Research Article

Characterization of Bacterial Microbiota in the Gastrointestinal Tract (GIT) of Buffaloes Using PCR-Based Analysis

Phoebe Lyndia Tolentino Llantada1,2*, Midori Umekawa2, Shuichi Karita2

1Philippine Carabao Center National Headquarters and Genepool, Philippines; 2Graduate School of Bioresources, Mie University, 1577 Kurimamachiya-cho, Tsu city, Mie 514-8507, Japan.

Received | September 28, 2023; Accepted | January 24, 2024; Published | February 07, 2024

*Correspondence | Phoebe Lyndia Tolentino Llantada, Philippine Carabao Center National Headquarters and Genepool, Philippines; Email: llantadaphoebe@gmail.com

Citation | Llantada PLT, Umekawa M, Karita S (2024). Characterization of bacterial microbiota in the gastrointestinal tract (GIT) of buffaloes using PCR-based analysis. Adv. Anim. Vet. Sci., 12(3):479-489.

DOI | https://dx.doi.org/10.17582/journal.aavs/2024/12.3.479.489

ISSN (Online) | 2307-8316

Copyright: 2024 by the authors. Licensee ResearchersLinks Ltd, England, UK.

This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https://creativecommons.org/licenses/by/4.0/).

INTRODUCTION

In tropical regions like the Philippines, buffalo (Bubalus bubalis) plays an important role in the livestock sector of the country. This animal serves as a source of milk and meat production as well of draft power and hide. Since buffalo can adapt to extreme climatic conditions and tolerate tropical diseases, it requires less management. Moreover, buffalo can feed on locally available forages and farm by-products, making it the most sustainable livestock in the country. As of July 1, 2017, the current population of buffalo (both backyard and commercial farm) in the country is 2.88 million heads (PSA, 2017).

In the Philippines, there are two major types of buffalo, the swamp buffalo and the riverine buffalo (Figure 1). The swamp buffalo commonly known as the Philippine carabao is usually utilized for draft power and meat production whereas the riverine buffalo or the dairy type is mainly raised for milk and meat production. The latter has an average milk production of 8-10 kg in a 305 days lactation period (Sarabia et al., 2009), thus a promising breed to improve the dairy industry of the country to meet the increasing demand for milk and meat of the growing population.

 

The unabated increase in the population growth of the country, which demands more milk and meat offers a great opportunity to small hold dairy farmers as a promising source of income. This scenario made the dairy sector of the government to do several actions such as the implementation of upgrading programs for the swamp buffalo wherein crossbreeding and backcrossing schemes were done in villages to produce Philippine dairy buffaloes. Another is the establishment of forage pastures. However, the efforts in the genetic improvement of the buffalo and pasture establishment were insufficient to address the challenge in increasing the ruminants’ productivity. This is because the major constraint relies on the nutrition aspect especially under the small hold ruminant production system. The animals were predominantly fed on locally available forages and farm by-products which are characterized as low in energy and protein contents, fibrous, and highly lignified which restrict the feed intake and nutrients digestibility of the animals. Hence, the genetic potential of the animals is not fully expressed, resulting in low production of the animals.

Several studies on dietary manipulations have been carried out in the past, suggesting that buffalo have higher digestibility of fibrous materials as compared to cattle (Norton et al., 1979; Devendra, 1983; Moran et al., 1983; Katiyar and Bisth, 1988; Wanapat et al., 1994). Therefore, it is necessary to investigate the complexity and the role of microorganisms in the rumen (Firkins and Yu, 2006) and other parts of the digestive tract which determine the efficiency of the GIT functions for fiber digestion. In the feeding of ruminants like the buffalo, the feed consumed is first exposed to rumen fermentation by the action of the microbes before gastric and intestinal digestion. Then, dietary polysaccharides are degraded into volatile fatty acids (VFA’s) such as acetate, butyrate, and propionate which serve as sources of energy both for the microbes and the animal (Madigan et al., 2000). The complex rumen digestion process makes the manipulation of microbial processes a challenge, especially in the rumen. Thus, the application of molecular biology techniques and bioinformatics in conducting a comprehensive survey on the microbial diversity in the rumen and other parts of the digestive tract (Morgavi et al., 2013) of ruminants is necessary to have a better understanding of the biological function of the whole gastrointestinal tract ecosystem.

