Evaluation of a therapy for Idiopathic Chronic Enterocolitis in rhesus macaques (Macaca mulatta) and linked microbial community correlates

Subjects & housing

This study was conducted between June 2014 and July 2015. Housing and handling of animals were in accordance with the Animal Welfare Act and the Guide for the Care and Use of Laboratory Animals (US Public Health Service). TNPRC is an AAALACi-accredited institution. All protocols and procedures were reviewed and approved by the Tulane University Institutional Animal Care and Use Committee. Subjects included 18 (12 male, six female) juvenile (2.00–3.99 years old) Indian-origin rhesus macaques (Macaca mulatta) reared in outdoor breeding groups in the TNPRC specific pathogen free (SPF) breeding colony. The SPF breeding colony has been derived to exclude the following: Simian retrovirus Type D (SRV), simian T-lymphotrophic virus (STLV1), simian immunodeficiency virus (SIV), macacine herpesvirus 1 (MHV-1), and tuberculosis (Mycobacterium tuberculosis). SPF animals are sampled and monitored at least biannually.

Subjects were selected from animals admitted to the hospital for evaluation of gastrointestinal disease. Inclusion criteria included the following: 45 days of soft or liquid stool observed in a 90 day period, three negative fecal parasite evaluations within a five day period, and three negative fecal bacterial cultures within a five day period. Stool samples were collected into sterile feces tubes (Sarstedt, Nümbrecht, Germany) using the scoops provided in each individual feces tube. Samples were collected from each subject’s cage pan within approximately 2 h of the AM light cycle. After collection the samples were immediately placed into −80 °C storage until shipment. Fecal parasitological evaluation was performed via standard zinc sulfate flotation and fecal smear light microscopic analysis. Aerobic and anaerobic fecal microbiological culture was performed to detect the presence of common nonhuman primate enteric bacterial pathogens (Shigella flexneri, Campylobacter jejuni, Yersinia enterocolitica, Salmonella spp., Escherichia coli). Stool samples were plated on MacConkey agar (BBL, Cockeysville, MD, USA), Hektoen enter agar (BBL), Campy CVA agar (BBL), CIN Yersinia agar (BBL), and Yersinia enrichment broth (Oxoid, Ltd., Basingstoke, UK). Balantidum coli, trichomonads, and Campylobacter coli were considered to be opportunistic organisms. Macaques yielding positive test results for the above pathogens were excluded from the study.

Twelve animals met inclusion criteria and were determined to be suffering from idiopathic chronic enterocolitis (ICE). Of these subjects, six were randomly chosen for ICE treatment and six assigned to the untreated group. No subject received antibiotics or other medications during the 90-day pre-treatment observation period. In addition, six animals matched for age and viral status with no history of gastrointestinal disease, normal physical examinations, and normal complete blood counts and serum chemistries were identified and enrolled to serve as untreated, healthy controls. Over the course of the study, all subjects were housed in the same room in standard indoor nonhuman primate cages at TNPRC (Covington, LA, USA). In an attempt to maximize conspecific social contact without interfering with individual animal fecal observations, animals within the same experimental group were initially allowed full contact pair housing on an intermittent basis during the day (approximately 0800 to approximately 1500) and separated into single housing overnight during the 90 day observation periods. During the treatment and sample collection phases of this study, animals were singly housed. All cages were furnished with perches, foraging devices, and manipulable objects.

Macaques were fed Purina LabDiet 5000 commercial monkey biscuits (PMI Nutritional International, Brentwood, MO, USA) twice daily. Major ingredients (in order as per the manufacturer’s label) included dehulled soybean meal, ground corn, wheat middlings, ground soybean hulls, porcine animal fat preserved with BHA and citric acid, fish meal, sucrose, whole wheat, calcium carbonate, salt, ground oats, wheat germ and brewers dried yeast. Water was available ad-libitum, lighting was on a 12:12-h light:dark cycle, and temperature was maintained at 72 ± 5 °F (22.2 ± 1.2 °C).

