Alcohol intoxication combined with burn injury can lead to life-threatening complications, including sepsis, multiple organ failure, and death. After an acute burn, the gastrointestinal system becomes hypoxic because of fluid loss and reduction of intestinal blood flow. This can cause perturbations in the intestinal epithelial barrier, immune function, and the composition of the gut microbiome. Increased gut permeability leads to proinflammatory signaling, contributing to further damage to the intestinal barrier. Recent studies have suggested that IL-27 plays an anti-inflammatory role, which may be beneficial in intestinal barrier repair. Therefore, in this study, we examined the effect of ethanol and burn injury on IL-27 in the small intestine, as well as the potential beneficial role of IL-27 in restoring the intestinal barrier after intoxication and burn. Male C57BL/6 mice were gavaged with 2.9 g/kg ethanol before receiving a ∼12.5% total body surface area scald burn with or without rIL-27 in resuscitation fluid. Our results demonstrate that IL-27–producing cells are reduced in the small intestine after injury. When IL-27 is supplemented in resuscitation fluid, we were able to restore intestinal barrier integrity and transit, mediated through increased intestinal epithelial cell proliferation, reduced inflammatory cytokines, and increased anti-inflammatory cytokine IL-10. We also observed increased gene expression of tight junction proteins. These findings suggest that IL-27 may be a contributor to maintaining proper intestinal barrier function after injury through multiple mechanisms, including preventing excess inflammation and promoting intestinal epithelial cell proliferation and tight junction integrity.
Alcohol intoxication is a major causal factor in adverse health outcomes. Alcohol misuse contributes to roughly 2.5 million deaths worldwide and increases the likelihood of other life-threatening conditions such as acute respiratory distress syndrome and multiple organ failure (1–3). Intoxication is a common confounding factor in numerous traumatic injuries including burns, because it leads to increased risk-taking behavior. Up to 50% of burn patients in the United States each year have detectable alcohol levels at the time of presentation to the emergency department (4, 5). It is well-known that intoxication at the time of burn injury contributes to detrimental sequelae, such as increased secondary infections, multiple organ failure, and death (4, 6, 7). After an acute burn injury, reduction in intestinal blood flow leads to hypoxic conditions in the gut, leading to disruptions in the gastrointestinal (GI) barrier (8–10). Previous work in our laboratory has demonstrated disturbances in intestinal barrier function, immune responses, and increased bacterial translocation from the GI lumen into the host after alcohol intoxication and burn injury (11–15). In addition, our laboratory has demonstrated decreased proliferation of intestinal epithelial cells (IECs) and reduced tight junction protein expression postinjury (16, 17). Although consequences of alcohol and burn injury on the intestinal immune response are complex, previous findings include impaired T cell activity, increased neutrophil infiltration and reactive oxygen species production, and abundant inflammatory mediators in small intestinal tissue (14, 18–20). Thus, after injury, the cross-talk between immune cells and IECs is altered. Improper or disrupted cytokine signaling has been proposed to contribute to gut barrier dysfunction.
IL-27 is a cytokine in the IL-12 family that has been previously studied in T cells for its role in promoting Th1 cell commitment (21). IL-27 is known to be produced by APCs, such as macrophages and dendritic cells, in response to TLR agonist binding, such as LPS (22, 23). It signals through a heterodimeric receptor composed of gp130 and IL-27Rα, which is also known as T cell cytokine receptor (WSX-1) (24). IL-27 has also been shown to induce production of anti-inflammatory cytokine IL-10 (25). Recently, studies have demonstrated a protective role for IL-27/IL-27R signaling in models of intestinal inflammation. IECs have been found to express both subunits of the IL-27R, and models of DSS-colitis and Clostridium difficile intestinal infection demonstrated protective effects of IL-27 on the intestine (26–28). However, changes in intestinal IL-27 after alcohol and burn injury have not been studied.
Therefore, we profiled changes in the production of IL-27 in intestinal immune cells after alcohol and burn injury. We also sought to characterize whether IL-27 administration could help restore intestinal function through improving IEC proliferation and tight junction gene expression and reducing intestinal inflammation after injury. We also investigated whether these combined effects could prevent gut barrier leakiness and delayed intestinal transit. Our results demonstrate that production of IL-27 in the mesenteric lymph nodes (MLNs) and lamina propria (LP) is reduced after injury. Therefore, we used in vitro and in vivo models to determine the beneficial effects of IL-27. IL-27 treatment induced proliferation of IECs and restored gene expression of tight junction proteins Claudin-4 (cldn4) and Claudin-8 (cldn8). IL-27 treatment also reduced inflammation in the intestine and restored gut barrier integrity and transit after injury. These effects occurred even in the presence of inflammatory bacteria present in the intestinal lumen, indicating that IL-27 has beneficial effects on the host after injury.