This study was conducted to profile the bacterial communities across the dairy buffaloes’ gut using PCR-DGGE (Muyzer et al., 1993) analysis by PCR with species specific primer sets.

MATERIALS AND METHODS

All the experimental procedures, including experimental animal maintenance and sample collection, were conducted following the guidelines of the ethical committee at the Philippine Carabao Center with the research code AN19004-RC.

Study location

Sample collection and DNA extraction were carried out in the Philippines whereas sample analysis was carried out in Japan.

Animals

Two buffaloes, 35-month-old healthy male island-born dairy types each with an average body weight of 464 + 32 kg, were used in the study. The animals were raised for 6 months in Gene Pool Farm, Philippine Carabao Center National Headquarters, Science City of Munoz, Nueva Ecija, following the standard dairy buffalo production management practices.

Feeding management before slaughter

The animals were kept in complete confinement, provided with the same diet of rice straw, freshly chopped grasses supplemented with commercial grower concentrate with 18 % CP, and were sometimes given with urea-treated rice straw. The ration was estimated to provide the necessary amount of nutrients needed for the animal’s growth and maintenance. Clean drinking water was also provided ad libitum; the animals were reared and maintained until their target slaughter weight.

Slaughter and gut sample collection

The animals were slaughtered at the Animal Products Development Center (APDC) of the Bureau of Animal Industry (BAI) following the standard procedure under the Humane Slaughter Guidelines of National Meat Inspection Services (NMIS). Twenty grams of fresh luminal samples were collected in all the sites within the three gut regions as follows, foregut: the rumen, reticulum, omasum, and abomasum, midgut: Duodenum, jejunum, and ileum and hindgut: Cecum, colon, and rectum (Figure 2). Three types of samples were collected in the rumen: rumen fluid, rumen digesta, and rumen mucosa. Sterilized gauze was used to filter the rumen samples to separate the liquid and solid samples (fiber-adherent). Aside from the reticulum digesta, reticulum mucosa was also collected. Samples were collected in the intestine from the duodenum (beginning of the midgut) through the rectum (end of the hindgut). Samples from the different sites were thoroughly mixed before placed in centrifuge tubes. Three replicates in each site and sample type were collected, a total of 78 samples were immediately frozen in liquid nitrogen, and transported to the laboratory for genomic DNA extraction.

 

Genomic DNA extraction

The samples collected from the rumen and reticulum mucosa were scraped to remove the attached food particles and were rinsed three times with sterilized phosphate-buffered saline (pH 7.0) before extraction. Bacterial DNA was extracted from the samples using the QIAampTM Fast DNA Stool Mini Kit (Qiagen, Valencia, CA, USA). A portion of the DNA extracted from each site of collection and sample type were pooled; samples were stored at -20°C until further analysis was done.

PCR and DGGE analysis

The hyper-variable V3 region of the 16S rDNA gene was amplified from all samples using the primers 357F and 517R for the confirmation of the target DNA and 357GCF and 517R (Table 1) for DGGE analysis. The forward primer contains a GC clamp at the end to prevent the dissociation of DNA strands (Yu and Morrison, 2004). The PCR reaction cycle was performed with Applied Biosystem 2720 TM thermal cycler (Life Technology). A total of 50 µl PCR reaction mixture contained 2.5 µl template DNA, 1x Ex Taq reaction buffer, 4 µl (2.5 mM each) deoxynucleotide triphosphate (dNTP mixture), 0.25-unit Ex Taq DNA polymerase and 0.5 g l-1 bovine serum albumin (BSA). Touchdown PCR was performed to prevent the production of spurious products. The reaction cycle was set for amplification with an initial denaturation at 94 °C for 4 min followed by denaturation for 30 s at 94 °C, 30 s at 61 °C and a decrease of 0.5 °C per cycle, 30 s at 72 °C (10 cycles), 30 s at 94 °C for denaturation, 30 s at 56 °C, 30 s at 72 °C (35 cycles), and 10 min at 72 °C (Lodge- Ivey et al., 2009). The PCR products were analyzed using electrophoresis on 1.5% agarose gel. DGGE was carried out using the Bio-Rad D-CodeTM system (Bio-Rad, Hercules, CA, USA) with 16 x 16 cm glass plates separated by 1 mm spacers. An equal volume of PCR product and dye mix was loaded in each well for PCR amplicons separation. The 16S rDNA gene products were resolved at 10% acrylamide gel with 30-60% linear denaturant in 1x Tris acetate-EDTA buffer. DGGE was conducted at 60 °C for 14 h at 100 V. After electrophoresis, the gel was stained with SYBR Green and the image was captured using an image analyzer (Printgraph; ATTO, Tokyo, Japan).

