Analysis of the hydrogenotrophic microbiota of wild and captive black howler monkeys (Alouatta pigra) in palenque national park Mexico.код для вставкиСкачать
American Journal of Primatology 73:909–919 (2011) RESEARCH ARTICLE Analysis of the Hydrogenotrophic Microbiota of Wild and Captive Black Howler Monkeys (Alouatta pigra) in Palenque National Park, Mexico NORIKO NAKAMURA1,2, KATHERINE R. AMATO3, PAUL GARBER3, ALEJANDRO ESTRADA4, RODERICK I. MACKIE1,2,5, AND H. REX GASKINS1,2,5,6 1 Institute for Genomic Biology, University of Illinois at Urbana-Champaign, Urbana, Illinois 2 Department of Animal Sciences, University of Illinois at Urbana-Champaign, Urbana, Illinois 3 Department of Anthropology, University of Illinois at Urbana-Champaign, Urbana, Illinois 4 Estación de Biologı´a Tropical Los Tuxtlas, Instituto de Biologı´a, Universidad Nacional Autónoma de México, Mexico 5 Division of Nutritional Sciences, University of Illinois at Urbana-Champaign, Urbana, Illinois 6 Department of Pathobiology, University of Illinois at Urbana-Champaign, Urbana, Illinois Intestinal methanogenesis is one of the major pathways for consumption of hydrogen produced by bacterial fermentation and is considered to affect the efficiency of host energy harvest; however, little information is available regarding the hydrogenotrophic pathways of nonhuman primates in the wild, in general, and of howler monkeys, in particular. Microbial fermentation of plant structural carbohydrates is an important feature in wild howlers owing to the high fiber and low available energy content of leaves, which make up the primary component of their diet. In contrast, captive howlers may consume greater quantities of fruits and vegetables that are higher in water, lower in fiber, and, along with commercial monkey chow commonly added to captive monkey diets, more readily digestible than the natural diet. In this study, we analyzed the composition of methanogens and sulfate-reducing bacteria (SRB) from fecal samples of black howler monkeys (Alouatta pigra) in the wild and in captivity. The hydrogenotrophic microbiota of three groups of monkeys was evaluated by PCR-denaturing gradient gel electrophoresis (DGGE) fingerprinting, small clone library construction, and quantitative real-time PCR. Abundance of methanogens was lower than SRB in all howler monkey groups studied. DGGE banding patterns were highly similar within each wild and captive group but distinct among groups. Desulfovibrionales-enriched DGGE showed reduced microbial diversity in the captive animals compared with their wild counterparts. Taken together, the data demonstrate that environmental or dietary changes of the host imposed by captivity likely influence the composition of intestinal hydrogenotrophs in black howler monkeys. Am. J. Primatol. 73:909–919, 2011. r 2011 Wiley-Liss, Inc. Key words: intestinal hydrogenotrophs; Alouatta pigra; DGGE; quantitative PCR INTRODUCTION Howler monkeys (Alouatta) are among the most folivorous of New World primates [Eisenberg et al., 1972]. Owing to the high fiber and low available energy content of leaves, microbial fermentation of plant structural carbohydrates in the colon is an important digestive feature in howlers. Different metabolic pathways for fermentation result in differences in the flow of carbon and electrons, energy yield from the substrate, and the final fermentation products. However, all bacterial fermentation of dietary fiber results in the production of volatile fatty acids (VFA), hydrogen (H2), and carbon dioxide (CO2). The production of H2 is an efficient mechanism of disposing of reducing power generated during the bacterial metabolism of carbohydrates and protein. However, accumulation of H2 results in the production of electron sink products, such as ethanol, lactate, or succinate. The symbiosis between fermentative bacteria and hydrogenotrophic microbes r 2011 Wiley-Liss, Inc. enables the former to shift fermentation patterns toward the production of more reduced substrates, such as VFA, as opposed to the production of the electron sink products. VFA can be absorbed by the host and may make an important contribution to its energy budget. It has been suggested that howlers are highly dependent on VFA to meet as much as 31% of their daily energy requirement [Milton & McBee, 1983]. Methanogenesis and sulfate reduction are the major hydrogenotrophic pathways in the human gut and play an important role in terminal carbon and Contract grant sponsor: Agricultural Experiment Station at UIUC. Correspondence to: H. Rex Gaskins, University of Illinois, 1207 W. Gregory Drive, Urbana, IL 61801. E-mail: email@example.com Received 23 August 2010; revised 1 April 2011; revision accepted 7 April 2011 DOI 10.1002/ajp.20961 Published online 9 May 2011 in Wiley Online Library (wiley onlinelibrary.com). 