Materials and Methods
Ten- to twelve-week-old C57BL/6 male mice (23–25 g body weight) were obtained from Charles River Laboratories. IEC-specific VillinCreSTAT3flox/flox knockout (henceforth referred to as STAT3−/−) mice used for small intestinal organoid cultures were a gift from Dr. B. Gao at the National Institute of Alcohol Abuse and Alcoholism and were rederived at Jackson Laboratories (Bar Harbor, ME). Mice were maintained in animal housing facilities at Loyola University Chicago Health Sciences Division, Maywood, Illinois.
Murine model of acute alcohol intoxication and burn injury
Mice were randomly assigned into experimental groups: sham injury + vehicle (water) or burn injury + ethanol. On the day of injury, mice were gavaged with 400 μl of 25% ethanol in water (2.9 g/kg), and vehicle animals were gavaged with 400 μl water. One hour before burn injury, mice were given 1 mg/kg buprenorphine s.c. Four hours after the gavage, mice were anesthetized with a ketamine hydrochloride/xylazine mixture (∼80 and ∼1.2 mg/kg, respectively) given by i.p. injection. The dorsal surface of the mice was shaved before placing the mice in a prefabricated template exposing ∼12.5% total body surface area (TBSA) calculated using Meeh’s formula as described by Walker and Mason (29). Burn group animals were immersed in ∼85°C water bath for ∼7 s to induce a full-thickness scald burn injury. Sham injury animals were placed in a 37°C water bath for ∼7 s (30, 31). After burn or sham injury, the animals were dried gently. Sham animals were given 1.0 ml normal saline resuscitation, and burn animals received 1.0 ml normal saline resuscitation fluid with or without 5 μg rIL-27 (R&D Systems, Minneapolis, MN) by i.p. injection (32). Animals were returned to their cages and allowed food and water ad libitum (14). All animal experiments were conducted in accordance with the guidelines set forth in the Animal Welfare Act and were approved by the Institutional Animal Care and Use Committee at Loyola University Health Sciences Division.
Isolation of MLNs
One day after injury, mice were humanely euthanized. The abdominal cavity was exposed by midline incision, and MLNs were identified by locating the cecum and mesentery surrounding the nodes. The nodes were gently removed using forceps and placed in complete RPMI 1640 media containing 2 mM l‐glutamine, 10 mM HEPES, 50 µg/ml gentamicin, 100 U/ml penicillin, 100 µg/ml streptomycin, and 10% FCS. After collection, MLNs were crushed and passed through a 70-µm cell strainer using the end of a 3-ml syringe. Cells were spun at 300 × g and washed once with complete RPMI media before cell stimulation.
Isolation of Peyer’s patches
The total small intestine was exposed, and between 7 and 11 Peyer's patches (PPs) were collected and placed in RPMI 1640 media supplemented with 2 mM l‐glutamine, 10 mM HEPES, 50 µg/ml gentamicin, 100 U/ml penicillin, 100 µg/ml streptomycin, and 10% FCS. PPs were digested in HBSS containing 0.5 mg/ml collagenase D for 15 min at 37°C. After digestion, PPs were crushed and filtered through a 70-μM cell strainer to obtain single-cell suspensions (11).
Isolation of LP
LP cells were isolated as described previously (33). After removal of PPs, ∼8 cm of the distal small intestine was harvested and opened longitudinally and washed in cold PBS. Small intestines were incubated in HBSS supplemented with 10 mmol/l HEPES, 50 μg/ml gentamicin, 100 U/ml penicillin, 100 μg/ml streptomycin, 5 mM EDTA, and 1 mM DTT (predigestion solution) for 20 min at 37°C with agitation to remove epithelial cells. Tissues were cut into small pieces and incubated in RPMI 1640 supplemented with 2 mmol/l l-glutamine, 10 mmol/L HEPES, 50 μg/ml gentamicin, 100 U/ml penicillin, 100 μg/ml streptomycin, 10% FCS, 0.5 mg/ml Collagenase D (Roche), 200 U/ml DNase I (Sigma-Aldrich), and 0.5 mg/ml Dispase II (Roche) for 20 min at 37°C with shaking, repeated two to three times. The digested tissues were passed through 70-μM cell strainers and centrifuged at 1200 rpm for 10 min at 20°C to pellet isolated cells. Cells were resuspended in complete media before stimulation (11).