The PCR reaction mixture (25 mL) contained 12.5 mL of Amplitaq, 0.5 mmol/L of each primer, and 1 mL of the template. The PCR cycle consisted of an initial denaturation at 94 °C for 4 min, followed by denaturation for 30 s at 94°C, 30 s at 61°C and a decrease of 0.5 °C per cycle, 30 s at 72 °C (10 cycles), 30 s at 94 °C for denaturation, 30 s at 56 °C, 30 s at 72 °C (35 cycles), and 10 min at 72 °C (Lodge-Ivey et al., 2009).

Clone library construction and DNA sequencing

Several predominant bands were chosen in the DGGE gel, excised and preserved overnight with 20 µl Milli Q water at 4 °C. One microliter of the eluted DNA was amplified to obtain a clear, single target band using forward primer without GC clamp (Table 1). Two cloning techniques were performed in the experiment. In-vivo cloning was carried out for samples from Buffalo 1, whereas TA (Thymine and Adenine) cloning was conducted for samples from Buffalo 2. The amplified DNA (using the primers listed in Table 2) of samples from Buffalo 1 was ligated using pUC 19

 

Table 1: Primers used in DNA amplification prior to DGGE analysis.

Name	Sequence (5' → 3')	Product size (bp)	Reference
357F	CCTACGGGAGGCAGCAG	194	Tajima et al., 2001
357 GCF	CGCCCGCCGCGCGCGGCGGGCGG-GGCGGGGGCACGGGGGGcctacgggaggcagcag
517R	ATTACCGCGGCTGCTGG

 

Table 2: Primers used in the amplification of eluted DNA samples from DGGE gel prior to in-vivo cloning.

Name	Sequence (5' → 3')
pUC_357F	GCCAGTGAATTCGAGCTCGGTACCCCCTACGGGAGGCAGCAG
pUC_517R	TGCAGGTCGACTCTAGAGGATCCCCATTACCGCGGCTGCTGG

 

Table 3: Primers used in the amplification of pUC19 prior to in-vivo cloning.

Name	Sequence (5' → 3')
iVECpUC19_Top	GGGTACCGAGCTCGAATTCA
iVECpUC19_Bot	GGGGATCCTCTAGAGTCGACCT

 

(amplified using primers in Table 3 before cloning) and PCR 2.1 (Invitrogen, Carlsbad, CA) for samples from Buffalo 2 followed by transformation to iVEC 3 (National Institute of Genetics, Japan) and Escherichia coli JM109 (Toyobo, Kyoto, Japan) respectively. Ampicillin was added in the Luria Bertani (LB) media plate to screened positive clones for the samples from Buffalo 1, while for Buffalo 2, 50 µg/ mL ampicillin and X-gal were used. Five positive transformants (white-colored colony) were randomly selected from each target band. Colonies from Buffalo 1 samples were subjected to colony PCR to confirm if the plasmid (pUC19) contained the insert (bacterial DNA). Samples were sequenced with M13 forward and M13 reverse primer (Takara, Otsu, Japan) using Bigdye Terminator v3.1 protocol. Sequencing reactions were run on ABI 3130. Homology search of the amplified DNA sequences was performed with the BLAST program (Atshul et al., 1997) and EZBioCloud (Yoon et al., 2017) for phylum and class classification. Species richness index R= s was performed using Excel (Microsoft, Redmond, Washington, USA), where s is the number of bands in each sample.

Detection of fibrolytic and non-fibrolytic bacteria using species-specific primer

Eleven species-specific primer sets (Table 4) were used to amplify the 16S rDNA of Clostridium IV, Butyrivibrio fibrosolvens, Fibrobacter succinogenes, Prevotella byantii,

 

 

Table 5: Band clones from the gut sections of Buffalo 1 (B1DGGE1 to B1DGGE11 with their highest percentage of similarity to known sequences in the gene bank.