Data collection

Animals were observed daily by three trained and experienced staff for presence and character of feces as per standard husbandry practices at TNPRC. Staff were blinded to treatment group. Standard husbandry practice utilizes a simple 3-point scoring system for stool consistency. Feces were characterized as either normal, soft, or liquid based upon the appearance of the stool in the cage. Feces observed to be soft or liquid were classified as abnormal for the purposes of this study. In order to assess the ongoing health status of healthy controls and subjects with ICE, physical examinations were performed and blood was collected for complete blood counts and serum chemistry. Blood was collected via femoral venipuncture using standard venipuncture techniques in nonhuman primates. Physical examinations on each subject were performed at each access under ketamine HCl anesthesia (10 mg/kg, IM; Butler Animal Health, Dublin, OH, USA). Complete blood count (CBC) and serum chemistries (Chem) were performed on blood samples obtained at the time points listed below. Blood and stool from all groups were collected for analysis 24 h prior to a duodenoscopy/colonoscopy procedure during which 10–20 duodenal and colonic mucosal biopsy samples were collected via standard endoscopic methods described here. All animals were maintained under general anesthesia. For upper GI endoscopy, an appropriately-sized mouth gag was inserted between the upper and lower dental arcades to prevent damage to the flexible endoscope. An Olympus EVIS Exera GIF Type H180 (Olympus Corporation of the Americas, Center Valley, PA, USA) flexible gastrointestinal endoscope was passed through the mouth gag and advanced through the esophagus into the gastric lumen. The stomach was insufflated with air to facilitate visualization of the pylorus. The stomach was visually inspected for abnormalities, and the endoscope was advanced through the pylorus into the proximal duodenum. The endoscope was then advanced approximately 5–10 cm past the major duodenal papilla. Olympus FB-25K-1 round-cup biopsy forceps (Olympus Corporation of the Americas, Center Valley, PA, USA) were inserted through the biopsy channel of the endoscope and were used to obtain 10–20 mucosal biopsies. Tissue biopsies were immediately placed on ice and stored in −80 °C until shipment. For colonoscopy, no prior preparation was used. An Olympus EVIS Exera GIF Type H180 flexible gastrointestinal endoscope was lubricated with sterile lubricating jelly (Henry Schein, Inc, Melville, NY, USA) and introduced per rectum and advanced retrograde approximately 15–20 cm. Visualization was achieved via saline irrigation. Olympus FB-25K-1 round-cup biopsy forceps were inserted through the biopsy channel of the endoscope and were used to obtain 10–20 mucosal biopsies. Tissue biopsies were immediately placed on ice and stored in −80 °C until shipment. The same veterinarian performed all endoscopic sample collections for this project. Immediately following duodenoscopy/colonoscopy, the six affected animals chosen at random to serve as the ICE-treated group received the following oral medications: 125 mg total vancomycin hydrochloride (Hospira, Inc., Lake Forest, IL, USA) PO four times daily, 50 mg/kg neomycin (Bimeda, Inc., Le Suer, MN, USA) PO twice daily, and 2.5 mg/kg fluconazole (PACK Pharmaceuticals, LLC., Buffalo Grove, IL, USA) PO twice daily beginning on Day 0 and continuing for 14 days. This drug combination was chosen based on anecdotal reports of improvement observed in human cases of chronic gastrointestinal disease. (L Albenberg and G Wu, pers. comm., 2014). We chose to add fluconazole due to reports of yeast overgrowth in the GI tract during antibiotic therapy (Kennedy & Volz, 1985; Samonis et al., 1993). During pre-treatment and inter-treatment observation, no potentially confounding food enrichment was administered with the exception of orange halves through which the oral therapeutic agents were administered. The ICE-untreated and healthy control groups also received unadulterated orange halves on the same schedule as outlined for the ICE-treated group above. The following sample collection schedule applied to all three experimental groups (ICE-treated, ICE-untreated, and healthy controls). On Day 7 post treatment initiation, stool was collected from all animals for analysis. On Day 14 blood, stool, and mucosal biopsies were collected (as described above) from all animals. On Day 21 blood was collected from all participants. On Day 28 blood and stool were collected from all groups prior to a final endoscopy/colonoscopy with mucosal biopsy collection. Stool and mucosal biopsies were stored at −80 °C until shipped on dry ice for processing and sequencing. Additionally, monkey biscuits (Purina LabDiet 5000) were sequenced and analyzed as described below for the fecal and mucosal biopsy samples.

DNA isolation and sequencing

DNA was isolated from stool, mucosal biopsies and food samples using the MoBio PowerSoil Kit (96-well format) in a sterile class II laminar flow hood. Prior to isolation, standard aliquots of sample were weighted and placed into a plate well with beads and buffer, and homogenized for 20 min on the TissueLyser II (Qiagen Valencia, CA, USA). DNA was isolated using the manufacturer’s protocol and then quantified using Picogreen. Amplicons were sequenced using the MiSeq 2 × 250 kit.