Cell stimulation and flow cytometry of lymphocytes
Mixed cells from MLN, LP, and PP were stimulated with cell stimulation mixture (eBioscience) containing PMA and ionomycin in the presence of protein transport inhibitor for 4 h at 37°C. After stimulation, cells were washed with FACS buffer (PBS with 5% FCS). Cells were incubated with Cell Viability Dye (Thermo Fisher) for 30 min to stain dead cells, followed by anti-CD16/32 Abs for Fc receptor blocking. Intracellular staining for IL-27 was performed using Foxp3 Transcription Factor Staining Buffer Set (eBioscience) according to the manufacturer’s instructions with PE-conjugated anti-mouse IL-27 (eBioscience). Flow cytometry data were collected with FACSCanto II (BD Biosciences) and analyzed with FlowJo software (Tree Star) (30).
Isolation of IECs was performed as described previously by Weigmann et al. (33). Small intestines were removed from the peritoneal cavity, and the distal 10 cm of the small intestine was separated from the remainder of the small intestine for analysis. The tissues were cut longitudinally and placed in ice-cold PBS + 1% penicillin/streptomycin mixture. After two washes in PBS + penicillin/streptomycin, tissues were placed in a digestion solution containing 10 mmol/L HEPES, 50 μg/ml gentamicin, 100 U/ml penicillin, 100 μg/ml streptomycin, 5 mM EDTA, and 1 mM DTT in HBSS. Tissues were placed in a 37°C incubator and shaken on a rotator at 250 rpm for 20 min. Tissues were vortexed to separate the epithelial cells from the tissue and passed through a 100-μm filter. Cells were counted on a hemocytometer to determine epithelial cell purity. IECs were then processed for downstream applications (34).
RNA isolation and cDNA synthesis
RNA isolation was performed using a RNeasy Mini Kit (Qiagen, Valencia, CA) according to the manufacturer’s instructions. Genomic DNA was removed by DNase digestion using an RNase-free DNase Set (Qiagen). Concentration of isolated RNA was determined using a NanoDrop 2000 spectrophotometer (Thermo Scientific, Bannockburn, IL). Only samples with a 260/280 ratio of ≥2.0 were used for cDNA synthesis. cDNA synthesis was performed using a High-Capacity cDNA Reverse Transcription Kit (Life Technologies, Carlsbad, CA), and reactions were run on a StepOne Plus Thermocycler (Applied Biosciences, Beverley Hills, CA) per the manufacturer’s instructions.
Expression levels of cldn1, cldn2, cldn4, cldn8, occludin (ocln), tight junction protein 1 (tjp1), Ly6G, and lipocalin 2 (Lcn2) mRNA were analyzed by quantitative real time PCR (qRT-PCR) using TaqMan primer probes and TaqMan Fast Advanced Master Mix (Life Technologies). Target gene cycle threshold (Ct) values were normalized to housekeeping control GAPDH Ct values. Data were calculated using the ΔΔCt method (35).
Small intestinal organoid isolation, culture, and IL-27 treatment
Small intestines were harvested from mice and rinsed in cold PBS. Intestines were cut into 2-mm sections and rinsed 15–20 times in cold PBS. Tissue pieces were resuspended in Gentle Cell Dissociation Reagent (STEMCELL Technologies, Vancouver, BC, Canada) and incubated on a rocker for 15 min. Tissue was rinsed in PBS containing 0.1% BSA five times, collecting the supernatant after each wash into separate tubes. Supernatant was filtered through a 70-μm filter and centrifuged at 290 × g, washed once in PBS containing 0.1% BSA, and centrifuged at 200 × g. Pelleted organoids were resuspended in equal volumes of IntestiCult Organoid Growth Medium (STEMCELL Technologies) and growth factor reduced Matrigel Matrix (STEMCELL Technologies). A total of 50 μl of crypt suspensions was pipetted to form domes on 24-well plates and incubated with IntestiCult Organoid Growth Medium at 37°C and 5% CO2 overnight (36). Recombinant mouse IL-27 (R&D Systems) was added the next morning at 100 ng/ml, and organoids were cultured for 48 h (21). Images for organoid growth were taken on EVOS cell imaging system (Thermo Fisher Scientific) at 10× magnification. Ten random image fields were taken at day 0 (before IL-27 treatment) and day 2. ImageJ Software (National Institutes of Health, Bethesda, MD) was used to quantify organoid areas of the image files.