Sample code	Gut sampling site	Closest relatives (similarity %)	Isolation Source	Accession number
B1DGGE1	Rumen fluid	Unidentified rumen bacterium RFN17 (99)	Bos taurus rumen	LC578747
B1DGGE2	Rumen digesta	Klebsiella sp. strain AAUGM-17 (100)	Insect gut	LC578748
B1DGGE3	Rumen digesta	Uncultured rumen bacterium clone CF376 (99)	Bos taurus rumen	LC578749
B1DGGE4	Omasum	Butyrivibrio sp. CA23 gene (99)	Bovine rumen	LC578750
B1DGGE5	Abomasum	Uncultured bacterium gene, clone: B1_4_20 (99)	Swamp buffalo rumen	LC578751
B1DGGE6	Ileum	Uncultured bacterium clone HY1_a03_2 (99)	Spotted hyena feces	LC578752
B1DGGE7	Ileum	Uncultured bacterium clone AFEL2_aao31g02 (99)	African elephant feces	LC578753
B1DGGE8	Ileum	Paeniclostridium sp. strain HVul.ww1 (100)	Gyps himalayensis	LC578754
B1DGGE9	Rectum	Uncultured bacterium clone Hmb2-50 (99)	Descending colon mucosa of Bos taurus	LC578755
B1DGGE10	Rectum	Uncultured bacterium clone 5-3K21 (99)	Fecal sample from MARC beef Cattle feedlot animal #4281	LC578756
B1DGGE11	Rectum	Uncultured bacterium clone AS1_aao37b02 (97)	Argali sheep feces	LC578757

 

 

Prevotella ruminocola, Ruminococcus albus and Ruminococcus flavefaciens, Anaerovibrio lypolitica, Ruminobacter amylophillus, Selenomonas ruminantium, and Streptococcus bovis. PCR was performed using ExTaq kit (Takara, Otsu, Japan) with Applied Biosystem 2720 thermal cycler (Life Technology). A total of 50 µl PCR reaction mixture for each sample were amplified following the PCR reaction cycle: Initial denaturation at 94 °C for 2 min, followed by thermal cycles consisting of denaturation at 94 °C for 30 s, annealing for 30 s, and extension at 72 °C for 10 min. The annealing temperatures used were based on the annealing temperature of each primer (Table 4). The PCR products were separated into 2% agarose gel with ethidium bromide in electrophoresis (1x TAE buffer) using 100 DNA bp ladder as a molecular marker. The gel images were captured using a gel image analyzer (Printgraph; ATTO, Tokyo, Japan).

RESULTS and Discussion

Bacterial microbiome profile revealed by DGGE

The bacterial diversity (Table 5), DGGE banding patterns, band densities, and the number of bands (Figure 3A, B) varies across the gut sections of the dairy buffaloes. Both animals displayed a higher number of bands (which could represent the gut sections bacterial communities) in the foregut and hindgut as compared in the midgut (Figure 3A, B). PCR-DGGE fingerprints further revealed that samples from the rumen, reticulum, omasum, abomasum, cecum, colon, and rectum shared similar banding patterns. It was further observed that the bacterial population (represented by bands) with higher density in the foregut decreased in the hindgut and the lower density bacterial population in the foregut increased in the hindgut. Moreover, the

 

midgut (duodenum, jejunum, ileum) which was the least diverse segment has distinct bands. Six individual bands were seen in the ileum of Buffalo 1 (Figure 3A) whereas in Buffalo 2, 2 to 3 highly distinct bands were observed in the duodenum, jejunum, and ileum, respectively (Figure 3B).