910 / Nakamura et al. electron flow in anaerobic gut ecosystems. In the human colon, direct competition among methanogens and SRB may occur for the common substrate, H2 [Nakamura et al., 2010]. Intestinal methanogenesis is considered to affect the efficiency of host energy harvest [Samuel & Gordon, 2006; Turnbaugh et al., 2006; Zhang et al., 2009]. A greater abundance of archaeal sequences was observed in obese humans [Zhang et al., 2009] and mice [Turnbaugh et al., 2006], indicating that interspecies H2 transfer between bacterial and archaeal species is an important mechanism for increasing uptake of VFA by the host. In a large comparative zoo study, it was reported that fecal incubation from one black and gold howler monkey (Alouatta caraya) resulted in methane production [Hackstein et al., 1995], indicating the contribution of methanogenesis as a mechanism of H2 disposal in the colon of howlers. However, no further information is available regarding the hydrogenotrophic pathways of howlers and, in particular, a potentially competing pathway for H2 disposal, sulfate reduction. Thus, in this study, we analyzed the composition of methanogens and sulfatereducing bacteria (SRB) in black howler monkeys (A. pigra) in the wild and in captivity in Mexico. The hydrogenotrophic microbiota was evaluated by PCR-denaturing gradient gel electrophoresis (DGGE) fingerprinting, small clone library construction, and quantitative real-time PCR. This is the first study that provides information on the hydrogenotrophic microbial composition in the colon of wild and captive howler monkeys, and provides insights into how hydrogenotrophic microbes may enable efficient fiber fermentation by this host species. METHODS Animals and Samples The fecal samples used in this study were collected at Palenque National Park, Mexico, by K.R. Amato as part of a larger research project on feeding ecology and gut microbiota. The study included two groups (n 5 14 total) of wild Mexican black howler monkeys (A. pigra), inhabiting an area of continuous forest within the park (Motiepa and Balam) and a group of captive animals (n 5 7) of the same species at a nearby rehabilitation center (AcaJungla). The Motiepa group contained three adult males, two adult females, and three juveniles (o4 years of age). The Balam group contained two adult males, two adult females, and two juveniles. The captive group contained six juveniles and one infant that were confiscated from illegal pet owners and are part of a program for reintroduction to the wild. Feces from each group member were collected once per week over a period of 8 weeks (total eight samples per individual May–July 2009) and those from captive animals during a single day in July 2009. Feces were collected immediately after defecation and stored in Am. J. Primatol. 96% ethanol at 41C, then transported to the University of Illinois at Urbana-Champaign (UIUC). Fecal collection, handling, and storage procedures were approved by the UIUC Institutional Animal Care and Use Committee and Institutional Biological Safety Committee. This research adhered to the American Society of Primatologists’ principles for the ethical treatment of nonhuman primates. Before collection, permits were obtained from Mexico’s National Commission on Protected Natural Areas and the Secretary of the Environment and Natural Resources, as well as the Secretary of Agriculture, Livestock, Rural Development, Fisheries and Food for the export of materials from Mexico and from the Centers for Disease Control and Prevention for import of materials to the United States. DNA Extraction and PCR Amplification Fecal DNA was extracted by using the UltraCleanTM Soil DNA Isolation Kit (MO BIO Laboratories, Carlsbad, CA). PCR amplifications for Archaea 16S V3 rDNA using the primers 344F [Raskin et al., 1994] and 519R [Amann et al., 1995] and enrichment of Desulfovibrionales 16S V3 rDNA were performed as follows: each 25 ml PCR reaction mixture contained 10 ng of extracted DNA, 10 PCR buffer (TaKaRa, Shiga, Japan), 8 mM bovine serum albumin (BSA; New England Biolabs, Ipswich, MA), 200 mM of each dNTP (TaKaRa), 0.5 or 1 mM of each primer, and 0.5 U of TaKaRa Taq polymerase (TaKaRa). PCR was performed by initial denaturation at 941C for 2 min, followed by 40 cycles of denaturation at 941C for 30–60 sec, primer annealing at defined temperature (Table I) for 30–60 sec, and extension at 721C for 1 min. Final extension was performed at 721C for 7 min, or 30 min to reduce the occurrence of artifactual double bands in DGGE [Janse et al., 2004]. Two-step nested amplification was performed to obtain Desulfovibrionales-enriched 16S rDNA fragments [Dar et al., 2005]. The DNA template was first amplified with the Desulfovibrionales group-specific primers DSV 230F and DSV 838R [Daly et al., 2000]. The product obtained was used as a template to produce fragments suitable for DGGE with bacterial universal 16S V3 primers 344F [Raskin et al., 1994] and 519R [Amann et al., 1995]. PCR was performed as mentioned previously, except that a GC-clamp, a series of G and C bases, was added to the forward primers (344F and DSV230F) to improve resolution of bands in the denaturing gradient gel (Table I). PCR-Denaturing Gradient Gel Electrophoresis PCR products were loaded in an 8% (w/v) polyacrylamide gel of 35–60% linear DNA-denaturing gradient. (The 100% [w/v] denaturant solution contains 7 M urea and 40% [w/v] formamide). DGGE was performed using the BioRad D-code system Gut Microbiota of Black Howler Monkeys / 911 TABLE I. Primers and Annealing Temperatures Used for PCR Target Archaea 16S V3 rDNA mcrA Bacteria 16S V3 rDNA Desulfovibrionales 16S rDNA aprA dsrA dsrB Primer Sequence (50 -30 ) 344Fa 519R qmcrA-F qmcrA-R-dc 341Fa 518R DSV230 DSV838 APS-FW APS-RVa DSR1F1 DSR-R DSRp2060Fa DSR4R ACG GGG HGC AGC AGG CGC GA GWA TTA CCG CGG CKG CTG TTC GGT GGA TCD CAR AGR GC GBA RRT CGW AWC CGT AGA AWC C CCT ACG GGA GGC AGC AG ATT ACC GCG GCT GCT GG GRG YCY GCG TYY CAT TAG C SYC CGR CAY CTA GYR TYC ATC TGG CAG ATM ATG ATY MAC GG GGG CCG TAA CCG TCC TTG AA ACS CAC TGG AAG CAC GGC GG GTG GMR CCG TGC AKR TTG G CAA CAT CGT YCA YAC CCA GGG GTG TAG CAG TTA CCG CA Annealing temperature 561Cb 601C 551Cb 611C 601C 601C 551C Reference               a A 40 bp GC clamp (CGC CCG CCG CGC GCG GCG GGC GGG GCG GGG GCA