Organoid staining and immunofluorescence
A total of 20 μl of isolated intestinal crypt suspensions was placed on an eight-well chamber slide and cultured overnight before rIL-27 treatment (100 ng/ml) for 48 h. Organoids were fixed with 4% paraformaldehyde for 30 min, followed by permeabilization with blocking buffer (5% FCS and 0.5% Triton X-100 in PBS) overnight at 4°C. Ki67 Ab (Cell Signaling) and Alexa 488–conjugated secondary Ab (Thermo Fisher Scientific) were used. Organoids were counterstained with ProLong Gold Antifade Reagent with DAPI (Thermo Fisher Scientific). Images were obtained on a Zeiss Axiovert 200M microscope (Zeiss, Jena, Germany) at 100× total magnification. Due to the peripheral staining pattern of Ki67 on organoids, Ki67-positive organoids were classified and counted based on ≥75% of the periphery stained with Ki67. Organoids containing <75% peripheral staining of Ki67 were considered Ki67 negative. A minimum of 50 organoids was counted for each treatment group (37).
Protein isolated from IECs was separated on an SDS-PAGE gel and transferred to a polyvinylidene difluoride membrane. Membranes were probed using rabbit anti-mouse Cyclin D1 Ab (Abcam, Cambridge, U.K.), and an HRP-conjugated anti-rabbit secondary Ab (Cell Signaling) was used for detection. Membranes were exposed on a ChemiDoc (Bio-Rad, Hercules, CA), and densitometric analysis was performed with ImageLab software (Bio-Rad). Bands were normalized to β-actin protein expression.
Total small intestinal tissue or IECs were lysed with cell lysis buffer (Cell Signaling) supplemented with protease and phosphatase inhibitor (Thermo Fisher). Levels of IL-10 (BD Biosciences), IL-6 (BD Biosciences), and KC (R&D Systems) were determined using their respective ELISA kits per the manufacturer’s instructions. Cytokine levels are expressed as picogram per milligram of protein for total small intestinal tissue. Relative cell apoptosis of IECs was determined with Cell Death ELISA Kit (Roche, Basel, Switzerland) per the manufacturer’s instructions. Cell death levels are expressed as relative absorbance measurements at 405 nm and normalized to the amount of protein loaded per well.
16s rRNA analysis of fecal bacteria
Small intestinal contents from animals were aseptically collected. Genomic DNA was isolated using QIAamp PowerFecal Pro DNA Kit (Qiagen, Valencia, CA) according to the manufacturer’s instructions. DNA concentration was measured using NanoDrop 2000 Spectrophotometer. Primers specific for the SSU 16s rRNA of total bacteria (domain level analysis), Enterobacteriaceae, and Lactobacillus were used for qRT-PCR. Primers include forward (F): 5′-ACTCCTACGGGAGGCAGCAGT-3′ and reverse (R): 5′-ATTACCGCGGCTGCTGGC-3′ for total bacteria, F: 5′-GTGCCAGCMGCCGCGGTAA-3′ and R: 5′-GCCTCAAGGGCACAACCTCCAAG-3′ for Enterobacteriaceae, and F: 5′-AGCAGTAGGGAATCTTCCA-3′ and R: 5′-CACCGCTACACATGGAG-3′ for Lactobacillus (Thermo Fisher Scientific). Equal amounts of fecal DNA were loaded, and quantitative PCRs were performed using 1× iTaq Universal SYBR Green Supermix (Bio-Rad) with 300 nM forward and reverse primers. Data are represented as a ratio of Enterobacteriaceae/total bacteria and Lactobacillus/total bacteria (34).
Intestinal barrier permeability measurement
One day after injury, mice were given a gavage of 0.4 ml of FITC-dextran (4D) (22 mg/ml) in PBS. Three hours postgavage, mice were euthanized, and blood was collected by cardiac puncture. Blood was centrifuged at 8000 rpm for 10 min at 4°C, and plasma was collected to measure FITC-dextran levels in circulation. Plasma was read at 480-nm excitation and 520-nm emission wavelengths (30).
Intestinal transit measurement
Mice were given FITC-dextran gavage 1 d after injury. Three hours after gavage, mice were humanely euthanized, and intestinal contents from the stomach, small intestine, and large intestine were collected. The small intestine was divided into three sections (proximal, middle, and distal). Luminal contents were suspended in PBS according to weight and homogenized. Contents were centrifuged, and supernatant was collected for fluorometric analysis at 480-nm excitation and 520-nm emission wavelengths (30).