Homology search using NCBI data base and EZBio cloud

Bacterial communities within the buffalo’s GIT were investigated by sequencing the 16S rRNA of 22 distinct bands in DGGE analysis: 11 bands from Buffalo 1 (Figure 3A) designed as B1DGGE1-B1DGGE11 with accession number LC578747 to LC578757 (Table 5) and 11 bands from Buffalo 2 (Figure 3B) coded as B2DGGE1- B2DGGE11 with accession number LC578758 to LC578768 (Table 6). The randomly selected bands were cloned and a total of 110 clones were sequenced. Table 5 displayed the results of the sequence alignment in the nucleotide collection (nr/nt) database. Most of the sequences were identified as uncultured bacteria (Figures 4A, 5A), therefore, the sequences were rerun using EzBioCloud (Yoon et al., 2017). The results obtained in EZBioCloud showed that almost all the sequences from the samples in both animals were classified as Firmicutes followed by Bacteriodetes then Proteobacteria (Figures 4B and 5B). Phylum Firmicutes was composed of Class Clostridia and Bacilli, bacteria under Phylum Bacteroidetes belongs to Class Bacteroidia, and bacteria from Phylum Proteobacteria were classified under Class Gammaprotobacteria (Figures 4C and 5C). Firmicutes and Bacteriodetes dominated the foregut and hindgut while Proteobacteria was found mostly in the foregut. Interestingly, some of the sequence obtained from the midgut of both animals was unique in terms of closest relatives compared to the other sites of GIT (Tables 5 and 6). Also, the species richness index (Figure 6) showed that the foregut and hindgut have higher species richness compared to the midgut.

 

Table 6: Band clones from the gut sections of Buffalo 2 (B2DGGE1 to B2DGGE11) with their highest percentage of similarity to known sequences in the gene bank.

Sample code	Gut sampling site	Closest relatives (similarity %)	Isolation Source	Accession number
B2DGGE1	Rumen digesta	Saccharofermentans sp. G8 (100)	Bovine rumen	LC578758
B2DGGE2	Rumen digesta	Bacteroidales bacterium P59 (95)	Bovine rumen	LC578759
B2DGGE3	Omasum	Uncultured Proteobacterium clone L2l14UD (100)	Cow rumen	LC578760
B2DGGE4	Rumen fluid	Uncultured rumen bacterium clone BRC57 (99)	Rumen fluid of Bubalus bubalis	LC578761
B2DGGE5	Rumen mucosa	Uncultured bacterium clone 1103200832064 (98)	Bovine rumen fluid fiber adherent microbiome from steer 71	LC578762
B2DGGE6	Abomasum	Uncultured rumen bacterium clone CTRS1B06 (98)	Cow rumen	LC578763
B2DGGE7	Abomasum	Uncultured bacterium clone: I26_4_14 (100)	Cattle rumen	LC578764
B2DGGE8	Duodenum	Paenibacillus xylaniclasticus strain NLG20 (99)	Boselaphus tragocamelus feces	LC578765
B2DGGE9	Cecum	Paenibacillus xylaniclasticus strain NLG20 (99)	Boselaphus tragocamelus feces	LC578766
B2DGGE10	Rectum	Uncultured Bacteroidales bacterium clone CO1 (98)	Cow feces	LC578767
B2DGGE11	Cecum	Uncultured bacterium clone Hda2-82 (99)	Bos taurus descending colon ingesta	LC578768

 

 

Fibrolytic and non-fibrolytic bacteria in the gut of dairy buffaloes

Fibrolytic and nonfibrolytic bacteria were detected across the gut of buffaloes by PCR using species-specific primer sets (Table 4). The results of the experiment showed that Prevotella ruminocola and Selenomonas ruminantium were detected only in the foregut (Table 7). On the other hand, Butyrivibrio fibrosolvens, Fibrobacter succinogenes, Prevotella bryantii, Ruminococcus albus, Anaerovibrio lypolitica, and Streptococcus bovis were located both in the foregut and hindgut. R. flavefaciens, Clostridium IV, and R. amylophilus were observed in the whole GIT. In addition, Table 6 displayed that fibrolytic bacteria were mostly detected in the foregut and hindgut; only a few were observed in the midgut. On the other hand, non-fibrolytic bacteria were mainly found in the foregut, whereas only a few of them were found in the midgut. In the hindgut, non-fibrolytic bacteria were hardly detected.

The symbiotic relationship of microbial flora and the host is essential in balancing the immune response, digestion, and the development of the animal’s GIT. However, the microbial diversity within the ruminants’ gut is understudied. Most studies rely on a few species and only utilize either the ruminal or fecal microbial communities because accessing the ruminants’ microbiota is very difficult. Aside from that, rumen bacteria are difficult to culture, only about 10% to 11% could be cultured (de Oliveira et al., 2013). This is because a vast number of rumen bacteria cannot grow in a single culture medium (Ishaq and Wright, 2014). Thus, in our present study, a culture-independent method was used. The genomic DNA was directly extracted from the samples collected. PCR-DGGE analysis, characterized as low resolution but effective way to identify the dominant microbial community (Sadet et al., 2010) was carried out to profile the bacterial communities present along the gut sections of dairy buffaloes. To detect major fibrolytic and non-fibrolytic bacteria, PCR amplification using species-specific primer sets (Table 4) for 16S rDNA fragments was conducted.