Bacterial 16S rRNA gene quantification

Bacterial communities were assessed using the V1–V2 region of the bacterial 16S rRNA gene. The 16S gene was amplified using universal 16S primers 27F and 338R with Golay barcodes for each sample (see Table S1) (Liu et al., 2007). PCR amplifications were conducted in triplicate with AccuPrime Taq DNA Polymerase High Fidelity (ThermoFisher Scientific, Waltham, MA, USA). The reaction mixture, prepared in a PCR clean room, contained 7.21 uL PCR-grade H2O, 2.5 ul 10× buffer II, 0.19 ul Taq, 5 ul each forward and reverse primer (2 uM) and 5 uL DNA template.

The PCR was performed on an Applied Biosystems GeneAmp PCR System 9700 (ThermoFisher Scientific Inc., Waltham, MA, USA) using the following cycling conditions: initial denaturation at 95 °C for 5 min, 30 cycles of denaturation at 95 °C for 30 s, annealing at 56 °C for 30 s, and extension at 72 °C for 90 s, and then a final extension of 8 min at 72 °C. Replicate amplicons were pooled and bead purified using Agencourt AMPure XP (Beckman Coulter, Brea, CA, USA) using the manufacturer’s protocol. The amplicons were sequenced on an Illumina Miseq. Numbers of reads recovered are summarized in Figs. S1A and S1B.

DNA sequencing of the fungal ITS rRNA locus

Fungal communities were assessed by sequencing the eukaryotic ITS1 rRNA locus. This region was amplified using ITS1F (CTTGGTCATTTAGAGGAAGTAA) and ITS2 (GCTGCGTTCTTCATCGATGC) primers specific for fungal ITS regions and Golay barcodes for each sample (see Table S1) (Gardes & Bruns, 1993; Charlson et al., 2012). The reaction mixture was comprised of 12.1 ul PCR-grade water, 2.5 ul Buffer II, 3 ul 10 uM each forward and reverse primer, 0.4 ul of AccuPrime Polymerase, and 4 ul of template. Each reaction was performed in triplicate with the following cycling conditions: initial denaturation at 94 °C for 3 min; 35 cycles of 94 °C for 45 s, then 56 °C for 60 s, then 72 °C for 90 s; and then a final extension at 72 °C for 10 min. The replicates were pooled and bead-purified using Agencourt AMPure XP beads at a 1.0 ratio, and then purified again with a 0.8 ratio to remove primer dimers. Amplification products were checked on a BioAnalyzer and quantified using Picogreen. The amplicons were pooled and a final bead purification at 0.8 ratio was performed to remove remnant primer dimers. The amplicons were sequenced on an Illumina Miseq.

Bacterial community analysis

The 16S rRNA gene sequences were analyzed using the QIIME 1.91 pipeline; additional analysis was conducted in the R programming language. Quality filtering of reads was performed using standard QIIME parameters. Operational taxonomic units (OTUs) were created by clustering the quality-controlled reads at 97% identity using SWARM (Mahé et al., 2014). Representative sequences from each OTU were aligned using PyNAST (Caporaso et al., 2010). OTUs studied are compiled in Table S2. Lineages were attributed using the Greengenes database (McDonald et al., 2012).

Fungal community analysis

ITS reads were processed using the PIPITS (Gweon et al., 2015) pipeline as follows: Reads were joined using PEAR and quality filtered using FAST-X toolkit. ITS1 regions in each read (if present) were identified and extracted using a hidden Markov model via ITSX (Pal et al., 2014). The ITS1 regions were then clustered at 97% identity using VSEARCH (Rognes et al., 2016). Representative sequences were then assigned a taxonomy using the RDP classifier and Warcup V1 ITS database (Deshpande et al., 2016). Subsequent analysis was performed in R. OTUs studied are compiled in Table S3.

Sequence data access

All DNA sequences determined in this study are available at NCBI under PRJNA395230.

Statistical analysis

For both bacterial and fungal communities in stool, indicator lineages were identified via the method of Dufrêne & Legendre (1997) and as implemented in the indicspecies R package (Dufrêne & Legendre, 1997; De Cáceres & Legendre, 2009). The abundances of the OTUs most strongly identified as indicator lineages were used as inputs to a random Forest classifier (R package randomForest, Liaw & Wiener, 2002), which was trained and tested using the default parameters for the package. The OTUs were ranked by importance by measuring the decrease in classification accuracy that resulted from removing each OTU from the classifier. For bacterial and fungal communities in colon biopsy samples, differentially abundant lineages were determined using a negative binomial model as implemented in the R package DESeq2 (Love, Huber & Anders, 2014) with an FDR-corrected p-value less than 0.05. For all differential abundance tests, the study groups were as follows: “Healthy” were samples belonging to monkeys in the Day 0 healthy, untreated group or Day 21 ICE-treated group (n = 12). “Unhealthy” samples were those belonging to monkeys in the Day 0 ICE-treated group or the Day 0 ICE-untreated group (n = 12).