CGG GGG GA, 344F and 341F; CGC CCG CCG CGC CCC GCG CCC GGC CCG CCG CCC CCG CCC C, DSRp2040F; CGC CCG CCG CGC CCC GCG CCC GGC CCG CCG CCC CCG CCC G, APS-RV) was added to the 50 -end. Touchdown protocol was used with the starting temperature of 51C higher than the indicated annealing temperature, which was lowered in the first ten cycles of amplification at a rate of 0.51C/cycle. c Degeneracy was added to two additional nucleotides of the original primer qmcrA-R to avoid mismatches with the reported mcrA sequence of Methanobrevibacter smithii (DQ251046). b (BioRad, Los Angeles, CA). Gels were run in 601C 1 TAE buffer at 150 V for 2 hr and then at 200 V for 1 hr. Following electrophoresis, gels were silver stained and scanned using a GS-710 calibrated imaging densitometer (BioRad). Images were analyzed using Diversity Database software (BioRad). Each band on the gel was identified regardless of the band intensity. Bands of interest were excised from the gel using a sterile pipette tip, and incubated in water overnight to facilitate diffusion of DNA out of the gel. The solution was used for reamplification with the corresponding primer pair (without GC clamp) and the amplification protocol originally used to produce fragments for DGGE. Cloning and Sequencing of Archaea 16S rDNA Sequences Target genes were amplified using Archaea 16S rDNA primers SDArch0333aS15 and SDArch1378aA20 [Lepp et al., 2004], with 35 cycles of PCR and an annealing temperature of 581C. The PCR products were visualized on 2% agarose gels and cloned in One Shot Competent E. coli using the TOPO TA cloning kit (Invitrogen, Carlsbad, CA). Clones were grown overnight on Luria-Bertani agar containing ampicillin. White colonies of ampicillin-resistant transformants were selected and checked for correct size insert by EcoR1 digestion and electrophoresis on 2% agarose gel. To confirm the taxonomic identification of Archaea obtained by DGGE, small clone libraries of 1,032 bp archaeal 16S rDNA sequences were constructed for each wild or captive group. Ten clones were randomly selected from each group, checked for correct size insert, and sequenced using an automated sequencing system (Applied Biosystems, Foster City, CA) at the W. M. Keck Center for Comparative and Functional Genomics (University of Illinois Biotechnology Center, Urbana, IL). Phylogenetic Analysis Because DGGE banding patterns were highly stable within individuals during the 8-week period (Fig. 1), only samples collected in May were used for subsequent analyses and comparisons among individuals or groups. Twenty-one Desulfovibrionales and 3 Archaea 16S rDNA DGGE bands were excised from the respective gels, reamplified, and sequenced. Four of the Desulfovibrionales bands and band A2 of the Archaea 16S bands were not included in the analysis owing to low sequence quality. A phylogenetic analysis was performed to determine the relatedness of amplified sequences to other methanogen sequences in the GenBank database. Sequences were analyzed using the BLASTn program to search for similar nucleotide sequences in the Greengenes database [DeSantis et al., 2006]. Phylogenetic analysis was performed using MEGA version 4 [Tamura et al., 2007]. A phylogenetic tree was constructed using the neighbor-joining method [Saitou & Nei, 1987]. Real-Time Quantitative PCR The mcrA and dsrA sequences were amplified using the primers qmcrA-F and qmcrA-R [Denman et al., 2007] and DSR1F1 and DSR-R [Kondo et al., 2004], respectively. Results were normalized to total bacteria abundance estimated by using universal bacterial 16S rDNA primers 341F and 534R [Muyzer et al., 1993]. Assays were performed using the ABI PRISM 7900HT Sequence Detection System Am. J. Primatol. 912 / Nakamura et al. A B M1 (J-M) B2 (J-F) M2 (J-M) B1 (J-M) M3 (SA-M) B5 (A-F) M4 (A-M) B6 (A-F) M5 (A-M) B3 (A-M) M6 (A-M) B4 (A-M) Fig. 1. Desulfovibrionales-enriched DGGE profiles of Motiepa (A) and Balam (B) groups sampled in May, June, and July, 2009 (left to right). Individual identifications are shown on the top with their ages (J, juvenile; SA, subadult; A, adult) and genders (-M, -F). Samples for B2 were collected only in May and July. Profiles from two samples collected on consecutive days in June are presented for B6 for four DGGE profiles (May, June 20, June 21, and July 2009). (Applied Biosystems, Foster City, CA). The 10 ml reaction mixture contained 10 ng template DNA, 250 nM (dsrA) or 500 nM (mcrA) of each primer, and 10 mM of BSA (New England Biolabs). SYBR Green PCR Master Mix (2 ; Applied Biosystems) was used Am. J. Primatol. for the detection of target sequences. PCR cycles consisted of one cycle of 501C for 2 min and 951C for 2 min, and 40 cycles of 951C for 15 sec and 601C for 1 min. Measurements were done in triplicate. Specificity of amplification was confirmed by dissociation Gut Microbiota of Black Howler Monkeys / 913 16S rDNA profiles using Primer-E software (http:// www.primer-e.com/). curve analysis of qPCR end products by increasing the temperature at a rate of 11C/30 sec from 60 to 901C. Standard curves were generated from 1 to 106 cloned plasmids, which contain the target sequence amplified from the primate fecal samples. Detection limits of both mcrA and dsrA were 100 copies. RESULTS Diversity Analysis of Desulfovibrionales 16S rDNA by DGGE The DGGE banding patterns were highly stable within individuals during the 8-week period (Fig. 1) and highly similar among the individuals of the two wild groups, whereas those of captive animals were less diverse and exhibited different banding patterns compared with the wild howlers (Fig. 2A). PCA Statistical Analysis Shapiro–Wilks normality and Kruskal–Wallis tests were performed by Systat 11 (Systat, Chicago, IL). Principal component analysis (PCA) was performed on the presence/absence of bands in the Desulfovibrionales A Motiepa Juvenile 1 2 3 4 Balam Juvenile Adult 5 6 7 8 AcaJungla (captive) Adult Juvenile 9 10 11 12 13 14 15 16 17 18 19 20 21 14 1 3 4 5 8 22 2 9 15 10 16 23 11 19 20 12 6 7 13 17 21 18 B Fig. 2. Desulfovibrionales 16S rDNA-enriched DGGE profiles of wild (Motiepa and Balam) and captive groups. 