 

Our PCR-DGGE profiles demonstrated that the composition of the microbial community varied along the gut of buffaloes (Figure 3A, B). The data showed that the foregut and the hindgut have higher diversity as compared to the midgut. These results were similar to previous studies on cattle (Romero-Perez et al., 2011), sheep (Neumann and Dehority, 2008; Zeng et al., 2015), and bison (Bergman, 2017). It is known that the microbial community which thrives in the foregut become fermenters providing the primary dietary protein to the ruminant (Bian et al., 2013). On the other hand, the midgut was the least diverse among the three regions due to two major reasons, first is the short retention time of digesta which gives almost no time for microbes to proliferate (Jami and Mizrahi, 2012) and the second one is the low pH level of around 2-4 which was caused by the abomasum (true stomach), gall bladder and the enzymes secreted by the pancreas (Li et al., 2012). The harsh environment created was very detrimental for most microorganisms like the Gram-negative Bacteroidetes; only the Firmicutes which have a thick peptidoglycan Gram-positive cell wall can thrive in this condition (Li et al., 2012). Meanwhile, our DGGE profiles showed that most microbes in buffalo GIT belonged to Firmicutes, Bacteroidetes, and Proteobacteria (Figures 4B and 5B) which is in agreement with the results of previous studies for microbes in other animals GIT (Muyzer, 1999; Lodge-Ivey et al., 2009; Russell, 2002; Zeng et al., 2015). Furthermore, the result of this study revealed that Firmicutes which is a dominant species in the gut and mainly consists of diverse fibrolytic bacterial genera was found largely in the hindgut of Buffalo 1 (Figure 4D) and the foregut of Buffalo 2 (Figure 5D).

In the evaluation of fibrolytic and non-fibrolytic bacteria along the gut sections, the results (Table 7) showed that Prevotella ruminocola and Selenomonas ruminantium were found only in the foregut of both buffaloes. On the other hand, Butyrivibrio fibrosolvens, Fibrobacter succinogenes, Prevotella bryantii, Ruminoccocus albus, and Ruminobacter amylophilus were detected in the midgut and hindgut. Anaerovibrio lypolitica was detected in the foregut and midgut, whereas Clostridium cluster IV and Ruminococcus flavefaciens were observed in the whole GIT (Table 7). It was discussed earlier that the GIT localization of bacteria and their function greatly affect their diversity. Ruminococcus flavefaciens which is a Firmicute have a thick peptidoglycan Gram-positive cell wall which made it survive in harsh conditions like low pH level (Li et al., 2012). On the other hand, Prevotella ruminocola, Ruminococcus albus, and Fibrobacter succinogenes, known to be highly cellulolytic bacteria, were found in the foregut where fermentation occurs (Table 7). Furthermore, both Fibrobacter succinogenes and Selemonas ruminantium are found in the rumen. This is presumably because Fibrobacter succinogenes produce succinate during fiber digestion, while Selemonas ruminantium converts succinate to propionate (Muyzer et al., 1993).

 

 

CONCLUSION ans Recommendations

In conclusion, this study showed that bacterial community composition differs among gut sections but is similar among those within the same region. In addition, it was revealed that most of the fibrolytic bacteria species used in this study were detected in the foregut and hindgut. Since this study only provides a qualitative and semi-quantitative way of analyzing the bacterial composition along with the GIT of dairy buffaloes, the use of quantitative real-time PCR (qPCR) to estimate the population of the major fibrolytic bacteria must be considered. Next Generation sequence (NGS) analysis is another good option because it can provide a more detailed analysis of the bacterial composition, diversity, and function which could lead to the discovery of metabolically important species and potentially novel species that play roles in animal health and productivity.

