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Analysis of the hydrogenotrophic microbiota of wild and captive black howler monkeys (Alouatta pigra) in palenque national park Mexico.

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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: hgaskins@illinois.edu
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
[47]
[1]
[10]
[10]
[36]
[36]
[7]
[7]
[11]
[11]
[29]
[29]
[14]
[57]
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. [2000], 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. [1971]
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.
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