Clinical effects of treatment were assessed by comparing affected and unaffected animals. For the treated subjects, the total proportion of days in which abnormal stool was observed was compared between a 90 day pre-treatment phase after admission to the hospital for diarrhea, and a 90 day post-treatment phase beginning immediately after the 14 day course of treatment. For untreated ICE-affected subjects, values were calculated over corresponding 90-day phases. A repeated measures ANOVA with phase as the within-subjects variable and treatment group as the between subjects variable was conducted with an alpha of 0.05 and a trend defined as p < 0.10. Where significant interactions were found, planned post-hoc comparisons were conducted with an alpha of 0.025 to adjust for multiple comparisons.

Clinical effect

Animals were tested for abnormal stool composition (i.e., liquid, soft, or normal), and the effects of treatment were assessed (n = 6 in each of the three groups). An effect in the treatment group was found (F_(1,10) = 19.3; p < 0.001) with a trend towards interaction between phase (i.e., before or after treatment) and treatment (F_(1,10) = 4.04; p > 0.001). Among the treated subjects, days with abnormal stool fell from 51.9% to 1.7% (p < 0.0005) but remained unchanged in the untreated subjects (55.2% vs. 36.7%; N.S.; Fig. 1). A more dramatic pattern was found for liquid stool alone, the most severe manifestation of the disease, with a main effect of treatment group (F_(1,10) = 32.67; p < 0.0001) and an interaction between treatment group and phase (F_(1,10) = 12.62; p > 0.001). Whereas during the pre-treatment phase 36.5% of affected subjects showed liquid diarrhea, none was observed following treatment (p < 0.0004), and no change was detected among untreated subjects (31.0% vs. 22.2%; N.S.). Abnormal stool was observed on one day for one animal in the healthy untreated control group.

No pathogenic agents were identified via fecal parasitological analysis and microbiological culture of rectal swabs taken serially in each of the ICE-affected animals as part of their initial screening for inclusion. Serial complete blood count (CBC) and serum chemistry (Chem) results from samples taken throughout the observation, treatment, and sample collection periods did not differ from healthy control animals. These findings are consistent with cases of ICE observed at TNPRC and other facilities housing nonhuman primates (McKenna et al., 2008; Broadhurst et al., 2012; Ferrecchia & Hobbs, 2013; Kanthaswamy et al., 2014).

Microbiome analysis of bacterial taxa

We sought to determine whether microbial community structure in macaque gut could be associated with ICE. Samples were collected from all animals in the three groups on specific treatment days in the ICE-treated group and on corresponding days in the untreated groups (ICE-treated and healthy untreated). Collection from treated and untreated animals occurred on Days 0, 14, and 21. Duodenal and colonic biopsies were also collected endoscopically.

After DNA extraction, PCR amplification with 16S rRNA gene primers and sequencing, we recovered an average of 16,484 reads from stool samples, 4,784 from colonic tissue, 246 from duodenum, and 73 from negative controls (Fig. S1). The major bacterial lineages detected are shown in Fig. 2. The most abundant lineages in stool included Prevotellaceae, Ruminococcaceae, Lactobacillaceae, and Streptococcaceae. Other lineages less commonly found in humans, such as Treponema (family Spirochaetaceae) and Bacteriodes S24-7, were also identified. Some macaques in the ICE-treated group showed higher levels of Streptococcaceae than macaques in the ICE-untreated group at baseline. During the period of antibiotic and antifungal treatment of the ICE-cohort, Lactobacillaceae predominated, which was a departure from the baseline composition. We infer that the Lactobacillaceae lineages detected were insensitive or less sensitive to the antibiotic treatment than other community members.

Biopsies yielded fewer sequences, and not all samples yielded useable data. The bacterial populations associated with the tissue sites were distinct from those in feces. Major bacterial lineages in the colon included Helicobacteraceae and Brachyspiraceae and Pasteurellaceae in the duodenum. Here, too, a notable feature was the presence of high Lactobacillaceae in colon biopsies during the period of antibiotic treatment (4/6 samples).

