The Role of Morphine- and Fentanyl-Induced Impairment of Intestinal Epithelial Antibacterial Activity in Dysbiosis and its Impact on the Microbiota-Gut-Brain Axis

Abstract

Recent evidence suggests that chronic exposure to opioid analgesics such as morphine disrupt the intestinal epithelial layer and cause intestinal dysbiosis. Depleting gut bacteria can preclude the development of tolerance to opioid-induced antinociception, suggesting an important role of the gut-brain axis in mediating opioid effects. The mechanism underlying opioid-induced dysbiosis however remains unclear. Host-produced antimicrobial peptides (AMPs) are critical for the integrity of the intestinal epithelial barrier as they prevent the pathogenesis of the enteric microbiota. Here, we report that chronic morphine or fentanyl exposure reduces the antimicrobial activity in the ileum, resulting in changes in the composition of bacteria. Fecal samples from morphine-treated mice had increased levels of Akkermansia muciniphila with a shift in the abundance ratio of Firmicutes and Bacteroidetes. Fecal microbial transplant (FMT) from morphine-naïve mice or oral supplementation with butyrate restored a) the antimicrobial activity, b) the expression of the antimicrobial peptide,Reg3γ, c) prevented the increase in intestinal permeability and d) prevented the development of antinociceptive tolerance in morphine-dependent mice. Improved epithelial barrier function with FMT or butyrate prevented the enrichment of the mucin-degrading Akkermansia muciniphila in morphine-dependent mice. These data implicate impairment of the antimicrobial activity of the intestinal epithelium as a mechanism by which opioids disrupt the microbiota-gut-brain axis.

Graphical Abstract

1. Chronic opioid treatment disrupts intestinal epithelial barrier function by reducing antimicrobial peptides, thus causing an expansion of the mucin-degrading, Akkermansia muciniphila, “leaky” epithelium and inflammation. This contributes to the development of antinociceptive tolerance.

2. Oral administration of butyrate or FMT prevents opioid-induced epithelial barrier disruption by increasing antimicrobial activity, and reducing epithelial permeability and Akkermansia muciniphila. Improved epithelial barrier function results in a significant decrease in antinociceptive tolerance.

INTRODUCTION

It is well established that the chronic use of μ-opioid analgesics such as morphine results in the development of tolerance to the antinociceptive effects. Thus, a larger dose may be required for adequate pain control. Dose escalation can increase the propensity for unwanted effects such as addiction, constipation, and death due to overdose in severe cases since tolerance to opioid-induced respiratory depression does not develop at the same rate as tolerance to antinociception¹. Despite the known risks, opioids remain the gold standard for managing pain in the clinic. It is, therefore, important to 1) understand the mechanisms that form the basis for the development of tolerance and 2) apply this information to formulate novel strategies to confront the ongoing opioid crisis.

Traditionally, nausea, vomiting, and constipation are reported as common gastrointestinal (GI) effects of μ-opioid analgesics². Recent clinical evidence indicates that opioid use is associated with intestinal dysbiosis^3–7. While nutritional deficiencies, poor hygiene, and comorbidities can contribute to an altered microbiota in opioid users^8–10, the absence of a microbial shift in opioid users treated with opioid receptor antagonists, naloxone or naltrexone⁴, suggests that the dysbiosis may be an opioid receptor-dependent effect.

In addition to the burgeoning clinical evidence for opioid-induced dysbiosis, several preclinical studies have reported changes in the intestinal microbiome with chronic opioid exposure. Mice chronically treated with opioids exhibit altered intestinal microbiota, intestinal inflammation, perturbed intestinal epithelial barrier, and systemic translocation of luminal bacteria^11–18. The exact sequence of events is unclear. Several studies have demonstrated that the disruptive effects of opioids on the intestinal microbiota and the intestinal epithelium contribute to the development of tolerance to opioid-induced antinociception in vivo^11,12,16 and in primary afferent dorsal root ganglia neurons^11,12 as noted by the reversal of in vivo and cellular tolerance to morphine after antibiotic treatment. However, there is a significant gap in our current understanding of how opioids induce dysbiosis and its impact on antinociceptive tolerance.

The host and autochthonous microbiota maintain a commensal relationship through several mechanisms, including the secretion of antimicrobial proteins and peptides (AMPs) and IgA^19,20. AMPs are released primarily by specialized epithelial cells called Paneth cells that reside within small intestinal crypts²⁰. Enterocytes in the villi also secrete AMPs²¹. AMPs help create a chemical barrier against bacterial colonization of the intestinal epithelium and maintain the composition and diversity of the intestinal microbiota by inhibiting pathogenic strains. Indeed, several studies have demonstrated that depletion of Paneth cells, or disruption of mechanisms that regulate the synthesis or secretion of AMPs, enhances susceptibility to intestinal inflammation, promotes systemic translocation of luminal bacteria, and increases vulnerability to pathogenic infections²⁰. Since chronic treatment with morphine alters the intestinal microbiome and results in bacterial translocation, we sought to test the hypothesis that tolerance to opioid-induced antinociception results from disruption of the epithelial barrier function due to alteration in the antimicrobial activity of the intestinal epithelium.

Interestingly, fecal samples of opioid users were deficient in the short-chain fatty acid (SCFA), butyrate, and butyrate-producing bacteria^4,7. Anerobic bacteria in the intestines produce butyrate by fermenting dietary fiber and resistant starches²². Butyrate is utilized as a significant energy source by colonocytes; it improves epithelial barrier function by inducing the expression of tight junction proteins, increasing AMP production, and inhibiting pro-inflammatory cytokines²². We previously reported that exacerbating intestinal inflammation in an experimental model of colitis increased the rate at which tolerance developed to morphine’s antinociceptive effects¹⁸. Since butyrate is critical for maintaining the physiology of the intestinal mucosa and inhibits intestinal inflammation, in the present study, we also tested the hypothesis that improving the intestinal epithelial barrier with butyrate can prevent the development of tolerance to opioid-induced antinociception.

MATERIALS AND METHODS

Animals:

Male Swiss Webster mice (ENVIGO, Indiana, IN, USA) and male ICR mice (ENVIGO, Indiana, IN, USA) six-eight weeks old, weighing 25–30 g and 30–35 g, respectively, were housed five to a cage with ad libitum access to food and water in animal care quarters maintained under a 12-hour light/dark cycle (lights on from 7 am to 7 pm). Animals were randomly assigned to control and treatment groups. All animal procedures were in accordance with the protocols reviewed and approved by the Institutional Animal Care and Use Committee at Virginia Commonwealth University (VCU IACUC). Results of the animal experiments were reported in accordance with the recommendations of the ARRIVE 2.0 guidelines. Swiss Webster mice were used in morphine experiments and ICR mice were used in fentanyl experiments.

Test Drugs:

Morphine:

1. 75-mg morphine or placebo pellets, obtained from the National Institute on Drug Abuse (NIDA, Bethesda MD), were aseptically implanted in the subcutaneous cavity on the dorsum under isoflurane (2.5%) anesthesia as described previously¹¹. Mice were allowed to recover in their home cages. On test day 7, the mice were either subjected to antinociceptive response experiments (Figures 1 and 4B); used for fecal microbiome analysis (Figures 1, 2, 11, and S1–S4); used for harvesting blood, stool, and colon tissue for evaluating butyrate concentration (Figure 3); for measuring GI transit (Figure 5) and body weight loss (Figures 6A and 6C); for determining intestinal epithelial permeability (Figure 7A); in antibacterial activity experiments (Figures 8 and 10); or for evaluating Regenerating islet-derived 3 gamma (Reg3γ) gene expression in the ileum (Figure 9A) as described in the subsequent methods sections. The same animals were not used to generate different types of data unless indicated.

Figure 1. Effect of FMT on the development of tolerance to morphine-induced antinociception.

(A) Alpha diversity was assessed in the fecal samples of saline-treated PP and saline-treated MP mice using the Chao1 index. (B) Principal coordinate analysis (PCoA) of the beta diversity of the fecal bacteria in saline-treated PP and saline-treated MP mice was measured with the Bray-Curtis Index. (C) Schematic of FMT administration to PP and MP mice. Tolerance to the 10 mg/kg morphine challenge was assessed in pelleted mice using the (D) warm-water tail-withdrawal and (E) hot-plate tests. Mice were either sham-treated (i.e., no FMT) or administered PP-FMT or MP-FMT. Low %MPE values indicated the development of tolerance. Data in A was analyzed by two-tailed unpaired Student’s t-test. Data in D and E were analyzed by two-way ANOVA, and Tukey’s multiple comparisons test was used for post hoc analysis. P values for relevant interactions have been indicated. Data in A, D and E are mean ± SEM, and scatter in A, B, D, and E represents data from individual mice. Sample sizes in A and B are 5 mice per group. Sample size in D are: 5 (PP-Sham and MP-Sham) and 10 (PP + PP-FMT, MP + PP-FMT, PP + MP-FMT, and MP + MP-FMT) mice. Sample sizes in E are: 5 (PP-Sham and MP-Sham), 4 (PP + PP-FMT), 5 (MP + PP-FMT), and 4 (PP + MP-FMT and MP + MP-FMT) mice.

Figure 4. Effect of sodium butyrate on the development of tolerance to morphine-induced antinociception.