1: M1, 2: M2, 3: M3, 4: M4, 5: M5, 6: M6, 7: M7, 8: M8, 9: B1, 10: B2, 11: B3, 12: B4, 13: B5, 14: B6, 15: AJ2, 16: AJ3, 17: AJ4, 18: AJ5, 19: AJ1, 20: AJ6, and 21: AJ7. Numbers on the gel indicate the bands excised for sequencing. Am. J. Primatol. 914 / Nakamura et al. 2.0 1.5 Motiepa (M) Balam (B) AcaJungla (A) A5 PC2 (16.8%) 1.0 A1 M7 B3 B2 B1 M6 M8 B4 M2 A7 0.5 A4 0 A3 -0.5 Diversity Analysis of Archaea 16S rDNA by DGGE M1, M3, M4, M5 cluster B5 A6 B6 -1.0 A2 -1.5 -2.0 -1.5 -1.0 -0.5 0 0.5 1.0 1.5 PC1 (47.0%) Fig. 3. Principal coordinate analysis based on presence and absence of DGGE bands in Desulfovibrionales 16S rDNA profiles. The x- and y-axes denote the eigenvectors of the first dimension, PC1, and second dimension, PC2, respectively. Percentages in parentheses indicate the percentage of total variance accounted by thatprincipal component. PC1 accounts for 47% of total variance and is the most important component in accounting for the clustering spread of samples. performed on the presence/absence of bands within these DGGE profiles confirmed this result, and showed weak separation between the two wild groups but a clear differentiation from the captive group (Fig. 3). Interestingly, two of the Balam wild group (B5 and B6) differed from all other wild monkeys, and observation records confirmed that these females were pregnant at the time of sample collection (S. Van Belle, personal communication). Conversely, two of the captive group (AJ6 and AJ7) were more similar to the wild groups than their other captive counterparts. Of these, one was an infant (AJ7; 2 month old) suffering from diarrhea at the time of fecal collection and had received antibiotic medication as well as a probiotic supplement. The sequences amplified from the Desulfovibrionales DGGE bands (Fig. 2A) were compared with those archived in the Greengenes database using BLAST search. Closest relatives and percentage identities for each band are shown in Table II. Data revealed amplification of bacteria belonging to the family Lachnospiraceae, owing to high sequence similarity in the Desulfovibrionales-specific 16S rDNA primer binding site. Figure 2B shows taxonomic group assignment for major DGGE bands. There was a clear difference in the composition of the family Lachnospiraceae between the wild and captive populations. Detection of Roseburia intestinalis and Lachnospira pectinoschiza was characteristic of the wild and captive populations, respectively. Overall diversity of Lachnospiraceae species was higher in Am. J. Primatol. the wild than the captive populations. Similarly, most wild animals showed two bands related to Desulfovibrio species, whereas only one of those was dominant in the captive animals. The banding pattern of the infant captive monkey AJ7 was distinct from those of any other wild or captive animal. Sequences of the two dominant bands characteristic of this sample were associated with Eubacteriaceae and D. piger. The DGGE banding patterns were highly similar within each wild and captive group, and distinct between groups (Fig. 4). Most animals of the Balam population did not possess Archaeal species that were detectable by this PCR-DGGE method. One of the sequences obtained from the Motiepa population (A1) grouped with a putative novel archaeal order [Wright et al., 2004], represented by uncultured members commonly found in the rumen or animal intestinal tract. The sequence from the dominant band in the captive population (A3) was highly similar to Methanosphaera stadtmanae (97% bootstrapping support). Small Clone Library Analysis Clones randomly selected from each group showed that the target Archaea 16S rDNA sequence was successfully cloned in all ten clones originating from Motiepa, eight clones from AcaJungla, and two from Balam. Results of a phylogenetic analysis of these sequences are shown in Figure 5. Nine of the ten sequences from Motiepa formed a group clustering with the putative new archaeal order [Wright et al., 2004]. Another cluster was formed within this archaeal group, which consisted of sequences from each of the wild and captive groups. Two sequences from the captive group were nearly identical (499%) to Methanobrevibacter smithii and five others were closely related (97%) to M. stadtmanae. Quantitative Real-Time PCR The abundance of methanogens and SRB was estimated by quantitative real-time PCR (qPCR) targeting the functional genes mcrA and dsrA, respectively. The copy numbers of mcrA in 10 ng of the fecal DNA were below the detection limit of 102 in all samples. The dsrA gene abundance ranged from 102–105 copies per 10 ng DNA. The data were normalized and represented as percentage of total Bacteria (Table III). The statistical tests indicated no significant effects of group, age, sex, or sampling time. DISCUSSION In this study, we present evidence of clear groupspecific variation in the intestinal hydrogenotrophic microbial community between wild and captive black Gut Microbiota of Black Howler Monkeys / 915 TABLE II. Closest