiment (Table 3 and Figure 1) (Hung et al., 2016). The MICs of all 11 tested isolates ranged from 0.08 to 0.16 mg/mL; however, the MBCs were all 0.31 mg/mL. As the MBCs were all the same, we assumed the inhibitory effect of lauric acid on C. difficile is likely not strain-dependent; hence, all subsequent experiments were performed using the laboratory strains 630 and R20291. To gain further insight into the inhibitory effect of lauric acid on C. difficile, the IC₅₀ was determined. As depicted in Figures 2A,B, the IC₅₀ of lauric acid against C. difficile strains R20291 and 630 was 12.48 and 33.67 μg/mL, respectively. The antibacterial activity of lauric acid against strain R20291 was further evaluated in liquid nutrient broth containing different concentrations of lauric acid (Figure 2C). When lauric acid was applied at 2× and 4× MBC, cell lysis appeared to occur immediately, as evidenced by the drop in OD. In short, lauric acid was revealed to display an inhibitory effect on multiple strains of C. difficile, and the inhibition mechanism is likely bactericidal.

To further understand the effect of lauric acid on C. difficile, we determined whether lauric acid treatment affects biofilm formation and stability. In this study, we measured the effect of lauric acid treatment on biofilm formation by R20291 and 630 strains, as these two strains were reported to exhibit different biofilm forming abilities (Dapa et al., 2013). Clinical C. difficile strains are known to form robust biofilms in vitro, and these biofilm-dwelling cells are more resistant to antibiotics and perhaps to even host defenses than planktonic cells are (Dapa and Unnikrishnan, 2013; Dapa et al., 2013; Crowther et al., 2014). A previous study reported that vancomycin applied at 20 μg/mL (100× MIC) can significantly reduce the survival of C. difficile biofilm cells (Dapa and Unnikrishnan, 2013). For the strain R20291, vancomycin applied at 100× MIC could reduce biofilm formation by 15.5-fold compared with the DMSO control (P < 0.0001). However, 0.25× MBC of lauric acid could significantly reduce biofilm formation by 24.9-fold compared with the control (P < 0.0001) (Figure 3A). Similarly, for the strain 630, although vancomycin treatment led to a 20.8-fold reduction in biofilm formation, 0.25× MBC of lauric acid led to a 47.2-fold reduction (Figure 3B). To determine whether lauric acid treatment also disrupts established biofilms, a static culture of C. difficile was grown in multi-well plates for 24 h before lauric acid addition. Interestingly, although vancomycin applied at 100× MIC did not reduce the biofilm mass, lauric acid at least 2× MBC and 1× MBC could reduce the mass of biofilms formed by strains R20291 and 630, respectively (Figures 3C,D). Reduction of the mass of biofilms formed by the strain R20291 required a higher concentration of lauric acid than that from strain 630.

In addition to biofilm formation, the spore-forming ability of C. difficile contributes to its transmission (Deakin et al., 2012). Spores are known to be resistant to multiple environmental stresses, including cold, heat, desiccation, antiseptics, and antibacterial products (Rodriguez-Palacios and Lejeune, 2011; Deng et al., 2015; Edwards et al., 2016), and they are therefore a critical component of the pathogenesis of CID. One of the key germinating signals for C. difficile spores is the presence of TA. In the presence of TA, spores will undergo core hydration, and this process can be visualized microscopically; previously dormant phase-dark spores will become phase-bright due to core hydration and the eventual degradation of the cortex peptidoglycan. To measure the effect of lauric acid on spore germination, purified C. difficile strain R20291 spores were treated with taurocholic acid and various concentrations of lauric acid. As revealed in Figure 4A, in the presence of TA, the OD₆₀₀ of the spore suspension decreased significantly within 20 min of exposure (PC group, P < 0.0001), whereas the DMSO control remained phase-dark (NC group). Interestingly, when various concentrations of lauric acid were added along with TA, the decrease in OD₆₀₀ was faster over time than that in the PC group, suggesting that core degradation occurred at a faster rate (P < 0.0001 for all concentrations compared with the PC group). We also performed the same assay in the absence of TA, and no significant decrease in OD₆₀₀ was observed for all concentrations of lauric acid tested throughout the 20 min of observation, suggesting that the effect was TA-dependent (Figure 4B). During spore germination, DPA release can be measured and is often used as a sign of spore germination. To further investigate the role of lauric acid in spore germination, we measured DPA released for 20 min (Figure 4C). As expected, DPA released from boiled spores was detected by its high fluorescence signals, which were higher than those of non-treated spores. Similarly, in the presence of TA, an increase in the fluorescent signal was detected, suggesting that DPA release was initiated at approximately 4 min and continued to increase until the end of the experiment. However, the addition of lauric acid, regardless of concentration, did not significantly alter DPA release over the course of the experiment (P > 0.9999). Furthermore, the addition of lauric acid alone did not induce any DPA release (data not shown). Finally, the viability of spores in the presence of lauric acid was measured (Figure 4D). Compared with spores exposed to TA only, lauric acid treatment in the presence of TA considerably decreased the rate of spore outgrowth in a dose-dependent manner. Spore outgrowth decreased to 64.5, 61, 60.4, and 39.6% of typical growth when subjected to lauric acid concentrations of 0.5× MBC, 1× MBC, 2× MBC, and 4× MBC, respectively (Figure 4D). These combined results demonstrate that lauric acid treatment can inhibit C. difficile biofilm formation and stability and can affect spore outgrowth.