(A) Mice were treated with increasing doses of sodium butyrate and the dose dependent effect of butyrate on the development of tolerance to morphine-induced antinociception was evaluated in the warm-water tail-withdrawal test. These mice were intermittently treated with escalating saline or escalating doses of morphine saline to induce tolerance to morphine-induced antinociception. (B) The effect of sodium butyrate on the development of tolerance to morphine-induced antinociception was also evaluated in pelleted mice using the warm-water tail-withdrawal test. Data in A and B were analyzed by two-way ANOVA analysis and Tukey’s test was used for comparisons between groups. (C) Mice were treated with 0.3 mg/kg fentanyl i.p. b.i.d and orally administered saline or 0.250 M sodium butyrate. Tolerance to 10 mg/kg morphine-induced antinociception was tested using the warm-water tail-withdrawal test. Data were analyzed by one-way ANOVA analysis and Tukey’s test was used for multiple comparisons. P values for relevant comparisons have been indicated. Data are expressed as mean ± SEM and scatter represents data from individual mice. Samples sizes for A are 5/dose/group, sample sizes for B are 5/group, and sample sizes for C are 7 in the Saline + Butyrate group, 10 in the Fentanyl + Saline group, and 9 in the Fentanyl + Butyrate group

Figure 2. Fecal microbial analysis in FMT-treated mice.

(A) Chao1 diversity index in the fecal microbiota of FMT donor and recipient mice. (B) Principal coordinate analysis (PCoA) of the beta diversity of FMT-treated mice. (C) Relative abundance of Akkermansia muciniphila in FMT-treated mice. Data from PP + PP-FMT, PP + MP-FMT, MP + PP-FMT, and MP + MP-FMT mice in A and C were analyzed by two-way ANOVA, and Tukey’s multiple comparisons test was used for post hoc analysis. P values for relevant interactions in A and C have been indicated. Data in A and C are mean ± SEM, and scatter in A, B, and C represents data from individual mice. Sample sizes are 6 (PP + PP-FMT), 5 (MP + PP-FMT), 4 (PP + MP-FMT) and 4 (MP + MP-FMT) mice.

Figure 11. Bacterial composition in butyrate-treated mice.

(A) The ratio of the relative abundance of Firmicutes/Bacteroidetes showed that butyrate did not prevent the reduction in Firmicutes/Bacteroidetes ratio in chronic morphine-treated mice. The relative abundance used to calculate the Firmicutes/Bacteroidetes ratio in PP + Saline and MP + Saline mice are from Figure S1. (B) Relative abundance of Verrucomicrobia in butyrate-treated mice. (C) Relative abundance of Akkermansia muciniphila in butyrate-treated mice. Data in A, B and C were analyzed using two-way ANOVA, and Tukey’s multiple comparisons was used for post hoc analysis. Significant P values have been indicated. Data are mean ± SEM, and the scatter represents data from individual mice. Sample sizes are 5 mice per group

Figure 3. Concentration of endogenous butyrate in stool, colon tissue and blood.

The concentration of endogenous butyrate was determined in (A) stool, (B) colon, and (C) blood samples from pelleted mice using GC-MS. Chronic exposure to morphine reduced endogenous butyrate in stool but not in colon or blood. Oral administration of 0.250 M sodium butyrate replenished butyrate levels in the stool samples of morphine-treated mice. Data in A were analyzed using two-way ANOVA with Tukey’s multiple comparisons post hoc test. Data in B and C were analyzed using unpaired two-tailed Student’s t-test. P values for comparisons between groups have been indicated. Data are mean ± SEM and scatter represents data from individual mice. Sample sizes for A are: 8 (PP, and MP), and 5 (PP + sodium butyrate, and MP + sodium butyrate). Sample sizes for B and C are: 5 (PP and MP).

Figure 5. Butyrate did not prevent chronic morphine-induced inhibition of GI motility.

The effect of 0.250 M sodium butyrate on the GI motility of chronic morphine-treated mice was determined on day 7 using the carmine red dye assay. Elevated GI transit times indicate reduced motility. Data are expressed as mean ± SEM and scatter represents data from individual animals. Data were analyzed by two-way ANOVA. The P value for the main effect of morphine treatment on GI transit time is indicated. Sample sizes are 5/group.

Figure 6. PP-FMT or butyrate did not attenuate chronic opioid-induced body weight loss.

Percent change in body weight was measured in mice treated with (A) FMT or (B, C and D) 0.250 M sodium butyrate. Body weight data in A, B, and C are of morphine-treated mice used in the warm-water tail-withdrawal assay in Figures 1D, 4A, and 4B, respectively. Body weight data in D are of fentanyl-treated mice used in the warm-water tail-withdrawal assay in Figure 4C. Data are expressed as mean ± SEM and scatter represents data from individual animals. Data in A, B, and C were analyzed using two-way ANOVA and data in D were analyzed using one-way ANOVA and Tukey’s post hoc test for multiple comparisons. P values for the main effect of morphine treatment on percent body weight change have been indicated in A-C, and P values for the post hoc analysis of fentanyl treatment on percent body weight change have been indicated in D. Sample sizes in A are 10/group, B and C are 5/group, and D are 7 in the Saline + Butyrate group, 10 in the Fentanyl + Saline group, and 9 in the Fentanyl + Butyrate group.

Figure 7. Effect of FMT or butyrate on morphine-induced disruption of epithelial permeability.

FITC-dextran was fluorometrically quantified from whole blood samples in morphine-treated mice after (A) FMT or (B) sodium butyrate administration. Data in A and B were analyzed by two-way ANOVA with Tukey’s multiple comparisons test. Relevant P values have been indicated. Data are expressed as mean ± SEM and scatter represents data from individual mice. Sample sizes in A are: 5 (PP-Sham, and MP-Sham) and 10 (PP + PP-FMT, and MP + PP-FMT). Sample sizes in B are 5/group.

Figure 8. Effect of morphine and sodium butyrate on the activity of the ileum against L.reuteri.

(A) Schematic of the antibacterial activity assay protocol. Each agar plate is divided into quadrants and in quadrant I ileum-derived conditioned media incubated with bacteria was smeared. In quadrant II ileum conditioned media containing only nutrient broth and no bacteria was applied; in quadrant III antibiotic-free DMEM/F12 media incubated with bacteria was smeared; and in quadrant IV antibiotic-containing DMEM/F12 media incubated with bacteria was applied. Conditioned media obtained from the ileum of (B) PP mice, (C) MP mice or (D) MP mice treated with 0.250 M sodium butyrate (MP + Butyrate) was used to evaluate the antibacterial activity of the ileum against the Gram-positive bacteria, L.reuteri. Representative images of L.reuteri growing on agar plates have been shown in B-D. Figure E is the normalized data of the %bactericidal activity of the ileum supernatants from PP, MP and MP + butyrate mice. Data in B-D were analyzed by repeated-measures one-way ANOVA and Tukey’s multiple comparisons test was used for post hoc analysis. Data in E was analyzed by one-way ANOVA and Holm-Sidak’s multiple comparisons test was used for post hoc analysis. P values for relevant comparisons have been indicated. All data are mean ± SEM and scatter represents data from samples from individual mice. Sample sizes are 6, 7, and 6 per group for B, C, and D, respectively. Sample sizes for E are: 6 (PP), 7 (MP) and 6 (MP + butyrate).

Figure 10. Effect of morphine and sodium butyrate on the activity of the ileum against E.coli.

Conditioned media obtained from the ileum of (A) PP mice, (B) PP mice treated with 0.250 M sodium butyrate (PP + butyrate), (C) MP mice or (D) MP mice treated with 0.250 M sodium butyrate (MP + Butyrate) was used to evaluate the antibacterial activity of the ileum against the Gram-negative bacteria, E.coli. Representative images of E.coli growing on agar plates have been shown in A-D. Each agar plate is divided into quadrants and in quadrant I ileum-derived conditioned media incubated with bacteria was smeared. In quadrant II ileum conditioned media containing only nutrient broth and no bacteria was applied; in quadrant III antibiotic-free DMEM/F12 media incubated with bacteria was smeared; and in quadrant IV antibiotic-containing DMEM/F12 media incubated with bacteria was applied. Figure E is the normalized data of the %bactericidal activity of the ileum supernatants from PP, PP + butyrate, MP and MP + butyrate mice. Data in A-D were analyzed by repeated-measures one-way ANOVA and Tukey’s multiple comparisons test was used for post hoc analysis. Data in E was analyzed by two-way ANOVA and Tukey’s multiple comparisons test was used for post hoc analysis. P values for relevant comparisons have been indicated. All data are mean ± SEM and scatter represents data from samples from individual mice. Sample sizes are 5 per group for A-D. Sample sizes for E are: 5/group.

Figure 9. Effect of morphine, fentanyl, FMT and sodium butyrate on the expression of Reg3γ in the ileum.