Identified Taxa of 16S rDNA Sequences Obtained From Desulfovibrionales-Enriched DGGE Band number 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 Closest identified taxon GenBank accession number % identity Roseburia intestinalis str. L1-8151 Low quality sequence Clostridium sp. ASF502 str. ASF 502 Lachnospiraceae genomosp. C1 Clostridium sp. ASF502 str. ASF 502 Lachnospiraceae genomosp. C1 Clostridium sp. ASF502 str. ASF 502 Lachnospiraceae genomosp. C1 Desulfovibrio sp. str. UNSW3caefatS Desulfovibrio sp. str. UNSW3caefatS Roseburia intestinalis str. L1-8151 Low quality sequence Clostridium sp. ASF502 str. ASF 502 Lachnospiraceae genomosp. C1 Lachnospiraceae genomosp. C1 Syntrophococcus sucromutans str. DSM 3224 Low quality sequence Desulfovibrio sp. str. UNSW3caefatS Lachnospira pectinoschiza Clostridium sp. ASF502 str. ASF 502 Lachnospiraceae genomosp. C1 Clostridium sp. ASF502 str. ASF 502 Lachnospiraceae genomosp. C1 Desulfovibrio sp. str. UNSW3caefatS Desulfovibrio sp. str. UNSW3caefatS Eubacterium callanderi str. DSM 3662 Pseudoramibacter alactolyticus str. 23263T Desulfovibrio piger str. ATCC29098 Desulfovibrio sp. str. UNSW3caefatS Desulfovibrio piger str. ATCC29098 Low quality sequence Clostridium sp. ASF502 str. ASF 502 Clostridium bolteae str. 16351 Lachnospiraceae genomosp. C1 AJ312386 99.42 AF157053 AY278618 AF157053 AY278618 AF157053 AY278618 AF056091 AF056091 AJ312386 94.74 AF157053 AY278618 AY278618 Y18191 97.08 94.29 91.75 91.79 97.71 97.11 92.57 AF056091 AY169414 AF157053 AY278618 AF157053 AY278618 AF056091 AF056091 X96961 AB036759 AF192152 AF056091 AF192152 91.75 95.95 98.25 AF157053 AJ508452 AY278618 95.38 94.22 91.75 92.27 94.22 95.88 91.75 Fig. 4. Archaea 16S rDNA-specific DGGE profiles of wild (Motiepa and Balam) and captive groups. 1: M1, 2: M2, 3: M3, 4: M4, 5: M5, 6: M6, 7: M7, 8: M8, 9: B1, 10: B2, 11: B3, 12: B4, 13: B5, 14: B6, 15: AJ2, 16: AJ3, 17: AJ4, 18: AJ5, 19: AJ1, 20: AJ6, and 21: AJ7. Numbers (A1–A3) on the gel indicate the bands excised for sequencing. Am. J. Primatol. 916 / Nakamura et al. Fig. 5. Neighbor-joining tree showing phylogenetic relationship of cloned Archaea 16S rDNA sequences. Methanopyrus kandleri was used as the outgroup. Sequences from this study are represented with letters indicating group (M, Motiepa; B, Balam; A, AcaJungla) and clone numbers (1–10). The bootstrap values on nodes are percentage confidence levels from 1,000 replications of resampling the alignments. howler monkeys. Our results indicate that later environmental and/or dietary changes by the host are likely to facilitate exposure to new or additional microbes, and therefore could exert a greater influence on the composition of intestinal hydrogenotrophs in black howler monkeys than the host genetic or early environmental effects. Alternatively, use of antibiotics to treat captive animals can result in perturbation of the gut microbiota and subsequent loss of specific archaeal and bacterial lineages. Although all captive monkeys from this group had been treated at various times with antibiotics (e.g., enrofloxacin for intestinal infections) and a commercial probiotic preparation during the period from at least a month before fecal collection for this study, only the captive monkey AJ7 was being treated at the time of collections. It has been shown that antibiotic treatment can cause a pervasive disturbance in the microbial community composition, which requires more than 4 weeks to recover to the pretreatment state [Dethlefsen et al., 2008]. Am. J. Primatol. The Desulfovibrionales-enriched DGGE indicated reduced microbial diversity in the captive animals compared with their wild counterparts. Most of the 16S rDNA sequences highly similar to Desulfovibrio spp. had relatively low percentage identity with sequences in the database, which may indicate that these howlers possess particular SRB that have not been characterized so far. The 16S rDNA sequences from Lachnospiraceae were detected in addition to Desulfovibrionales, owing to their high similarity to the Desulfovibrionales-specific primer sequences. The family Lachnospiraceae includes genera Butyrivibrio, Lachnospira, and Roseburia, which are normal inhabitants of the rumen and gastrointestinal tract of animals [Cotta & Forster, 2006]. R. intestinalis-related sequence was detected only from the wild groups. R. intestinalis is a butyrate-producing microbe that belongs to the clostridial cluster XIVa [Duncan et al., 2002b], and has an absolute requirement for acetate as a cosubstrate of butyrate production [Duncan et al., Gut Microbiota of Black Howler Monkeys / 917 TABLE III. Abundance of dsr Gene Copies Normalized to Bacteria 16S rDNA Copies (%) Group Motiepa Motiepa Motiepa Motiepa Motiepa Motiepa Motiepa Motiepa Balam Balam Balam Balam Balam Balam Individual Age Sex May June July M1 M2 M3 M4 M5 M6 M7 M8 B1 B2 B3 B4 B5 B6 Juvenile Juvenile Subadult Adult Adult Adult Adult Adult Juvenile Juvenile Adult Adult Adult Adult Male Male Male Male Male Male Female Female Male Female Male Male Female Female 0.12 0.31 0.12 0.23 0.42 0.57 1.73 0.11 0.27 0.11 0.09 0.05 NDb 1.39 2.71 1.27 1.61 1.15 0.11 0.74 0.28 0.39 0.98 NDa 1.64 0.18 0.22 1.01 6.39 1.11 1.19 0.10 0.94 1.12 0.57 2.14 0.72 0.21 0.20 0.14 0.34 0.29 Sampled one time AcaJungla AcaJungla AcaJungla AcaJungla AcaJungla AcaJungla AcaJungla AJ1 AJ2 AJ3 AJ4 AJ5 AJ6 AJ7 Juvenile Juvenile Juvenile Juvenile Juvenile Juvenile Juvenile Female Male Male Male Male Female Female 0.31 0.40 0.21 12.56 NDc 0.03 2.03 a Sample not available. Not calculated owing to low copy number of