The data, wherever applicable, are presented as means + SEM and were analyzed using one-way ANOVA with Tukey post hoc test, Student t test, or χ2 test (GraphPad Prism 9). A p value <0.05 was considered statistically significant. Unless noted otherwise, significance is represented as follows: *p < 0.05, **p < 0.01, ***p < 0.001, and ****p < 0.0001.
Alterations in immune function in the intestine from ethanol and burn injury can have consequences that affect numerous pathways, including regulation of the intestinal barrier, defense against invading pathogens, and intestinal motility. As such, elucidating changes in intestinal cytokine production can provide information regarding the effects of trauma, such as intoxication and burn injury. Previous data from our laboratory have demonstrated decreases in cytokine production in intestinal immune cells, including IFN-γ, IL-17, and IL-22, following ethanol and burn injury in our mouse model compared with sham animals (31, 38). However, whether alcohol and burn injury alters release of IL-27 remains to be established. To identify changes in production of IL-27 in GI-associated lymphoid tissues, the MLN, PP, and LP were isolated 1 d after alcohol and burn injury. Mixed cell populations were cultured with cell stimulation mixture containing PMA, ionomycin, and protein transport inhibitor to analyze IL-27 production by FACS analysis. Cells were also stained with Live/Dead viability dye. FACS analysis demonstrated that mice subjected to burn injury in conjunction with ethanol showed a significant decrease (p < 0.05) in the percentage of IL-27–producing cells in the small intestine LP (∼40% reduction) and MLN (∼90% reduction) compared with sham + vehicle group (Fig. 1). Levels of IL-27–producing cells in the PP were unchanged between groups. No difference was observed in IL-27 production between burn + vehicle and burn + ethanol groups, indicating that intoxication may not further decrease IL-27 production in intestinal tissue (data not shown). However, because our model of intoxication and burn does not demonstrate intestinal pathology with burn injury alone, the following in vivo studies were all conducted in the combined ethanol + burn injury (14, 19, 34, 39).
IL-27 has been documented to induce IL-10 production in T cells and provide protective effects in animal models of colitis (28, 40, 41). IL-10 is particularly important in regulating intestinal homeostasis by preventing inflammation. Therefore, we sought to identify whether rIL-27 treatment after ethanol and burn can increase IL-10 production in the small intestine. As shown in (Fig. 2, IL-10 levels are reduced after ethanol and burn injury compared with sham + vehicle levels. With administration of rIL-27, levels of IL-10 in total intestinal tissue are restored to sham + vehicle levels. Intestinal inflammatory mediators after intoxication and burn injury, including IL-6, are elevated after injury (19). To determine whether IL-27 can reduce inflammation in the intestine, total small intestinal tissue was harvested and homogenized for analysis of cytokines via ELISA. (Fig. 2 demonstrates that IL-6 is significantly reduced in IL-27–treated mice compared with the ethanol + burn group.
Neutrophil infiltration into the intestines is well documented after alcohol and burn injury and leads to further tissue damage by release of reactive oxygen species and inflammatory proteins, such as Lcn2 (neutrophil gelatinase-associated lipocalin). To determine whether IL-27 affects neutrophil recruitment to the intestine, we assessed neutrophil chemokine KC by ELISA. As shown in (Fig. 3, IL-27–treated mice have significant reductions in intestinal KC compared with ethanol + burn mice. Reductions in KC may be one potential mechanism to prevent intestinal damage by reducing recruitment of neutrophils. Similarly, qRT-PCR of intestinal tissue demonstrated significant reduction in gene expression of Ly6G, a marker that is abundantly expressed by neutrophils. Finally, gene expression of Lcn2, which can be induced by bacteria or inflammatory cytokines, was also significantly reduced in intestinal tissue when comparing ethanol + burn and rIL-27 treatment groups.