The expression of the AMP, Reg3γ, which is active against Gram-positive bacteria, was measured in the ileum in chronic morphine-treated mice administered (A) FMT or (B) 0.250 M sodium butyrate, and in the ileum in chronic fentanyl-treated mice administered (C) 0.250 M sodium butyrate. Data in A and B were analyzed by two-way ANOVA analysis and Tukey’s post hoc test was used for multiple comparisons between groups. Data in C were analyzed by one-way ANOVA and Tukey’s post hoc test was used for comparisons between groups. Relevant P values are indicated. Data are mean ± SEM and scatter represents data from ileum tissues of individual mice. Sample sizes for A are 9 (sham groups), 10 (PP-FMT groups), and 10 (MP-FMT groups). Sample sizes for B are 9 (Saline + Saline), 7 (Saline + Morphine), 5 (Butyrate + Saline), and 5 (Butyrate + Morphine). Sample sizes for C are 7 (Saline + Butyrate), 9 (Fentanyl + Saline), and 9 (Fentanyl + Butyrate).

2. Morphine sulfate (National Institute on Drug Abuse Drug Supply Program, Bethesda, MD) was diluted in saline to 1, 2, 4, and 8 mg/mL. Mice were injected intraperitoneally twice daily with saline or increasing doses of morphine as follows: Day 1– 20 mg/kg morphine, Day 2– 40 mg/kg morphine, Day 3– 40 mg/kg morphine, and Day 4– 80 mg/kg morphine. Mice were used on test day 5 either in the warm-water tail-withdrawal experiment in Figure 4A, for measuring body weight change in Figure 6B, for evaluating intestinal epithelial permeability in Figure 7B, for measuring the expression of Reg3γ in Figure 9B, or in antibacterial activity experiments in Figure 12 as described in the following methods. The same animals were not used to generate different types of data unless indicated.

Figure 12. Effect of naloxone on morphine-induced decrease in antibacterial activity of the ileum.

The effect of the non-selective opioid receptor antagonist, naloxone, was evaluated on the inhibitory effect of morphine on the antibacterial activity of the ileum against (A) L.reuteri, and (B) E.coli. Data are normalized %bactericidal activity and presented as mean ± SEM. Scatter represents data from ileum-derived conditioned media from individual mice. Data were analyzed by one-way ANOVA and Tukey’s multiple comparisons test was used for post hoc analysis. P values for relevant comparisons have been indicated. Sample sizes in A and B are 6, 6, and 7 for saline, morphine and morphine + naloxone groups, respectively.

Fentanyl:

Fentanyl HCl (National Institute on Drug Abuse Drug Supply Program, Bethesda, MD) was diluted in saline to 0.03 mg/mL. Mice were injected intraperitoneally twice daily with saline or 0.3 mg/kg fentanyl for 6 days. Mice were used on day 7 in the warm-water tail-withdrawal experiment in Figure 4C, and for measuring the expression of Reg3γ in Figure 9C as described in the following methods.

Sodium butyrate:

Sodium butyrate (ThermoFisher Scientific, Waltham, MA) was prepared in saline at concentrations of 0.125, 0.250, 0.500, and 1.000 M and administered twice daily by oral gavage. For the dose-response experiment in Figure 4A, mice injected with ramping doses of morphine (as described above) were orally administered saline or different concentrations of sodium butyrate (0.125, 0.250, 0.500, or 1.000 M) for four days. Antinociception was measured on day 5 using the warm-water tail-withdrawal test described below. In all subsequent experiments, 0.250 M sodium butyrate or its vehicle, saline, was administered twice daily through oral gavage. The number of treatments with sodium butyrate was contingent on the duration of exposure to morphine or fentanyl, such that mice subcutaneously implanted with morphine pellets were administered 0.250 M sodium butyrate for six days (Figures 3A, 4B, 5, 6C, 6C, 7B, 8, 9B, 10, 11 and S4), mice injected with ramping doses of morphine received 0.250 M sodium butyrate for four days (Figures 4A, and 6B), and mice injected with fentanyl received 0.250 M sodium butyrate for 6 days (Figures 4C, 6D, and 9C).

Naloxone HCl:

Naloxone HCl (Sigma-Aldrich, St. Louis, MO) was prepared in saline at concentrations of 0.2, 0.4, and 0.8 mg/mL and injected intraperitoneally twice daily at escalating doses 10 minutes before the administration of morphine sulfate in the following manner: Day 1– 2 mg/kg naloxone, Day 2– 4 mg/kg naloxone, Day 3– 4 mg/kg naloxone, and Day 4– 8 mg/kg naloxone. The doses of naloxone were 1/10^th of the doses of morphine sulfate. Ileum tissue was collected on day 5 for use in the antibacterial activity assay in Figure 12.

Fecal Microbiota Transplant (FMT):

Fresh fecal pellets (100 mg) from placebo or morphine-pelleted mice were collected on day 7 and suspended in 1.2 mL of cold (4°C) phosphate- buffered saline (PBS) containing 10% glycerol. The suspension was homogenized and then centrifuged at 800xg for three minutes. The supernatant was transferred to a separate tube and stored at −80°C. The concentration of total bacteria was determined by measuring optical density (OD), such that OD = 0.5 represented 1×10⁸ cells. 100 μL of the fecal supernatant (1 × 10⁹ cells/dose) was then administered twice daily for six days to recipient mice groups via oral gavage according to the following scheme: 1. Placebo-pelleted mice that did not receive fecal microbiota transplants (PP-Sham), 2. Morphine-pelleted mice that did not receive fecal microbiota transplants (MP-Sham), 3. Placebo-pelleted mice that received fecal microbiota from placebo-pelleted donor mice (PP + PP-FMT), 4. Morphine-pelleted mice that received fecal microbiota from placebo-pelleted donor mice (MP + PP-FMT), 5. Placebo-pelleted mice that received fecal microbiota transplants from morphine-pelleted donor mice (PP + MP-FMT), and 6. Morphine-pelleted mice that received fecal microbiota transplants from morphine-pelleted donor mice (MP + MP-FMT) (Figure 1C).

Antinociceptive response tests:

The warm-water tail-withdrawal and hot-plate assays were used in the present study. In the warm-water tail-withdrawal test, the distal 1/3 tail was immersed in a water bath at 56°C. The latency to withdraw the tail from the warm water was recorded. A maximum cutoff of 10 seconds was set to prevent tissue damage. On test day 7 in morphine-pelleted mice (Figures 1D, 4B, and S3), test day 5 in morphine-injected mice (Figure 4A), and test day 7 in fentanyl-injected mice (Figure 4C), baseline responses were recorded, following which the mice received acute morphine (10 mg/kg s.c.). 25 minutes later, tail-flick latencies were recorded to test for the development of tolerance to the 10 mg/kg morphine challenge. Antinociception induced by 10 mg/kg morphine was quantified as the percentage of maximum possible effect (%MPE), such that: %MPE = [(challenge latency−baseline latency) / (Maximum cutoff−baseline latency)] × 100.

In the hot-plate assay, individual mice were placed on a Syscom Model 35D hot-plate set at 56°C, and the latency to lick their hind paw or jump was recorded. A maximum cutoff of 30 seconds was set to prevent tissue damage. Baseline responses were recorded on test day 7 in morphine-pelleted mice (Figure 1E), following which the mice were injected with acute morphine (10 mg/kg s.c.). 25 minutes later, hot-plate responses were measured again to test for the development of tolerance to the acute morphine challenge. Antinociception induced by 10 mg/kg morphine was quantified as %MPE as described above.

Measuring body weight:

On day 1 the baseline body weight of individual mice was noted prior to any drug treatment. Body weights were also noted on test day 7 in morphine-pelleted mice, test day 5 in morphine-injected mice, and test day 7 in fentanyl-treated mice before utilizing them in the warm-water tail-withdrawal assay. The percent change in the body weight of mice was determined using the formula: [(Body weight on test day – baseline body weight on day 1) / (baseline body weight)] × 100.

Detection of butyrate in blood, colon, and stool:

Blood, colon tissue samples, and fecal material were collected from morphine or placebo-pelleted mice on day 7. Samples were immediately homogenized 1:4 with deionized water and stored at −30°C. Seven-point calibration curves of 10–1000 μg/g butyrate (Sigma-Aldrich, St. Louis, MO), a butyrate-free control, and a negative control free of butyrate and the internal standard (ISTD) were prepared. Butyrate was extracted and analyzed using a modified previously published method²³. In brief, 100 μL of methanol containing 20 μg butyrate-1,2-¹³C₂ (Sigma-Aldrich, St. Louis, MO), the ISTDs, was added to 0.20 g aliquots of each calibrator, control, or specimen except the negative control. Samples were mixed for five minutes, centrifuged for 30 minutes, and then left for 30 minutes at 4°C. 100 μL of the clear supernatant was transferred into a new tube and washed with 100 μL propyl formate. Samples were mixed for five minutes and centrifuged for 30 minutes before transferring 50 μL of the organic layer to GC vials for analysis. Gas chromatography-mass spectrometer (GC-MS) analysis was performed on a Shimadzu GC/MS-QP2020 NX Single Quadrupole GC-MS (Shimazu, Kyoto, Japan) controlled by GCMS solution software (Shimadzu, Kyoto, Japan). Chromatographic separation was performed using a ZB-FFAP column, 30 m × 32mm, 0.25 μm (Phenomenex, Torrance, CA). A sample volume of 2 μL was injected in splitless mode with an injector temperature of 200°C. The carrier gas was Helium with a 2 mL/minute flow rate. The initial oven temperature of 55°C was held for four minutes, then ramped to 130°C at 50°C/minute and held for 3.7 minutes. Finally, the temperature was raised to 250°C at 30°C/minute and held for two minutes. Linear regression of the peak area of ratios of the quantification ion for butyrate (72 m/z) and the ISTD quantification ion (75 m/z) was used to construct the calibration curves. For each analytical run, the coefficient of determination (r²) was higher than 0.996. The concentrations of each calibrator were determined to be within ± 20% of their expected concentration.