Bacteria 16S rDNA. c Not calculated owing to a dsrA copy number below detection limit. b 2002a]. Acetate is reported to be the major VFA produced by cecal contents of wild howler monkeys (93.7% of total VFA) [Milton & McBee, 1983]. Crossfeeding between the acetate-producing bifidobacteria and R. intestinalis has been suggested to contribute to the bifidogenic/butyrogenic effect observed after ingestion of prebiotics in human [Falony et al., 2006]. Further studies would be needed to determine the extent to which such interactions occur in the intestinal microbiota of wild black howler monkeys, and their quantitative contribution to butyrate production would be difficult to determine; however, the probability of this interaction occurring is high given the high levels of acetate produced in this species. The microbiota of captive animals was characterized by the dominance of L. pectinoschiza-like and M. stadtmanae sequences. L. pectinoschiza, which was isolated from the pig intestine [Cornick et al., 1994], is a pectinophile, a type of bacteria that utilizes pectin and a few related compounds as substrates. Pectin fermentation results in the production of acetate, formate, ethanol, methanol, and CO2. Syntrophy between pectinophiles and methanolutilizing bacteria has been observed during coculture growth on pectin [Rode et al., 1981]. M. stadtmanae utilizes only methanol and H2 for methanogenesis [Fricke et al., 2006; Miller & Wolin, 1985]. Together, these observations are consistent with increased pectin fermentation and potential cross-feeding between pectinophiles and methanogens in the intestinal microbiota of captive howler monkeys. Increased pectin fermentation may be expected, because the captive animals analyzed in this study were fed fruits daily at the rehabilitation center. The real-time PCR results indicated higher abundance of SRB than methanogens in all howler monkeys examined in this study. Relatively large inter- and intraindividual variations were observed in SRB abundances; however, we could not identify the factors that might have caused this variation. The level of methanogens in the Balam group was below the detection limit by both real-time PCR and PCR-DGGE methods. Although methanogens were not detected, the abundance of SRB in the Balam group was not significantly different from that in other groups. These results may indicate the presence of an alternative hydrogenotrophic pathway, such as reductive acetogenesis. Reductive acetogenesis is regarded as a relatively minor hydrogenotrophic pathway in the human colon. However, acetogens may predominate in nonmethanogenic human subjects, and the numbers of acetogens have a negative correlation with those of methanogens [Bernalier et al., 1996]. In a companion study, VFA analysis carried out on these howler monkey fecal samples showed that the molar proportions of VFA were 76.4% acetate, 14.3% propionate, 5.8% butyrate, 0.8% isobutyrate, 1.3% iso-valerate, and 1.4% valerate, consistent with an active fiber fermentation but do not provide any evidence for alternative H2 disposal pathways [Amato, 2011, unpublished]. The high similarity of hydrogenotrophic microbial profiles among captive animals was somewhat surprising. These animals were collected from various geographical areas in southern Mexico and were kept together in small enclosures at the rehabilitation center. The similarity in their microbiota likely reflects the shift to a more simplified microbial community composition owing to the dietary and/or environmental conditions under captivity. M. smithii and M. stadtmanae are commonly found in humans that harbor methanogens and are considered specific to human microbiota. However, both M. smithii and M. stadtmanae have been detected in sheep, cattle [Wright et al., 2004, 2007], and pigs [Ufnar et al., 2007]. Despite these unexpected findings in captive howler monkeys, differences between a wild and captive diet would certainly affect intestinal fermentation patterns. Staple foods consumed by these wild howlers, such as leaves, primarily of the species Poulsenia armata (Moraceae), and fruits, primarily Ficus spp. (Moraceae) [Amato, 2011, unpublished], contain considerable cell wall material (30–55% dry weight) [Milton et al., 1980]. On the other hand, fruits and vegetables fed routinely to captive animals may be considerably lower in dry matter and fiber and, thus, together with commercial monkey chow Am. J. Primatol. 918 / Nakamura et al. commonly added to captive monkey diets, contribute to a more readily digestible diet than that consumed by wild monkeys [Crissey & Pribyl, 1997]. Of interest was the difference in the Desulfovibrionales 16S rDNA banding profiles used for the PCA (Fig. 3) that showed that two pregnant females (B5 and B6) had community profiles different from all other wild howler monkeys. This difference in fecal community profile may reflect a change in foraging behavior by pregnant females in order to increase intake of protein and energy. An 8-year study of lactating female howler monkeys (Alouatta palliata) estimated that they consumed significantly more energy (41%) and protein (118%) compared with nonlacating females [Serio-Silva et al., 1999]. Environmental conditions and transmission of methanogens among individuals are also known to have important influences on the establishment of a methanogenic microbiota. In the cohabitation experiments of methanogenic and nonmethanogenic adult rats by Florin et al. , the methanogenic trait was transferred to nonmethanogenic adult rats. Once a methanogenic microbiota