Ultimately, the major consequence of alcohol intoxication and burn injury on the intestine is increased gut leakiness. This causes a series of downstream consequences, including bacterial translocation into the host, leading to further intestinal inflammation and barrier dysfunction. Due to the anti-inflammatory effects described earlier, we sought to determine whether administration of rIL-27 after ethanol and burn injury could prevent gut permeability. To determine intestinal permeability, we gavaged mice with FITC-dextran 3 h before euthanasia. Blood was collected to measure levels of FITC-dextran in circulation. As shown in (Fig. 4, mice with intoxication and burn injury demonstrate significantly higher levels of FITC-dextran in circulation, indicative of increased gut permeability. However, treatment with rIL-27 reduced circulating FITC-dextran to sham + vehicle levels, demonstrating the restored gut barrier after injury. Similar to increased intestinal permeability, a common complication in burn patients is paralytic ileus, or slow intestinal transit (42). To determine whether IL-27 improved GI transit after ethanol and burn injury, we collected intestinal contents from the stomach, three sections of the small intestine, and the large intestine to detect FITC dextran levels along the digestive tract 3 h postgavage. As shown in (Fig. 4, ethanol + burn–injured mice display delayed intestinal transit, with peak FITC fluorescence detected in the stomach and the proximal small intestine. In contrast, sham + vehicle and ethanol + burn + rIL-27–treated mice have normal intestinal transit, with the greatest FITC fluorescence intensity observed in the distal third of the small intestine and the large intestine. Taken together, the data indicate that rIL-27 treatment restores gut barrier integrity and intestinal transit after ethanol intoxication and burn injury.
Mechanisms of barrier dysfunction after alcohol intoxication and burn injury are complex and involve the host response and intestinal microbial changes. In previous studies, our laboratory has demonstrated that levels of inflammatory bacterial species Enterobacteriaceae are elevated and beneficial Lactobacillus is reduced in the GI lumen after alcohol and burn injury (34, 39, 43). To determine whether rIL-27 administration prevented microbial dysbiosis, we analyzed fecal contents from the small intestine by 16s rRNA amplification by qRT-PCR to determine ratios of Enterobacteriaceae and Lactobacillus normalized to total bacteria. As shown in (Fig. 5, relative levels of Enterobacteriaceae are increased in the ethanol + burn group and remain elevated with rIL-27 treatment. Lactobacillus is reduced in the ethanol + burn group and is still reduced with rIL-27 treatment, indicating that IL-27 does not seem to influence these intestinal bacterial species.
An important component that modulates intestinal barrier function and permeability is the expression of tight junction proteins. Previous studies from our laboratory have shown decreased gene expression of a variety of intestinal tight junction proteins in small IECs after ethanol and burn injury (34, 39). However, whether IL-27 influences tight junction gene expression has not been explored. Therefore, we analyzed expression of claudin-4 (cldn4) and claudin-8 (cldn8), tjp1, and ocln after in vivo treatment with rIL-27 of ethanol + burn–injured mice (44). As shown in (Fig. 6, mice treated with rIL-27 showed gene expression of cldn4 and cldn8 similar to sham levels in IECs. However, expression of other tight junction proteins (tjp1, ocln) did not improve after rIL-27 treatment.
In addition, rates of IEC death and proliferation are crucial regulators of intestinal barrier maintenance. To determine whether IL-27 had effects on apoptosis and proliferation of IECs in vivo, we treated mice with rIL-27 in resuscitation fluid. One day after alcohol intoxication and burn injury, mice were sacrificed, and small IECs were isolated. As shown in (Fig. 7, ethanol + burn leads to increased apoptosis of IECs as detected by ELISA. Treatment with rIL-27 does not reduce IEC apoptosis. To assess proliferation, we quantified Cyclin D1 expression in IECs by Western blot analysis. As shown in (Fig. 7, the level of cyclin D1 is reduced after intoxication and burn injury compared with sham animals. However, treatment of animals with rIL-27 after injury restores cyclin D1 to sham + vehicle levels, indicative of recovered IEC proliferation. Taken together, our data indicate that although IL-27 can promote IEC proliferation, it does not significantly reduce apoptosis.
To further determine whether IL-27 can promote proliferation of IECs, we used an in vitro small intestinal organoid culture system that recapitulates the morphology of the intestine through differentiation of intestinal stem cells into specialized cells and formation of villus-like structures (45–48). Images of organoids were taken on day 0 before treatment with or without 100 ng/ml rIL-27. On day 2, images of organoids were taken again to assess growth area. (Fig. 8 shows that rIL-27 treatment led to significantly increased growth of intestinal organoids compared with untreated organoids. Because IL-27 is known to signal through downstream signaling protein STAT3, we next analyzed whether STAT3 was required for IL-27–mediated growth of intestinal organoids. When STAT3−/−-derived intestinal organoids were treated with rIL-27 for 48 h, IL-27 treatment did not significantly increase growth and proliferation compared with untreated organoids. These results suggest that IL-27–mediated growth is dependent on STAT3. To confirm this, we stained organoids treated with or without rIL-27 with Ki67, a marker of cellular proliferation. As demonstrated in (Fig. 9, organoids isolated from wild type (WT) mice show increased staining for Ki67 when treated with IL-27. However, staining of Ki67 does not increase in intestinal organoids isolated from STAT3−/− mice, confirming that proliferation mediated by IL-27 is dependent on STAT3. When the number of Ki67-positive organoids was quantified as measured by >75% Ki67 staining along the periphery of the organoid, WT organoids showed a significant increase in the number of Ki67-positive organoids compared with untreated organoids (p < 0.01). This effect was not observed with STAT3−/− organoids.