Measuring GI transit using the carmine red dye assay:

On day 7, morphine or placebo-pelleted mice injected with saline or 0.250 M sodium butyrate were administered 150 μL of non-absorbable carmine red dye solution, 6% w/v carmine red dye powder in 0.5% w/v carboxymethylcellulose, via oral gavage. Each animal was placed in an empty cage with no bedding, and had free access to water, but no food. The time from gavage to the first appearance of the red dye in the fecal pellet was noted as total transit time. A maximum cutoff time of 360 minutes was set for the experiment.

Intestinal permeability assay:

On the test day, i.e., day 7 in pelleted mice and day 5 in intermittently injected mice, animals were orally gavaged with FITC-conjugated dextran (100 mg/ml in PBS, Sigma-Aldrich, St. Louis, MO) at a dose of 44 mg/100 g body weight of FITC-labeled dextran. After four hours, mice were anesthetized with isoflurane, and 300–500 μl of blood was collected by cardiac puncture. Serum collected from blood samples by centrifugation for 15 minutes at 1500xg and 4°C was diluted with an equal volume of PBS. 100 μl of diluted serum was transferred to a 96-well plate, and FITC concentration was fluorometrically quantified by emission spectrometry (Promega, Madison, WI) at 528 nm using an excitation wavelength of 485 nm. Serum from mice not administered FITC-dextran was used to determine background. All concentrations were measured against a standard curve of serially diluted FITC-dextran.

Bactericidal activity assay:

The bactericidal activity assay was performed based on the procedure described by Udden et al.²⁴.

Preparation of conditioned media from ileum tissue samples:

4–5 cm of the distal ileum was resected and immediately flushed with sterile-filtered ice-cold PBS to remove digesta. Ileum tissue samples were cut longitudinally, rinsed in sterile-filtered ice-cold PBS, and weighed. Tissue samples were disinfected in 5 ml DMEM/F12 media supplemented with 5% FBS and 1x antibiotics (penicillin, streptomycin, and vancomycin) for two hours at 37°C in a 95%O₂/5%CO₂ incubator. After disinfection, residual antibiotics were washed off by rinsing the samples three times with 5 ml antibiotic-free DMEM/F12 media supplemented with 5% FBS. Rinsed ileum tissue samples were then cut into 1 cm pieces using sterile scissors and transferred to 12-well cell culture plates containing fresh antibiotic-free DMEM/F12 media supplemented with 5% FBS. 1 ml of DMEM/F12 supplemented with 5% FBS was used per 100 mg of tissue. Samples were incubated at 37°C in an incubator with 5% CO₂ and 95% O₂ for 12 hours. Tissue supernatants (or conditioned media) were subsequently transferred into sterile 1.5 ml centrifuge tubes. Tissue debris was sedimented by centrifugation at 12,000 × g at 4°C for five minutes, and the conditioned media was used for the antibacterial activity assay.

Antibacterial activity assay:

The prototypical Gram-negative bacteria, Escherichia coli (E. coli strain HB101), was inoculated in 5 ml of Luria-Bertani (LB) broth and incubated overnight at 37°C with constant shaking at 250 rpm. Cultured bacteria were collected by centrifugation at 1,200 × g for 10 minutes at 4°C and then resuspended in fresh LB broth at a final concentration of 1×10⁵ cells/mL. 20 μL of the diluted bacteria were added to 500 μL of the ileum-derived conditioned media and incubated for one hour in an incubator maintained at 37°C and 95% O₂/5% CO₂. An additional 20 μL of the diluted bacteria were incubated with 500 μL of DMEM/F12 media supplemented with 5% FBS and 1 x antibiotics (penicillin, streptomycin, and vancomycin; positive internal control) and with 500 μL antibiotic-free DMEM/F12 media containing 5% FBS (negative internal control). 500 μL of the ileum-derived conditioned media supplemented with 20 μL of fresh LB broth was also incubated along with the other samples to check for the presence of contamination (sham control). Thus, an experiment with each ileum-derived conditioned media constituted four groups: A) ileum supernatant + E. coli, B) ileum supernatant + LB broth, C) negative control, and D) positive control. A BHI agar plate was divided into four quadrants, and 50 μL per group was evenly applied to each quadrant. The agar plate was incubated at 37°C, and bacterial colonies were counted 15–18 hours later. The experiment was repeated for ileum supernatants prepared from different mice.

The prototypical Gram-positive bacteria, Lactobacillus reuteri (L.reuteri strain ATCC 53608), was cultured in de Man, Rogosa, and Sharpe (MRS) broth supplemented with 0.001% Tween 80 in a 95% O₂/5% CO₂ incubator maintained at 37°C. The cultured bacteria were collected by centrifugation, resuspended in fresh MRS broth, and serially diluted to a final concentration of 1 × 10⁵ cells/mL. The activity of the ileum tissue supernatants against L. reuteri was tested using the methodology described above for E. coli. Briefly, each ileum tissue supernatant experiment consisted of four groups: A) ileum supernatant + L.reuteri, B) ileum supernatant + MRS broth, C) DMEM/F12 media + 5% FBS + L.reuteri (negative control), and D) DMEM/F12 media + 5% FBS + 1x antibiotics (penicillin, streptomycin, and vancomycin) + L.reuteri (positive control). 50 μL per group was uniformly smeared onto MRS agar plates divided into four quadrants, and the total number of bacterial colonies formed was determined after incubation for 15–18 hours.

RNA isolation and qRT-PCR:

Total RNA was extracted from the ileum using TRIzol reagent (ThermoFisher Scientific, Waltham, MA). RNA samples were treated with DNase 1 (RNase-free, ThermoFisher Scientific, Waltham, MA) to remove DNA contamination. Quantitative real-time polymerase chain reaction (qRT-PCR) was performed on a Mini-Opticon real-time PCR system (Bio-Rad, Hercules, CA) by using the iTaq Universal SYBR Green One-Step kit (Bio-Rad, Hercules, CA) as described previously²⁵. Gapdh was used as the internal control. Primers used in this study were: murine Reg3γ forward, 5′-CGTGCCTATGGCTCCTATTGCT-3′; murine Reg3γ reverse, 5′-TTCAGCGCCACTGAGCACAGAC-3′; murine Gapdh forward 5′-CCATGGAGAAGGCTGGGG-3′; and murine Gapdh reverse 5′-CAAAGTTGTCATGGATGACC-3′ (Integrated DNA Technologies, Inc., Skokie, Illinois).

Microbiome profiling in butyrate-treated fecal samples:

Fecal pellets were collected from saline-treated placebo-pelleted, butyrate-treated placebo-pelleted, saline-treated morphine-pelleted, and butyrate-treated morphine-pelleted mice. DNA was extracted using the QIAamp Fast DNA Stool Mini Kit (QIAGEN, Aarhus, Denmark) according to the manufacturer’s protocols, and DNA was sent to CosmosID (Cosmosid Inc, Rockville, MD) and subjected to whole genome shotgun sequencing using the Illumina platform. An average of 5.921M reads per sample was achieved with a minimum of 4.514M reads and a maximum of 8.816M reads. CosmosID’s k-mer based approach was used for taxonomic identification by comparing sequences to an in-house database. Profiles were analyzed using the filtered species-level data containing 297 bacterial species. Counts were renormalized to the mean number of reads with a pseudo count added to each bacterial species count and the counts were log10 transformed.

Microbiome profiling in FMT-treated fecal samples:

Fecal pellets were collected from PP + PP-FMT, PP + MP-FMT, MP + PP-FMT and MP + MP-FMT-treated mice. Fecal samples from mice that did not exhibit a tail-flick response at baseline in the warm-water tail-withdrawal test were not used for downstream processing. Microbial DNA was isolated from the fecal samples using the QIAamp PowerFecal Pro DNA Kit (QIAGEN, Aarhus, Denmark) according to the manufacturer’s protocols, and subjected to whole genome shotgun sequencing using the Nextseq 2000 platform from Illumina. An average of 10.456M reads were received and their quality was verified using FastQC v0.12.1²⁶. Reads that matched the mouse genome mm10 were removed using bowtie2 v2.5.2²⁷ and samtools v1.19²⁸. The unmapped reads were then used for taxonomy calling at the species level using Kraken2 v2.1.3²⁹, Bracken v2.0³⁰ and mouse gastrointestinal bacteria catalogue³¹. All the metagenomic data was assembled into a single table using the kraken-biom v1.0.1.

Blinding:

Experimenters were not blinded while performing experiments. However, separate investigators conducted the experiments to ensure reliability of results.

Data analysis:

The threshold for statistical significance was P < 0.05. Post hoc analysis of the ANOVA was performed only for significant main effects or significant interactions unless indicated. GraphPad Prism (version 10.1.2) was used for data analysis. Data are presented as mean ± SEM.