was established, all animals retained this trait over the 2-year study period, indicating the remarkable stability of methanogens [Florin et al., 2000]. Furthermore, Bond et al.  observed an unusually high incidence of methane producers among institutionalized children living together in closed units for a long period of time. Only a few studies have examined the impact of captivity on the primate intestinal microbiota [Benno et al., 1987; Fujita & Kageyama, 2007; Uenishi et al., 2007], all of which showed clear differences between wild and captive populations. Our study clearly indicated the impact of captivity on the hydrogenotrophic microbiota of howler monkeys. Hydrogenotrophic activity has important implications for host health [Nakamura et al., 2010]. In our study, one infant animal in captivity (AJ7, estimated to be 2 months old) that harbored a unique hydrogenotrophic microbiota dissimilar from any wild or other captive animal was experiencing diarrhea and being treated with the antibiotic enrofloxacin and a commercial probiotic preparation at the time of sample collection, indicating a dramatic impact of diarrhea, probiotics, and/or antibiotic treatment on the hydrogenotrophic microbiota. Thus, analysis of hydrogenotrophic microbiota may provide insights into the intestinal metabolism and health of captive primates, and thereby contribute to the development of dietary or environmental strategies to improve well being. Further studies are needed to sort out the multifactorial interactions among host genetic, environmental, and dietary influences on the composition of the microbiota of wild and captive nonhuman primates. Such studies may provide insights into the unanswered questions regarding the variation of human methanogenic microbiota observed among Am. J. Primatol. individuals of differential geographic origin [Levitt et al., 2006] or genetic relatedness [Bond et al., 1971; Hackstein et al., 1995]. ACKNOWLEDGMENTS This work was supported by Beckman, Tinker, and PEEC grants awarded to K.R. Amato at UIUC. N. Nakamura was supported in part by the Agricultural Experiment Station at UIUC. The authors thank Dr. Peiying Hong for performing the PCA, Dr. Franck Carbonero for constructive comments, and Ms. Ann Benefiel for editorial assistance. P.A.G. acknowledges Sara, Jenni, and Chrissie for their continued support. We thank Dr. Sarie Van Belle, UIUC, for kindly providing historical data on the two pregnant females and Dr. Salomon Gonzalez, President of Acajungla, for allowing us to collect samples from the captive howler monkeys housed in their facilities. REFERENCES Amann RI, Ludwig W, Schleifer KH. 1995. Phylogenetic identification and in situ detection of individual microbial cells without cultivation. Microbiology Reviews 59:143–169. Benno Y, Itoh K, Miyao Y, Mitsuoka T. 1987. Comparison of fecal microflora between wild Japanese monkeys in a snowy area and laboratory-reared Japanese monkeys. Nippon Juigaku Zasshi 49:1059–1064. Bernalier A, Lelait M, Rochert V, Grivet P, Gibson GR, Durand M. 1996. Acetogenesis from H2 and CO2 by methaneand non-methane-producing human colonic bacterial communities. FEMS Microbiology Ecology 19:193–202. Bond JHJ, Engel RR, Levitt MD. 1971. Factors influencing pulmonary methane excretion in man. An indirect method of studying the in situ metabolism of the methane-producing colonic bacteria. Journal of Experimental Medicine 133:572–588. Cornick NA, Jensen NS, Stahl DA, Hartman PA, Allison MJ. 1994. Lachnospira pectinoschiza sp. nov., an anaerobic pectinophile from the pig intestine. International Journal of Systematic Bacteriology 44:87–93. Cotta M, Forster R. 2006. The family Lachnospiraceae, including the genera Butyrivibrio, Lachnospira and Roseburia. In: Dworkin M, Falkow S, Rosenberg E, Schleifer K-H, Stackebrandt E, editors. The Prokaryotes, 3rd ed. New York: Springer. p 1002–1021. Crissey SD, Pribyl LS. 1997. Utilizing wild foraging ecology information to provide captive primates with an appropriate diet. Proceedings of the Nutrition Society 56:1083–1094. Daly K, Sharp RJ, McCarthy AJ. 2000. Development of oligonucleotide probes and PCR primers for detecting phylogenetic subgroups of sulfate-reducing bacteria. Microbiology 146:1693–1705. Dar SA, Kuenen JG, Muyzer G. 2005. Nested PCR-denaturing gradient gel electrophoresis approach to determine the diversity of sulfate-reducing bacteria in complex microbial communities. Applied and Environmental Microbiology 71: 2325–2330. Denman SE, Tomkins NW, McSweeney CS. 2007. Quantitation and diversity analysis of ruminal methanogenic populations in response to the antimethanogenic compound bromochloromethane. FEMS Microbiology Ecology 62:313–322. DeSantis TZ, Hugenholtz P, Larsen N, Rojas M, Brodie EL, Keller K, Huber T, Dalevi D, Hu P, Andersen GL. 2006. Greengenes, a chimera-checked 16S rRNA gene database Gut Microbiota of Black Howler Monkeys / 919 and workbench compatible with ARB. Applied and Environmental Microbiology 72:5069–5072. Dethlefsen L, Huse S, Sogin ML, Relman DA. 2008. The pervasive effects of an antibiotic on the human gut microbiota, as revealed by deep 16S rRNA sequencing. PLoS Biology 6:e280. Duncan SH, Barcenilla A, Stewart CS, Pryde SE, Flint HJ. 2002a. Acetate utilization and butyryl coenzyme A (CoA): acetate-CoA transferase in butyrate-producing bacteria from the human large intestine. Applied and Environmental Microbiology 68:5186–5190. Duncan SH, Hold GL, Barcenilla A, Stewart CS, Flint HJ. 2002b. Roseburia intestinalis sp. nov., a novel saccharolytic, butyrate-producing bacterium