To determine whether the inhibitory effect of lauric acid on C. difficile growth is due to the disruption of cell membrane integrity, C. difficile cells were treated with lauric acid, and the extracellular presence of released nucleic acid was measured (Figure 5A). In this assay, we included nisin, a broad-spectrum polycyclic antibacterial peptide produced by Lactococcus lactis, as a positive control, as it is known to attack bacterial cell membranes; this leads to cytoplasmic content release and cell lysis eventually (Ruhr and Sahl, 1985; Nobmann et al., 2010). As expected, in the presence of the antibacterial peptide nisin, a significant quantity of nucleic acids was detected in the culture supernatant at 30 min after treatment. When cells were treated with various concentrations of lauric acid, a considerably higher quantity of nucleic acid materials was also detected compared with the negative control, indicating that the addition of lauric acid compromised cell membrane integrity. The viability of C. difficile cells treated with lauric acid was further assessed using LIVE/DEAD staining and was visualized using confocal microscopy (Figure 5B). As lauric acid induced the rapid lysis of C. difficile cells, as indicated in Figure 2C, to elucidate the effects of lauric acid on C. difficile cells, these cells were treated with sublethal concentrations of lauric acid (0.25× MBC). When cell viability was quantified based on the percentage of cells that stained positive for propidium iodide, treatment with lauric acid at 0.25× MBC decreased viability to approximately 65% by 15 min after treatment (Figure 5C). By 30 min after treatment, the viability decreased to approximately 35%. By contrast, the viability of the DMSO-treated control remained relatively high (15 min: 87%, 30 min: 84%). Ultrathin-section TEM analysis revealed that 20 min of lauric acid treatment (0.25× MBC) was sufficient to induce substantial cell death, as indicated by abnormal cell morphology and cytoplasmic content leakage. These findings are similar to those of Shilling et al. (2013) (Figure 5D and Supplementary Figure 1).

To determine whether lauric acid also induces ROS generation, vegetative C. difficile R20291 cells were treated with a sublethal concentration of lauric acid (0.25× and 0.5× MBC) for up to 60 min, and intracellular ROS levels were measured (see section “Materials and Methods”). As depicted in Figure 6A, treatment with the antiseptic hydrogen peroxide and tert-butyl hydroperoxide (TBHP) generated a considerable level of ROS in a time-dependent manner. Interestingly, lauric acid treatment also generated a substantial level of ROS over the course of the experiment. As no molecular or biochemical studies have been performed in C. difficile on ROS-associated genes, from the genomic annotation, we selected four potential antioxidant defense-associated genes. As depicted in Figure 6B, in the presence of 0.25× MBC of lauric acid, the expression levels of genes encoding for a putative superoxide dismutase (locus tag CDR20291_1529) and a putative catalase (locus tag CDR20291_1465) were upregulated (9.1- and 3.4-0 fold, respectively, compared with untreated cells) (Figure 6B). However, the gene expression levels of CDR20291_C0757, which encodes for a putative superoxide reductase, and CDR20291_1716, which encodes for a putative peroxidase, were not significantly altered in the presence of either 0.25× or 0.5× MBC of lauric acid. Nevertheless, the results of these experiments suggested that the bactericidal activity of lauric acid was partly due to the induction of membrane damage and ROS generation, which resulted in cell lysis.

Finally, to determine whether lauric acid affects CDI in vivo, C57BL/6 mice were administered lauric acid orogastrically for 1 week prior to infection with purified C. difficile R20291 spores (Supplementary Figure 2). Mice administered only PBS prior to infection displayed symptoms of CDI, including a lack of well-formed feces due to diarrhea and a considerable decrease in body and cecum weight (Figures 7A–D). Gross views of the colon and cecum indicated severe colitis (Figure 7E). By contrast, both groups of mice that were administered either 12 mg/kg (LA-low) or 24 mg/kg (LA-high) of lauric acid displayed healthier colon and cecum morphology (Figure 7E), body weight recovery after infection (Figure 7B), a substantially lower body weight decrease (Figure 7C), and a higher cecum weight (Figure 7D) than the PBS control group. The protective effects of lauric acid appeared to be dose-dependent, as mice belonging to the LA-high group appeared to exhibit less severe symptoms of CDI compared with mice in the LA-low group. No differences were observed in the number of heat-resistant spores recovered from fecal samples (Figure 7F).

The expression levels of genes encoding for proinflammatory cytokines such as tumor necrosis factor α (TNF-α), interleukin 6 (IL-6), interleukin 1β (IL-1βb), macrophage inflammatory protein 2 (MIP-2), and monocyte chemoattractant protein 1 (MCP-1) were considerably decreased in the colon homogenates of the LA-low and LA-high groups compared with those in the PBS group (Figure 8A). In addition to gene expression, proinflammatory cytokines and chemokines from the GAL fluid of all three groups were analyzed. Levels of TNF-α, IL-1β, and MCP-1 were significantly decreased in the GAL lavage fluid of both lauric acid treatment groups compared with that in the PBS control group (P = 0.0254, P = 0.036, and P = 0.0285, respectively, compared with the PBS control group) (Figure 8B). No statistically significant differences were observed in the levels of IL-6 and MIP-2 detected, although the trend was similar to that observed above. Collectively, these results demonstrated that the administration of lauric acid could decrease the severity of C. difficile-induced inflammation in vivo.