Warm-water tail-withdrawal test and hot-plate test:

Data represented as %MPE in Figures 1 and S3 were evaluated by two-way ANOVA with FMT treatment and morphine treatment as the two independent variables. The Tukey’s multiple comparisons test was used for post hoc analysis. Data represented as %MPE in Figure 4A were evaluated by two-way ANOVA with butyrate dose and morphine treatment as the two independent variables, and in Figure 4B, by two-way ANOVA with butyrate treatment and morphine treatment as the two independent variables. Tukey’s post hoc test was used for multiple comparisons between groups. Data represented as %MPE in Figure 4C were evaluated by one-way ANOVA with treatment as the independent variable. Tukey’s post hoc test was used for multiple comparisons between groups.

Butyrate levels in blood, colon, and stool:

Data in Figure 3A were analyzed by two-way ANOVA with butyrate treatment and morphine treatment as the independent variables. Tukey’s multiple comparisons test was used for post hoc analysis. Data in Figures 3B and 3C were analyzed by unpaired two-tailed Student’s t-test.

Intestinal permeability assay:

Serum concentrations of FITC-dextran were evaluated in Figure 7A by two-way ANOVA with FMT treatment and morphine treatment as the independent variables and in Figure 7B by two-way ANOVA with butyrate treatment and morphine treatment as the independent variables. Tukey’s post hoc test was used for multiple comparisons between groups.

Bactericidal activity assay:

The total number of bacterial colonies formed on agar plates were converted to colony forming units (CFU)/mL, such that CFU/mL = (number of colonies × dilution factor) / 50 μL. Data in Figures 8A–C and 10A–D were evaluated by repeated-measures one-way ANOVA with the media as the independent variable. Tukey’s post hoc test was used for multiple comparisons between groups. CFU/mL data were transformed into percent bactericidal activity in Figures 8E, 10E, and 12, such that % Bactericidal activity = {[(CFU/mL of antibiotic-free DMEM/F12 media + bacteria) - (CFU/mL of Ileum supernatant + bacteria)] / (CFU/mL of antibiotic-free DMEM/F12 media + bacteria)} * 100. Data in Figure 8E was assessed by one-way ANOVA with treatment as the independent variable. The Holm-Sidak post hoc test was used for multiple comparisons between groups. Data in Figure 10E was evaluated by two-way ANOVA with butyrate treatment and morphine treatment as the two independent variables. Multiple comparisons between groups were made using Tukey’s post hoc test. Data in Figure 12 was evaluated by one-way ANOVA with treatment as the independent variable. Tukey’s multiple comparisons test was used for post hoc analysis.

qRT-PCR:

Relative expression of Reg3γ to Gapdh was calculated using the 2^−ΔΔCt method, and values were expressed as fold change. Data were analyzed in Figure 9A by two-way ANOVA with FMT treatment and morphine treatment as the independent variables and in Figure 9B by two-way ANOVA with butyrate treatment and morphine treatment as the independent variables. Data in Figure 9C were analyzed by one-way ANOVA with treatment as the independent variable. Multiple comparisons between groups were made using Tukey’s post hoc test.

Microbiome analysis of the butyrate-treated fecal samples:

The Permutational multivariate analysis of variance (PERMANOVA) using the Bray–Curtis dissimilarity index was utilized to evaluate the beta diversity of the fecal bacteria between the different groups. Results of the statistical analysis were obtained using the CosmosID Hub. The alpha diversity index, Chao1, for all the groups was determined using the CosmosID Hub. The data were analyzed by unpaired two-tailed Student’s t-test or two-way ANOVA, and Tukey’s post hoc test was used for pairwise comparisons using GraphPad Prism (version 10.1.2).

Microbiome analysis of the FMT-treated samples:

The data was analyzed in R v4.3.2 using phyloseq³². The beta diversity index and alpha diversity index (Chao1) were calculated using the distance and estimate_richness functions of the phyloseq package. The relative abundance for all samples was also determined. The Chao1 and relative abundance data were analyzed using two-way ANOVA, and Tukey’s post hoc test was used for pairwise comparisons using GraphPad Prism (version 10.1.2). PERMANOVA using the Bray–Curtis dissimilarity index was utilized to evaluate the beta diversity of the fecal bacteria between the different groups. P values were corrected for multiple corrections using the Benjamini-Hochberg test.

RESULTS

Chronic morphine exposure alters the composition of the fecal microbiota.

Evaluation of the alpha diversity (within group diversity) of fecal bacterial communities using the Chao1 Index revealed significantly increased bacterial abundance in the saline-treated morphine-pelleted (MP) animals compared to the saline-treated placebo-pelleted (PP) animals (Figure 1A). Principal coordinate analysis (PCoA) of the beta diversity (diversity between groups) of the fecal bacteria measured with the Bray-Curtis Index revealed discrete clustering of the saline-treated MP mice and saline-treated PP mice (Figure 1B, Table S1). Further analysis of the bacterial taxa at the phylum level showed contraction of Firmicutes and Actinobacteria, and expansion of Bacteroidetes in saline-treated MP mice (Figure S1). Altogether, the data indicated that chronic morphine treatment altered the composition of the fecal bacteria.

Fecal microbiota transplants from placebo-pelleted mice inhibited the development of tolerance to morphine-induced antinociception.

Opioid-induced dysbiosis has been implicated in the development of tolerance to opioid-induced antinociception¹¹. Since chronic morphine exposure induced dysbiosis, we investigated whether fecal microbiota transplants (FMT) altered the development of tolerance to morphine-induced antinociception (Figure 1C). A two-way ANOVA analysis of the antinociceptive effect produced by 10 mg/kg morphine in the warm-water tail-withdrawal test revealed a significant FMT x morphine treatment interaction [F (2, 44) = 19.36, P < 0.001] (Figure 1D). Chronic morphine-treated mice responded poorly to the 10 mg/kg morphine challenge compared to PP mice (28.5 ± 4.2 %MPE vs. 100 ± 0 %MPE in PP mice; Figure 1D), indicating the development of tolerance. FMT from PP mice (PP-FMT) prevented the development of tolerance to morphine-induced antinociception. 10 mg/kg morphine produced significant antinociception in MP mice treated with PP-FMT (88.9 ± 7.0 %MPE; Figure 1D). Interestingly, FMT from MP mice (MP-FMT) did not produce tolerance in PP mice.

A two-way ANOVA analysis of the hot-plate data indicated a significant interaction of FMT x morphine treatment [F (2, 21) = 6.154, P = 0.008] (Figure 1E). Chronic morphine-treated mice were tolerant to the 10 mg/kg morphine challenge dose in the hot-plate test (40.1 ± 14.6 %MPE vs. 100 ± 0 %MPE in PP mice; Figure 1E). Consistent with the warm-water tail-withdrawal test results, PP-FMT prevented the development of tolerance to morphine-induced antinociception in MP mice. MP mice treated with PP-FMT exhibited significant antinociception to 10 mg/kg morphine (89.2 ± 10.8 %MPE; Figure 1E). Thus, treating MP mice with the microbiome of PP mice prevented the development of tolerance to morphine-induced antinociception in the warm-water tail-withdrawal and hot-plate tests. However, treating PP mice with the microbiome of MP mice did not produce tolerance to morphine-induced antinociception (Figures 1D and 1E). 10 mg/kg morphine produced significant antinociception in PP mice treated with MP-FMT (Figures 1D and 1E). These results suggest that chronic exposure to morphine is required for the induction of tolerance to morphine-induced antinociception.

Fecal microbial composition in FMT-treated mice.

We next determined in a separate cohort if FMT treatment altered the fecal microbial composition of the recipient mice. Two-way ANOVA analysis of the Chao1 diversity indices revealed a significant main effect of morphine treatment on the alpha diversity of the recipient mice [F (1,16) = 6.118, P = 0.02] (Figure 2A). A significant interaction of morphine treatment x FMT treatment was not detected [F (1, 16) = 0.1373, P = 0.72] (Figure 2A). Post hoc analysis revealed that the alpha diversity of the PP + MP-FMT group was significantly decreased compared to that of the MP + PP-FMT group (Figure 2A). PCoA of the beta diversity revealed significant clustering based on morphine treatment (Figure 2B, Table S2). All MP mice clustered distinctly and more tightly along PC1 compared to PP mice, irrespective of FMT treatment (Figure 2B, Table S2). Significant variability was observed in the beta diversity of the PP mice and no distinct clustering was observed between the PP + PP-FMT and PP + MP-FMT groups (Figure 2B, Table S2). At the phylum level the Firmicutes/Bacteroidetes relative abundance ratio between the FMT-treated groups was similar (Figure S2A). However, differences were noted in the relative abundance of Verrucomicrobia (Figure S2B), specifically Akkermansia muciniphila was altered (Figure 2C). Increase in Akkermansia muciniphila correlated with the development of tolerance to morphine (Figures 2C and S3). MP + MP-FMT mice, which were tolerant in the warm-water tail-withdrawal test, exhibited increased abundance of Akkermansia muciniphila (Figures 2C and S3). The abundance of Akkermansia muciniphila was reduced in the MP + PP-FMT mice, which were not tolerant to morphine, (Figures 2C and S3). Administering MP-FMT to PP mice did not alter the relative abundance of Akkermansia muciniphila (Figure 2C).

Exposure to chronic morphine reduced endogenous butyrate concentration in the stool.