Finally, we analyzed expression of tight junction proteins in organoids after treatment with rIL-27. Organoids isolated from WT and STAT3−/− mice were treated with or without 100 ng/ml rIL-27 for 48 h as previously described. After treatment, organoids were collected for RNA isolation and gene expression analysis of tight junction proteins by qRT-PCR. As demonstrated in (Fig. 10, WT organoids showed increased expression of cldn1 (p < 0.01) and cldn2 (p < 0.01). Although not statistically significant, both cldn4 and cldn8 also demonstrated an increase in gene expression after rIL-27 treatment. Furthermore, ocln gene expression was significantly increased with rIL-27 treatment. When STAT3−/−-derived organoids were treated with rIL-27, the only significant increase in gene expression was observed in cldn4 (p < 0.001), as shown in (Fig. 11.
Intestinal homeostasis is a complex process due to the proximity of the microbiome to the host’s intestinal immune system. The consequences of combined ethanol intoxication and burn injury lead to a myriad of complications in the intestine, including inflammatory cell infiltration, inflammatory cytokine production, bacterial overgrowth, disruptions in the IEC barrier, and alterations in beneficial cytokine signaling (11, 18, 39, 43, 45, 49, 50). All of these factors coalesce to promote gut leakiness, which contributes to systemic inflammation and organ injury (51, 52). To our knowledge, no studies have explored the role of IL-27 in acute burn injury to date. Due to the complex interplay between proinflammatory and anti-inflammatory mediators in the intestine, it is important to characterize and determine changes after acute injuries, such as intoxication and burn. IL-27, which has demonstrated a unique and beneficial ability to stimulate production of anti-inflammatory cytokine IL-10 and promote wound healing in colonic epithelial cell lines, is a promising target for understanding the intestinal pathophysiology after alcohol and burn, as well as a potential therapeutic target (25, 27, 53).
In our studies, we determined that IL-27–producing cells were significantly decreased in the MLNs and LP of burn-injured and ethanol + burn–injured animals 1 d after injury. Although both burn injury and ethanol + burn groups demonstrate reduced IL-27 in intestinal immune cells, our laboratory has previously documented that alcohol potentiates the adverse outcomes associated with burn injury, including markers of inflammation, intestinal permeability, and bacterial translocation (11, 14, 18, 54). Our mouse model uses a relatively small burn area (∼12.5% TBSA), in which burn injury alone does not result in intestinal pathophysiology 1 d after injury unless combined with alcohol intoxication (14, 35). Due to the amplifying nature of alcohol intoxication on the intestinal pathophysiology after burn injury, we focused our studies on ethanol and burn–injured mice, with sham + vehicle animals as our control group.
It has been proposed that the intestinal barrier serves as an important impediment to prevent excess inflammation after a severe trauma, such as burn injury, because of its role in sequestering the microbiome away from the host. Innate and adaptive immune cells maintain homeostasis and tolerance through production of anti-inflammatory cytokines and suppressing inflammation (55). Failure or collapse of this regulatory network often leads to intestinal inflammation, as observed in our model of intoxication and burn injury. Our results demonstrated that IL-10 levels in the small intestine were reduced after injury; however, they were rescued to sham + vehicle levels with rIL-27. We also observed decreases in inflammatory mediators IL-6 and KC in small intestinal tissue with treatment. In particular, the reduction in neutrophil chemokine KC after rIL-27 administration is a likely mechanism contributing to the reduced intestinal pathophysiology. Previous studies demonstrated that neutrophil depletion before burn injury prevented intestinal permeability, illustrating the prominent role neutrophils hold as mediators of intestinal damage (56). After rIL-27 treatment, we observed almost no neutrophil-associated markers in intestinal tissue as detected by gene expression of Ly6G, a common neutrophil marker, and Lcn2, or neutrophil gelatinase-associated lipocalin, in total intestinal tissue (57, 58).
To confirm whether IL-27 administration after intoxication and burn injury ultimately restores the gut barrier, we assessed intestinal permeability by FITC-dextran gavage. Our results demonstrate that FITC-dextran leakage into the serum was significantly reduced with IL-27 treatment and normalized to sham + vehicle levels. Furthermore, intestinal transit rate was restored. Delayed GI emptying and postburn ileus is a well-documented phenomenon that contributes to GI dysfunction (59, 60).