from human faeces. International Journal of Systematic and Evolutionary Microbiology 52:1615–1620. Eisenberg JF, Muckenhirn NA, Rundran R. 1972. The relation between ecology and social structure in primates. Science 176:863–874. Falony G, Vlachou A, Verbrugghe K, De Vuyst L. 2006. Crossfeeding between Bifidobacterium longum BB536 and acetate-converting, butyrate-producing colon bacteria during growth on oligofructose. Applied and Environmental Microbiology 72:7835–7841. Florin TH, Zhu G, Kirk KM, Martin NG. 2000. Shared and unique environmental factors determine the ecology of methanogens in humans and rats. American Journal of Gastroenterology 95:2872–2879. Fricke WF, Seedorf H, Henne A, Kruer M, Liesegang H, Hedderich R, Gottschalk G, Thauer RK. 2006. The genome sequence of Methanosphaera stadtmanae reveals why this human intestinal archaeon is restricted to methanol and H2 for methane formation and ATP synthesis. Journal of Bacteriology 188:642–658. Fujita S, Kageyama T. 2007. Polymerase chain reaction detection of Clostridium perfringens in feces from captive and wild chimpanzees, Pan troglodytes. Journal of Medical Primatology 36:25–32. Hackstein JH, Van Alen TA, Op Den Camp H, Smits A, Mariman E. 1995. Intestinal methanogenesis in primates—a genetic and evolutionary approach. Dtsch Tierarztl Wochenschr 102:152–154. Janse I, Bok J, Zwart G. 2004. A simple remedy against artifactual double bands in denaturing gradient gel electrophoresis. Journal of Microbiological Methods 57:279–281. Kondo R, Nedwell DB, Purdy KJ, de Queiroz Silva S. 2004. Detection and enumeration of sulphate-reducing bacteria in estuarine sediments by competitive PCR. Geomicrobiology Journal 21:145–157. Lepp PW, Brinig MM, Ouverney CC, Palm K, Armitage GC, Relman DA. 2004. Methanogenic Archaea and human periodontal disease. Proceedings of the National Academy of Science of the United States of America 101:6176–6181. Levitt MD, Furne JK, Kuskowski M, Ruddy J. 2006. Stability of human methanogenic flora over 35 years and a review of insights obtained from breath methane measurements. Clinical Gastroenterology and Hepatology 4:123–129. Miller TL, Wolin MJ. 1985. Methanosphaera stadtmaniae gen. nov., sp. nov.: a species that forms methane by reducing methanol with hydrogen. Archives of Microbiology 141: 116–122. Milton K, McBee RH. 1983. Rates of fermentative digestion in the howler monkey, Alouatta palliata (primates: ceboidea). Comparative Biochemistry and Physiology—Part A: Comparative Physiology 74:29–31. Milton K, Soest PJV, Robertson JB. 1980. Digestive efficiencies of wild howler monkeys. Physiology and Zoology 53: 402–409. Muyzer G, de Waal EC, Uitterlinden AG. 1993. Profiling of complex microbial populations by denaturing gradient gel electrophoresis analysis of polymerase chain reactionamplified genes coding for 16S rRNA. Applied and Environmental Microbiology 59:695–700. Nakamura N, Lin HC, McSweeney CS, Mackie RI, Gaskins HR. 2010. Mechanisms of microbial hydrogen disposal in the human colon and implications for health and disease. Annual Review of Food Science and Technology 1:363–395. Raskin L, Stromley JM, Rittmann BE, Stahl DA. 1994. Groupspecific 16S rRNA hybridization probes to describe natural communities of methanogens. Applied and Environmental Microbiology 60:1232–1240. Rode LM, Genthner BR, Bryant MP. 1981. Syntrophic association by cocultures of the methanol- and CO2-H2-utilizing species Eubacterium limosum and pectin-fermenting Lachnospira multiparus during growth in a pectin medium. Applied and Environmental Microbiology 42:20–22. Saitou N, Nei M. 1987. The neighbor-joining method: a new method for reconstructing phylogenetic trees. Molecular Biology and Evolution 4:406–425. Samuel BS, Gordon JI. 2006. A humanized gnotobiotic mouse model of host-archaeal-bacterial mutualism. Proceedings of the National Academy of Science of the United States of America 103:10011–10016. Serio-Silva JC, Hernández-Salazar LT, Rico-Gray V. 1999. Nutritional composition of the diet of Alouatta palliata mexicana females in different reproductive states. Zoo Biology 18:507–513. Tamura K, Dudley J, Nei M, Kumar S. 2007. MEGA4: molecular evolutionary genetics analysis (MEGA) software version 4.0. Molecular Biology and Evolution 24:1596–1599. Turnbaugh PJ, Ley RE, Mahowald MA, Magrini V, Mardis ER, Gordon JI. 2006. An obesity-associated gut microbiome with increased capacity for energy harvest. Nature 444:1027–1031. Uenishi G, Fujita S, Ohashi G, Kato A, Yamauchi S, Matsuzawa T, Ushida K. 2007. Molecular analyses of the intestinal microbiota of chimpanzees in the wild and in captivity. American Journal of Primatology 69:367–376. Ufnar JA, Wang SY, Ufnar DF, Ellender RD. 2007. Methanobrevibacter ruminantium as an indicator of domesticated-ruminant fecal pollution in surface waters. Applied and Environmental Microbiology 73:7118–7121. Wright AD, Williams AJ, Winder B, Christophersen CT, Rodgers SL, Smith KD. 2004. Molecular diversity of rumen methanogens from sheep in Western Australia. Applied and Environmental Microbiology 70:1263–1270. Wright AD, Auckland CH, Lynn DH. 2007. Molecular diversity of methanogens in feedlot cattle from Ontario and Prince Edward Island, Canada. Applied and Environmental Microbiology 73:4206–4210. Zhang H, DiBaise JK, Zuccolo A, Kudrna D, Braidotti M, Yu Y, Parameswaran P, Crowell MD, Wing R, Rittmann BE, Krajmalnik-Brown R. 2009. Human gut microbiota in obesity and after gastric bypass. Proceedings of the National Academy of Science of the United States of America 106: 2365–2370. Am. J. Primatol.