Butyrate levels are substantially reduced in fecal samples of opioid users^4,7. Furthermore, the phylum Firmicutes, which comprises butyrate-producing bacteria²², was reduced in chronic morphine-treated mice (Figure S1). Therefore, we analyzed the concentration of endogenous butyrate in fecal samples, colon tissue, and whole blood in chronic morphine-treated mice using GC-MS (Figure 3). The concentration of butyrate in fecal samples of MP mice was significantly reduced compared to that of PP mice (386.1 ± 34.5 vs. 161.0 ± 12.3 μg/g, respectively). Daily oral supplementation with sodium butyrate restored butyrate levels in the fecal samples of MP mice (Figure 3A). Daily butyrate supplementation in placebo mice did not increase butyrate concentrations in the stool, likely due to quorum sensing³³. We observed no change in the concentration of butyrate in colon tissue [13.7 ± 2.5 vs. 12.8 ± 3.8 μg/g, t (df) = 0.186 (8), P = 0.86] or whole blood [8.6 ± 0.7 vs. 7.5 ± 0.4 μg/mL, t (df) = 1.466 (8), P = 0.18] between PP and MP mice (Figures 3B and 3C, respectively). These data suggest that in chronic morphine-treated mice, butyrate production is significantly impaired due to altered intestinal microbiota.

Butyrate administration prevented the development of tolerance to opioid-induced antinociception.

We next investigated if oral supplementation with sodium butyrate altered the development of tolerance to morphine-induced antinociception. Morphine was administered intermittently (Figure 4A) or continuously through subcutaneously implanted pellets (Figure 4B). Figure 4A shows that chronic morphine-treated mice were tolerant to morphine-induced antinociception when sodium butyrate was not administered. The 10 mg/kg morphine challenge did not produce significant antinociception (32.0 ± 17.6 % MPE vs. 100.0 ± 0.0 % MPE in the saline + 0 M butyrate-treated mice). Oral administration of sodium butyrate prevented the development of tolerance to morphine-induced antinociception in a dose-dependent manner. Significant inhibition of tolerance to morphine-induced antinociception was observed starting with the 0.250 M sodium butyrate dose (89.9 ± 5.0 % MPE vs. 32.0 ± 17.6 % MPE in the morphine + 0 M butyrate-treated mice; Figure 4A). A two-way ANOVA analysis comparing % MPE of the 10 mg/kg morphine challenge in saline or chronic morphine-treated animals administered with increasing doses of sodium butyrate revealed a significant butyrate dose x morphine treatment interaction [F (4, 40) = 4.015, P = 0.0008] (Figure 4A). Consistent with results in Figure 4A, tolerance to morphine-induced antinociception was observed in MP mice in the warm-water tail-withdrawal assay (2.0 ± 0.9 % MPE vs. 85.4 ± 14.6 % MPE in the PP + saline group) and oral administration of 0.250 M sodium butyrate significantly reduced tolerance to morphine-induced antinociception (45.1 ± 15.4 % MPE vs. 2.0 ± 0.9 % MPE in the MP + saline group, Figure 4B). A two-way ANOVA analysis of the % MPE of 10 mg/kg morphine in PP or MP mice revealed significant main effects of butyrate treatment [F (1, 16) = 7.373, P = 0.02] and morphine treatment [F (1, 16) = 42.49, P < 0.001] but no interaction effects [F (1, 16) = 1.804, P = 0.20]. These data implied that tolerance to morphine-induced antinociception is inhibited by microbial metabolites such as butyrate irrespective of the route of morphine administration or the method of inducing tolerance.

To test whether butyrate was effective at preventing tolerance induced by more potent opioids, we examined the effect of 0.250 M sodium butyrate on the development of antinociceptive tolerance in fentanyl-treated mice (Figure 4C). One-way ANOVA analysis of the % MPE of 10 mg/kg morphine in saline or fentanyl-treated mice revealed a significant main effect of treatment [F (2, 23) = 7.716, P = 0.003]. Post hoc analysis revealed tolerance to the 10 mg/kg morphine dose in fentanyl-treated mice (68.3 ± 10.4 % MPE in the Saline + Butyrate vs. 24.6 ± 5.3 % MPE in the Fentanyl + Saline group). However, oral administration of sodium butyrate prevented tolerance (66.1 ± 11.5 % MPE in the Fentanyl + Butyrate vs. 24.6 ± 5.3 % MPE in the Fentanyl + Saline group). These data suggested that butyrate prevented the development of tolerance to opioid-induced antinociception, and not tolerance to antinociception induced by morphine alone.

Oral butyrate administration did not prevent morphine-induced inhibition of GI transit.

It is well established that chronic exposure to morphine reduces GI motility³⁴. In Figure 5, we examined the effect of 0.250 mM sodium butyrate on the GI motility of chronic morphine-treated mice using the carmine red dye assay. A two-way ANOVA analysis of the transit time of the carmine red dye in saline or butyrate-treated PP or MP mice revealed a significant main effect of morphine [F (1, 16) = 6.994, P = 0.02]. A significant main effect of butyrate [F (1, 16) = 0.2227, P = 0.64] or a significant interaction of morphine x butyrate treatments [F (1, 16) = 0.3710, P = 0.55] was not detected. These data indicated that chronic morphine treatment increased GI transit time, and that oral sodium butyrate did not affect morphine-induced inhibition of GI motility.

Butyrate or fecal microbiota transplantation did not prevent opioid-induced body weight loss.

We next examined the effect of FMT or oral butyrate on the body weight of chronic morphine-treated mice used in the warm-water tail-withdrawal test from Figures 1D, 4A,4B, and 4C. Although oral administration of PP-FMT (Figure 1D) or 0.250 M sodium butyrate (Figures 4A and 4B) prevented the development of tolerance to morphine-induced antinociception in the tail-withdrawal test, no effect on morphine-induced body weight loss was observed (Figure 6). A two-way ANOVA analysis of the body weight change in FMT-treated pelleted mice revealed a significant main effect of morphine [F (1, 36) = 10.42, P = 0.003], but a significant main effect of FMT treatment [F (1, 36) = 0.1901, P = 0.67] or a significant FMT x morphine interaction [F (1, 36) = 0.1914, P = 0.66] was not detected (Figure 6A). Furthermore, a two-way ANOVA analysis of the body weight change in 0.250 M butyrate-treated mice injected with a ramping dose of morphine [F (1, 16) = 28.79, P < 0.001] (Figure 6B) or morphine-pelleted mice [F (1, 16) = 12.21, P = 0.003] (Figure 6C) revealed a significant main effect of morphine, but a significant main effect of butyrate [F (1, 16) = 0.2108, P = 0.65 in injected mice; F (1, 16) = 0.1095, P = 0.74 in pelleted mice] or a significant butyrate x morphine interaction [F (1, 16) = 1.012, P = 0.33 in injected mice; F (1, 16) = 1.192, P = 0.29 in pelleted mice] was not detected. Consistent with the results in morphine-treated mice, oral butyrate administration did not prevent fentanyl-induced body weight loss (Figure 6D). Altogether, these data suggested that neither FMT nor butyrate mitigate the weight loss caused by opioids.

Butyrate or fecal microbiota transplantation reduced chronic morphine-induced intestinal epithelial barrier disruption.

Increasing evidence indicates that chronic exposure to morphine results in the disruption of the intestinal epithelium^11,35. FMT of commensal microbiota and butyrate are known to protect epithelial barrier function through multiple mechanisms, including regulating the permeability of the epithelium^22,36. Here, the effect of FMT or sodium butyrate on the transepithelial permeability of chronic morphine-treated mice was assessed using FITC-dextran (Figure 7). We detected significantly higher serum levels of FITC-dextran in chronic morphine-treated mice compared to placebo (Figure 7A) or saline (Figure 7B) controls. Since the presence of FITC-dextran in serum indicates enhanced intestinal permeability, these results implicate that chronic morphine exposure caused an increase in passive diffusion by disrupting the intestinal epithelium. PP-FMT (Figure 7A) or sodium butyrate (Figure 7B) significantly reduced serum levels of FITC-dextran in chronic morphine-treated mice. No significant difference in serum levels of FITC-dextran was observed between PP mice and FMT-treated MP mice (Figure 7A) or between saline and chronic morphine + sodium butyrate-treated mice (Figure 7B). These data demonstrate that the intestinal epithelium remained intact despite chronic morphine treatment.

Butyrate or FMT prevented morphine-induced disruption of the antimicrobial activity of the small intestinal epithelium.