A proposed mechanism by which intestinal inflammation and dysfunction occur is the overgrowth of pathogenic bacterial species in the intestinal lumen. A study of burn patients by Earley et al. (43) demonstrated that while healthy patients had abundant Bacteroidaceae and Ruminococcaceae species present in their microbiome, burn patients had significantly elevated levels of pathogenic Enterobacteriaceae. This relative increase in opportunistic pathogens may exacerbate mucosal disruptions observed after alcohol intoxication and burn injury. In a study of fecal transplantation, burn-injured mice receiving a transplant from healthy mice demonstrated reduced intestinal permeability (61). However, we did not find changes in ratios of Enterobacteriaceae or Lactobacillus compared with total bacteria after rIL-27 treatment in injured mice. Mucosal delivery of IL-27 may provide a more direct approach to targeting the intestinal microbiome and dysbiosis. Our results indicate that although IL-27 can prevent inflammation in the intestine in this acute setting, this reduction in intestinal inflammation is independent of changes in levels of Enterobacteriaceae or Lactobacillus.
A major problem after intoxication and burn is the breakdown of the physical barrier composed of IECs, which are normally replaced every 3–5 d (62–64). In our in vivo model, we showed that IL-27 treatment restored expression of cyclin D1, a protein active during the G1 phase of cellular proliferation, to sham levels in IECs. However, this effect was not statistically significant. A study by Varedi et al. (16) identified that intestinal cell proliferation and apoptosis are time and position dependent after burn injury. Using a 60% TBSA burn injury model in rats, it was observed that cellular mitosis was decreased in stem cells within the crypt 6 h postinjury, whereas mitosis was decreased in differentiating cells farther up the crypt 12 h after injury (16). In this model, proliferation was restored by 24 h after injury. More studies are needed to explore the timing and position of IECs affected in our model of alcohol intoxication and burn injury.
We also chose to investigate whether IL-27 could also stimulate proliferation in small IECs by using a 3D organoid culture model. Our findings demonstrate that IL-27 treatment of intestinal organoids significantly increased growth as measured by area over the course of 48 h. When IL-27 binds to its receptor, it has been reported to signal through either STAT1- or STAT3-mediated pathways; however, only STAT3 has been reported to be essential for proliferation in T cells (65, 66). Therefore, we performed the same assay with IEC-specific STAT3 knockout organoids. We observed no change in the area growth when treating with IL-27, demonstrating that the increased growth mediated by IL-27 is dependent on STAT3 signaling in IECs. This was confirmed by no change in Ki67 staining between IL-27 treatment and untreated groups in STAT3−/− organoids.
Along with IEC turnover, expression of tight junction proteins is a crucial component that regulates intestinal barrier permeability. Our laboratory has previously demonstrated reduced expression of cldn4, cldn8, and ocln after acute ethanol intoxication and burn injury (39, 46, 67). We also saw beneficial effects on IEC tight junction protein expression in our in vivo model after rIL-27 treatment in resuscitation fluid. As reported previously, we observed significant decreases in gene expression of cldn4, cldn8, ocln, and tjp1 after ethanol and burn injury. However, the treatment with rIL-27 was able to restore gene expression of cldn4 and cldn8 in injured mice. Our results also demonstrated significantly increased gene expression across several tight junction proteins after a 48-h stimulation with rIL-27 in WT-derived organoids, including cldn1, cldn2, and ocln. In addition, cldn4, cldn8, and tjp1 showed an increasing trend in gene expression after treatment. This effect, however, was not observed in STAT3−/− organoids with the exception of cldn4. These results indicate that expression of tight junction proteins cldn1, cldn2, and ocln may be regulated by STAT3-mediated IL-27 signaling. However, cldn4 may be regulated by IL-27 independent of STAT3.
In summary, the activation of uncontrolled inflammation leads to numerous consequences that affect the function of the GI tract; however, IL-27 can mitigate the damage because of its anti-inflammatory activity and by promoting IEC proliferation and tight junction expression. These beneficial effects on the host were present even in the presence of intestinal dysbiosis. Taken together, our findings illuminate that IL-27 may be a viable therapeutic target after injury and provide protection to the intestinal barrier.
This work was supported by National Institutes of Health Grants R01 AA015731, R01 GM128242, T32 AA013527, and F30 DK123929.
Abbreviations used in this article
The authors have no financial conflicts of interest.