Association of bacterial community structure and ICE

Bacterial community structure was compared using generalized UniFrac, which measures a distance between pairs of communities by aligning all OTUs detected for each pair of samples onto a common phylogenetic tree and scores the shared branch length on the tree (Chen et al., 2012). Generalized UniFrac takes into account both presence/absence and the abundance of OTUs when calculating community distance. Comparison of the three communities at time zero showed a significant difference in community structure among the three groups (Fig. 3A, p < 0.001, PERMANOVA), leaving the influence of ICE in isolation unclear.

We next compared the treated and untreated stool samples over all time points (Fig. 3B). The most notable outliers were the samples collected at Day 14 from the ICE-treated group, associated with the antimicrobial treatment. As mentioned above, the Day 14 samples from the treated group were dominated by certain Lactobacillaceae which we infer are relatively less sensitive to the antibiotic regimen (Fig. S2). Apart from Day 14 treated samples, only a weak effect of ICE versus health was evident (p = 0.015, R2 = 0.1). We did not compare the tissue biopsies due to their lower sample numbers.

We also queried the association of stool consistency and community structure. No significant association was detected (Fig. S3).

Bacterial lineages potentially associated with ICE

We sought to identify bacterial lineages selectively associated with clinical signs of ICE. For the analysis, we pooled healthy and unhealthy specimens. Specifically for healthy, we included untreated control monkeys on Day 1 and ICE-treated monkeys on Day 21 (absence of clinical signs of ICE) in both stool and intestinal biopsy samples. For the unhealthy (clinical signs of ICE) group, we included the ICE untreated group and the ICE-treated group (clinical signs of ICE) on Day 1. We excluded samples from Day 14 of the ICE treated group in favor of Day 21 in order to avoid allowing the effect of active treatment to dominate the classifier.

A Random Forest classifier was developed to sort unhealthy and healthy monkey samples (Fig. 4). Testing of the classifier showed that it could classify the animals into their original groups (no clinical signs or clinical signs of ICE) with 95% accuracy (5% out of bag error rate). As a future control for over fitting, it will be useful to determine whether this model can also distinguish ICE cases from healthy controls in new data sets not used to generate the model.

Assessment of the lineages primarily responsible for discrimination between healthy and unhealthy animals in colonic biopsy samples (Fig. 4A) showed that OTUs of the families Lactobacillaceae and Ruminococcaceae were the most prominently enriched in ICE cases, and are therefore candidates for ICE-associated organisms.

A separate analysis, using the same approach but with only time zero samples, showed that healthy animals were distinguished by the presence of Christensenella (Fig. S4) when compared to macaques with ICE. In these samples, two OTUs (denovo326 and denovo557), annotated as belonging to the Christensenellaceae family, were present at higher levels in the healthy cohort, and were important features for distinguishing between cohorts for the Random Forest classifier.

If the cause of ICE is due to the presence of a pathogenic bacteria, we reasoned that any causal organism should decrease in abundance with treatment, and so investigated changes in colonization over the treatment period (Fig. 4B). All of the OTUs in stool contributing to discrimination changed only slightly after treatment, with none showing a greater than 10% absolute change in abundance.

Discriminative lineages identified in stool were then compared to lineages in intestinal biopsies. Sparse bacterial detections and limited time points precluded generating an effective Random Forest classifier, and further analysis using a negative binomial model failed to identify discriminating taxa. Thus no signal of ICE could be discerned in the biopsy samples.

Fungal taxa are derived primarily from food

Because the successful therapy included an antifungal agent, we investigated fungal lineages for association with ICE. DNA samples from stool were amplified with ITS primers, which are selective for sequences from the eukaryotic rRNA locus. A variety of fungal lineages were detected in the samples (Fig. 5)—relatively abundant taxa in stool included Dothidiomycetes, Sordariomycetes, Eurotiomycetes, and Saccharomycetes. Analysis using PERMANOVA on UniFrac distances between samples showed no significant difference between ICE and control samples (p = 1). Lineages detected in tissue samples were sparse (Fig. 5B), and again no discrimination could be obtained using PERMANOVA. A random forest classifier was constructed as above with the goal of discriminating ICE from healthy animals, but no strongly discriminating taxa were identified.

We next investigated the degree to which lineages detected in our study originated as transients in food. We extracted DNA from macaque biscuits (their major food material) and carried out 16S and ITS rRNA tag sequencing. We found little resemblance between bacterial lineages from stool or biopsy samples and biscuit samples (Fig. S2A). However, fungal lineages were nearly identical in stool and biscuits samples (Fig. 5A). Labeling on the biscuit packaging revealed that brewers dried yeast was a component of the biscuits. We thus conclude that the fungal sequences detected in stool corresponded primarily to transients or free DNA from food, whereas the bacterial sequences were mostly from gut-resident organisms.