AMPs contribute to the innate immune response of the intestinal epithelium by maintaining the intestinal microbiota and preventing colonization of the epithelium by pathogenic strains²¹. It is unclear if the altered antimicrobial activity of the intestinal epithelium contributes to opioid-induced dysbiosis. Therefore, we investigated whether chronic morphine treatment perturbed the antibacterial activity of the ileum and if sodium butyrate or FMT inhibited morphine’s effects (Figure 8A). We first examined antimicrobial activity against the prototypical Gram-positive bacteria, L.reuteri (Figure 8). Incubation of L.reuteri with conditioned media from the ileum of PP mice resulted in significantly reduced colonies on the agar plate compared to when the bacteria were incubated with antibiotic-free nutrient media (6.9 × 10⁶ ± 9.4 × 10⁵ CFU/mL vs. 2.3 × 10⁷ ± 9.2 × 10⁵ CFU/mL, respectively; Figure 8B), indicating that the conditioned media exhibited activity against Gram-positive bacteria. In comparison, conditioned media prepared from the ileum of MP mice yielded similar colony numbers to when the bacteria were incubated in antibiotic-free nutrient media (2.1 × 10⁷ ± 1.3 × 10⁶ CFU/mL vs. 2.0 × 10⁷ ± 1.1 × 10⁶ CFU/mL, respectively; Figure 8C), suggesting that the antibacterial activity of the conditioned media decreased after chronic morphine treatment. Interestingly, the antibacterial activity of the ileum-conditioned media was somewhat restored upon treatment with sodium butyrate (Figure 8C). The conditioned media from the ileum of mice treated with morphine and sodium butyrate yielded fewer colonies compared to when the bacteria were incubated with antibiotic-free nutrient media (1.5 × 10⁷ ± 1.2 × 10⁶ CFU/mL vs. 2.0 × 10⁷ ± 9.6 × 10⁵ CFU/mL, respectively; Figure 8D). A comparison of the normalized data in Figure 8E revealed that chronic morphine treatment significantly reduced bactericidal activity of the ileum against Gram-positive bacteria, which was partially restored with oral butyrate treatment (70.1 ± 3.6% in PP mice vs. −3.6 ± 9.9% in MP mice vs. 23.6 ± 8.6% in MP +butyrate-treated mice).

The REG3 family of antimicrobial peptides is highly expressed in the small intestine. Murine REG3γ, and its human analog, REG3α, have been reported to exhibit activity against Gram-positive bacteria^37,38. Since REG3γ is transcriptionally regulated, we investigated whether morphine altered the expression of the Reg3γ gene in the ileum and if FMT of commensal microbiota comprising butyrate-producing bacteria (Figure 9A) or direct administration of exogenous butyrate (Figure 9B) inhibited morphine effects. Reg3γ expression was significantly reduced after chronic morphine treatment, i.e., in MP mice compared to PP controls (Figure 9A) and morphine-injected mice compared to saline controls (Figure 9B). Interestingly, FMT of from placebo-pelleted mice (PP-FMT) prevented the downregulation of Reg3γ in morphine-pelleted mice. However, FMT from morphine-pelleted mice (MP-FMT) produced no effect on Reg3γ expression in placebo-pelleted mice, and Reg3γ expression in the ileum of these mice remained significantly upregulated (Figure 9A). The absence of the effect of MP-FMT on Reg3γ expression indicated that the antimicrobial response of the epithelial barrier remained intact and that chronic treatment with morphine was necessary for inducing barrier dysfunction. Consistent with the results of PP-FMT treatment on Reg3γ expression in morphine-treated mice, oral administration of sodium butyrate also prevented morphine-induced downregulation of Reg3γ in the ileum (Figure 9B). Sodium butyrate did not alter the expression of Reg3γ in saline-treated control mice (Figure 9B). The absence of an effect could be due to quorum sensing by the autochthonous microbiota³³. Consistent with the results in morphine-treated mice, fentanyl treatment downregulated Reg3γ in the ileum, but treatment with 0.250 M butyrate prevented this effect (Figure 9C). These data indicated that the commensal microbiota and/or butyrate regulate Reg3γ transcription to reverse chronic opioid-induced effects.

Next, we tested the activity of the ileum supernatants against the prototypical Gram-negative bacteria, E.coli (Figure 10). Bacterial colony counts were significantly diminished when E.coli were incubated with conditioned media from the ileum of PP mice instead of antibiotic-free nutrient media (5.9 × 10⁷ ± 2.7 × 10⁶ CFU/mL vs. 8.1 × 10⁶ ± 4.0 × 10⁶ CFU/mL, respectively; Figure 10A), indicating that the ileum-derived conditioned media exerted substantial antibacterial activity against Gram-negative bacteria. Sodium butyrate did not affect the antibacterial activity of the ileum-derived conditioned media. E.coli colony counts were substantially reduced when incubated with the conditioned media from the ileum of PP mice treated with sodium butyrate instead of antibiotic-free DMEM/F12 media (5.0 × 10⁷ ± 2.3 × 10⁶ CFU/mL vs. 6.2 × 10⁶ ± 1.9 × 10⁶ CFU/mL, respectively Figure 10B). Figure 10C shows the reduced antibacterial activity of ileum supernatants obtained from chronic morphine-treated mice. Incubation of E.coli with conditioned media from the ileum of MP mice resulted in a non-significant reduction in colony numbers compared to when the bacteria were incubated with antibiotic-free nutrient media (3.8 × 10⁷ ± 2.9 × 10⁶ CFU/mL vs. 4.8 × 10⁷ ± 3.4 × 10⁶ CFU/mL, respectively). Conditioned media derived from the ileum of chronic morphine-pelleted mice treated with sodium butyrate exerted significant antibacterial activity (Figure 10D). E.coli colony counts were substantially larger when antibiotic-free nutrient media was used instead of conditioned media from the ileum of sodium butyrate-treated chronic morphine-pelleted mice (5.9 × 10⁷ ± 3.4 × 10⁶ CFU/mL vs. 1.3 × 10⁷ ± 3.4 × 10⁶ CFU/mL, respectively; Figure 10D). A comparison of the normalized data in Figure 10E revealed that the % Bactericidal activity of morphine-treated ileum media was significantly reduced compared to that from PP mice (17.5 ± 10.0% vs. 87.1 ± 6.0%, respectively). While oral administration of sodium butyrate did not alter the antibacterial activity of the ileum from PP mice, it significantly increased the bactericidal activity of the ileum of MP mice (87.9 ± 3.9% and 77.3 ± 6.4%, respectively; Figure 10E). These data indicated that chronic treatment with morphine inhibited the homeostatic activity of the ileum from fighting against Gram-positive and Gram-negative bacteria and that oral administration of sodium butyrate can prevent these effects.

Oral butyrate administration altered the fecal bacterial composition in morphine-treated mice.

Analysis of the Chao1 Index of alpha diversity revealed that the bacterial abundance in butyrate-treated MP mice was diminished compared to saline-treated MP mice and was comparable to saline-treated PP animals. Oral butyrate administration did not alter the Chao1 diversity of the fecal bacteria of PP mice (Figure S4A). Evaluation of the beta diversity using the Bray-Curtis Index revealed distinct clustering of butyrate-treated MP mice and saline-treated MP mice (Table S1; Figure S4B). PERMANOVA analysis indicated a significant difference in the beta diversity of butyrate-treated MP mice and saline-treated PP mice (Table S1; Figure S4C). PERMANOVA analysis did not reveal differences in the beta diversity of saline-treated PP mice and butyrate-treated PP mice (Table S1; Figure S4D). At the phylum-level, butyrate prevented the enrichment of Verrucomicrobia, specifically, the abundance of Akkermansia muciniphila was reduced (Figure 11). Altogether, these data indicated that butyrate altered the fecal microbial composition of chronic morphine-treated mice.

Naloxone antagonized the morphine-induced decrease in the antimicrobial activity of the ileum.

We next tested if the inhibitory effect of morphine on the antibacterial activity of the ileum was opioid receptor-mediated. Morphine treatment reduced the activity of the ileum supernatants against L.reuteri (52.5 ± 2.1% in morphine-treated vs. 73.8 ± 1.7% in saline-treated; Figure 12A) and against E.coli (36.1 ± 14.5% in morphine-treated vs. 93.3 ± 2.8% in saline-treated) (Figure 12B). Concomitant treatment with naloxone, a non-selective opioid receptor antagonist, prevented morphine effects (L.reuteri: 69.9 ± 2.6%; E.coli: 83.4 ± 4.8%; Figure 12), thus indicating that the effect of chronic morphine exposure on the antibacterial activity of the ileum is opioid receptor-mediated.

DISCUSSION

In this paper we show that the antimicrobial activity and the AMP, Reg3γ, in the intestine is markedly reduced in an opioid-dependent mouse model. Oral supplementation with fecal transplants from morphine-naïve mice or the short chain fatty acid, butyrate, prevented the decrease in antimicrobial activity, downregulation of Reg3γ, and increase in the abundance of Akkermansia muciniphila. Treatment with butyrate or fecal transplants from opioid-naïve mice also attenuated tolerance to opioid-induced antinociception in mice. However, no effect on chronic opioid-induced decrease in body weight or GI motility was observed. Altogether, the results indicate that opioid-induced disruption of the antimicrobial activity of the intestinal epithelium contributes to enteric dysbiosis and impacts the development of antinociceptive tolerance.

In the present study, chronic morphine exposure altered the alpha and beta diversities of the fecal microbiota. We observed an increased abundance of Bacteroidetes and Verrucomicrobia, and depletion of Firmicutes in the feces of chronic morphine-treated mice. The intestinal epithelium restricts bacteria from colonizing the intestine tissue and entering systemic circulation through multiple contingencies that maintain homeostasis between the intestinal epithelium and the gut bacteria^19,21,39. One mechanism is the secretion of AMPs into the intestinal lumen by enterocytes and crypt Paneth cells²¹. Previous studies have reported reduced antimicrobial activity, and systemic translocation of luminal bacteria if Paneth cells are damaged, or their activity is disrupted²⁰. Genetic deletion of the AMP, Reg3γ, results in altered mucus distribution, increased colonization of the intestinal epithelium by mucosa-associated bacteria, bacterial translocation, and elevated inflammatory responses in the intestine^40,41. Similar effects were observed in mice chronically exposed to morphine or fentanyl. Chronic morphine treatment increased intestinal epithelial permeability and morphine or fentanyl treatment downregulated Reg3γ. Antimicrobial activity against both Gram-positive and Gram-negative bacteria was also reduced. Interestingly, Akkermansia muciniphila, a Gram-negative bacteria in the phylum, Verrucomicrobia, was enriched with chronic morphine treatment. Akkermansia muciniphila is a mucin-degrading bacteria that was previously reported to be abundant in hydromorphone-treated DSS-colitis mice¹⁷. It exerts strain-specific effects on intestinal inflammation such that certain strains are pro-inflammatory and disrupt the intestinal epithelial barrier^42–44. Our findings suggest that impaired ability to contain bacteria might contribute to opioid-induced intestinal dysbiosis, including expansion of Akkermansia muciniphila and predispose to systemic translocation of luminal bacteria via disruption of the epithelial barrier.

Naloxone, a non-selective opioid receptor antagonist, prevented morphine effects on the antimicrobial activity of the ileum, indicating that this effect is opioid receptor-mediated. In the mouse gastrointestinal tract, μ-opioid receptors have not been detected in intestinal epithelial cells; they are primarily expressed in enteric neurons and gut-innervating extrinsic neurons^1,45. There is increasing evidence that enteric neurons and gut-innervating extrinsic neurons can modulate the response of the intestinal epithelial barrier to gut microbiota^46–48. Morphine is known to modulate the excitability of these neurons by altering the activity of voltage-gated ion channels^12,49–52. Thus, altered activity of neurons might form the basis for opioid-induced intestinal barrier dysfunction and the resultant dysbiosis.

Fecal microbiota transplants from PP mice prevented morphine-induced disruption of the intestinal epithelial barrier. These results were consistent with findings reported previously by Banerjee et al.¹⁴. Interestingly, PP-FMT induced improvement in the epithelial barrier function but did not result in significant changes in the fecal microbial composition in chronic morphine-treated mice. Changes were only observed in the relative abundance of Akkermansia muciniphila, which could be attributed to the improved epithelial barrier function in the MP + PP-FMT mice. Further studies are required to investigate if FMT treatment alters the composition of other bacterial species that intimately interact with the intestinal epithelium as they may represent a population that is distinct from the fecal microbiota.

Metabolomic analysis of the fecal samples of PP and MP mice revealed reduced butyrate levels in chronic morphine-treated mice. Several factors could contribute to the low fecal butyrate levels in chronic morphine-treated mice, including the depletion of the phylum, Firmicutes, which comprises butyrate-producing bacteria²². In fact, reduced butyrate levels have been noted in the stool samples of human patients exposed to opioids⁷. Butyrate is well-known for its positive effects on host-gut microbiota interactions as it engages pathways that regulate epithelial permeability, inflammation, and production of AMPs, including Reg3γ^53–59, via activation of G-protein-coupled receptors (GPR41, GPR43, and GPR109A) and/or transcriptional regulation of genes by histone deacetylase inhibition and histone acetyltransferase activation^60–63. In the present study, butyrate administration protected against the morphine-induced increase in epithelial permeability, conserved the antimicrobial activity of the intestinal epithelium and inhibited morphine- or fentanyl-induced downregulation of Reg3γ. Although we did not measure butyrate levels post FMT, the metagenomic data shows reduced abundance of Verrucomicrobia with both butyrate and FMT in morphine treated mice. Thus, the improved epithelial barrier function of the intestinal epithelium in chronic opioid-treated mice supplemented with butyrate might explain the shifts in the bacterial community, especially Akkermansia muciniphila.

We observed that oral administration of PP-FMT or sodium butyrate to chronic morphine-treated mice and sodium butyrate to chronic fentanyl-treated mice—treatments that improved the epithelial barrier function—attenuated the development of tolerance to morphine-induced antinociception. The inhibition of tolerance in morphine-treated mice was dependent on the dose of sodium butyrate and independent of the opioid used to induce tolerance. Consequently, these results indicated that improving the intestinal epithelial barrier function prevented the development of tolerance to opioid-induced antinociception. These findings add to the growing body of evidence of the role of the gut in mediating substance use disorders^64–66. Although SCFAs can cross the blood-brain barrier, approximately 95% of orally administered butyrate is absorbed locally in the intestine and extremely low concentrations of oral butyrate are detected in systemic circulation or brain^67–69. Thus, the effect of orally administered sodium butyrate on the development of tolerance to morphine- or fentanyl-induced antinociception is most likely through peripheral processes mediating the crosstalk between the gut and the brain. Indeed, accumulating evidence indicates that gut microbes and their fermentation byproducts, such as SCFAs, can modulate neural circuits of the PNS that relay information to the CNS^{11,12,66,70–72}. For example, in vivo oral treatment with broad-spectrum antibiotics¹¹ or sodium butyrate⁶⁶ prevented the development of morphine-induced tolerance¹¹ or hyperexcitability⁶⁶, respectively, in dorsal root ganglia neurons.

Inhibiting myosin light-chain kinase using ML-7 has been previously used as a strategy to prevent morphine-induced intestinal epithelial barrier disruption and bacterial translocation³⁵. However, improving the intestinal barrier with ML-7 did not prevent morphine-induced constipation³⁵. Similarly, butyrate, which improved the intestinal epithelial barrier, did not alleviate morphine-induced constipation in mice. Furthermore, morphine- or fentanyl-induced body weight loss remained unaltered by butyrate. Nonetheless, butyrate precluded tolerance to opioid-induced antinociception (present study) and alleviated opioid-induced thermal hyperalgesia⁶⁶. This suggested a distinct role for the intestinal epithelium in modulating mechanisms underlying the nociceptive pathway.

In the present study, FMT from placebo-pelleted donors attenuated tolerance in chronic morphine-treated mice. The reverse was not observed when FMT from morphine-pelleted donors was given to placebo -pelleted mice that did not develop tolerance to morphine. These results differ from those previously reported by Zhang et al.¹⁶ that noted the induction of tolerance to morphine-induced antinociception in germ-free mice treated with FMT from chronic morphine-treated animals. A key difference between the two studies may be that germ-free mice exhibit significant developmental deficits, especially in the gut-associated immune system⁷³. In the present study we did not deplete the commensal bacteria in the recipient mice to allow for colonization by FMT in the presence or absence of antimicrobial activity.

LIMITATIONS OF THE PRESENT STUDY

In the present study, there was a significant reduction in the antimicrobial activity of the intestinal epithelium of chronic morphine-treated mice against Gram-negative bacteria. However, the identity of specific AMPs active against Gram-negative bacteria was not determined. One possible candidate could be Reg3β, an isoform of Reg3γ, which is inducible and exhibits activity against Gram-negative bacteria⁷⁴. Like Reg3γ, Reg3β limits mucosa-associated bacteria, and ablation of Reg3β results in dysbiosis, bacterial colonization of Peyer’s patches, systemic translocation of gut bacteria, and reduced survival of mice infected with pathogenic bacteria^75–77. In this study, we have used outbred mice, with a genetic diversity reflective of the human population, however this can induce “within-group” variability as noted in the beta diversity of placebo mice.

Another limitation of the current study is that only male mice were investigated. Studies in humans and rodents have reported sex as a biological variable influencing opioid-induced antinociception⁷⁸. Additionally, sexual dimorphism in the composition of gut bacteria has been noted in laboratory animals and humans, and studies have shown that sex hormones actively influence the gut microbiome⁷⁹. Further studies are planned to explore the influence of sexual dimorphism on the findings of this study.

CONCLUSION

The intestinal epithelium plays a critical role in controlling the composition of the intestinal microbiota and helps maintain a symbiotic relationship with commensal bacteria. Here, we report that chronic treatment with morphine or fentanyl reduced the antimicrobial activity of the intestinal epithelium via opioid receptors, implicating a potential mechanism underlying opioid-induced dysbiosis. Preventing opioid-induced disruption of the epithelial barrier function with FMT or sodium butyrate inhibited the development of tolerance to antinociception. Finally, dysbiosis alone is insufficient for inducing tolerance to morphine-induced antinociception and disruption of the intestinal epithelium is required for the development of tolerance. In conclusion, our results implicate a mechanism by which morphine disrupts homeostasis in the microbiota-gut-brain axis.

ABBREVIATIONS

AMP

Antimicrobial peptides

FMT

Fecal microbial transplant

Reg3γ

Regenerating islet-derived 3 gamma

SCFA

Short-chain fatty acid

MP

Morphine-pelleted

PP

Placebo-pelleted

FITC

Fluorescein isothiocyanate

CFU

Colony forming units

VIP

Vasoactive intestinal peptide

%MPE

Percentage maximum possible effect

LB broth

Luria-Bertani broth

MRS broth

de Man, Rogosa, and Sharpe broth

DATA AVAILABILITY

The data supporting these findings have been deposited with NCBI BioProject at https://www.ncbi.nlm.nih.gov/bioproject (Reference PRJNA992486).