Acknowledgement

This study is grateful for the generous support from the Japan Ministry of Education, Culture, Sports, Science, and Technology (MEXT) and the Mie University Graduate School of Bioresources. Special thanks are extended to the Philippine Carabao Center for permitting the collection of buffalo samples. The authors also acknowledge and appreciate the valuable support and assistance provided by Prof. Tsutomu Fujihara, the team of the Production System and Nutrition Section (PSNS), and the Gene Pool Farm.

Novelty Statement

The author asserts that this study is the first to conduct a comprehensive, non-culture-based analysis of the bacterial microbiota associated with buffaloes, particularly within different compartments of the gastrointestinal tract (GIT), as observed in buffaloes raised in the Philippines.

Author’s Contribution

PLTL: Conceptualization, data curation, formal analysis, investigation, methodology, validation, writing - original draft, writing review and editing. MU: Supervision, writing review and editing. SK: Conceptualization, formal analysis, funding acquisition, methodology, resources, supervision, validation, writing original draft, writing review and editing.

Conflicts of interest

The authors have declared no conflict of interest.

REFERENCES

Atshul SF, Madden TL, Schaffer AA, Zhang J, Zhang Z, Miller W, Lipman DJ (1997). Gapped BLAST and PSI-BLAST: A new generation of protein database search programs. Nucl. Acids, 25: 3389-3402. https://doi.org/10.1093/nar/25.17.3389

Bergmann GT (2017). Microbial community composition along the digestive tract in forage- and grain-fed bison. BMC Vet. Res., 13(1). https://doi.org/10.1186/s12917-017-1161-x

Bian G, Ma L, Su Y, Zhu W (2013). The microbial community in the feces of the white rhinoceros (Ceratotherium simum) as determined by barcoded pyrosequencing analysis. PLoS One, 8(7). https://doi.org/10.1371/journal.pone.0070103

de Oliveira MNV, Jewell KA, Freitas FS, Benjamin LA, Tótola MR, Borges AC, Suen G (2013). Characterizing the microbiota across the gastrointestinal tract of a Brazilian Nelore steer. Vet. Microbiol., 164(3–4): 307–314. https://doi.org/10.1016/j.vetmic.2013.02.013

Devendra C (1983). The utilization of nutrients, feeding system and nutrient requirements of swamp buffaloes. In: Shimizu H (eds), Proceedings of preconference and symposium 5th world conference in animal production. Tsukuba, Japan; pp. 34.

Fernando SC, Purvis HT, Najar FZ, Sukharnikov LO, Khrebiel CR, Nagaraja TG, De Silva U (2010). Rumen microbial population dynamics during adaptation to a high-grain diet. Appl. Environ. Microbiol. 76(22): 7482-7490. https://doi.org/10.1128/AEM.00388-10

Firkins JL, Yu Z (2006). Characterisation and quantification of microbial populations of the rumen. In: Hvelplund T, Nielson MO, Sejrsen k. (eds), Ruminant physiology: Digestion, metabolism and impact of nutrition on gene expression, immunology and stress. Waginengen, Academic publisher, UK. pp. 19-54. https://doi.org/10.3920/9789086865666_002

Ishaq SL, Wright AD (2014). High-throughput DNA sequencing of the ruminal bacteria from moose (Alces alces) in Vermont, Alaska, and Norway. Microb. Ecol., 68(2): 185–195. https://doi.org/10.1007/s00248-014-0399-0

Jami E, Mizrahi I (2012). Composition and similarity of bovine rumen microbiota across individual animals. PLoS One, 7(3). https://doi.org/10.1371/journal.pone.0033306

Katiyar RC, Bisth GS (1988). Nutrient utilization in murrah buffalo and hariana cattle. A comparative study with oat-hay- based rations. Proc. Second World Buffalo Cong. New Delhi, India; 2: 189-193.

Koike S, Kobayashi Y (2001). Development and use of com- petitive PCR assays for the rumen cellulolytic bacteria: Fibrobacter succinogenes, Ruminococcus albus and Ruminococcus avefaciens. FEMS Microbiol. Lett., 204: 361–366. https://doi.org/10.1111/j.1574-6968.2001.tb10911.x

Li RW, Connor EE, Li C, Baldwin Vi RL, Sparks ME (2012). Characterization of the rumen microbiota of pre-ruminant calves using metagenomic tools. Environ. Microbiol., 14(1): 129–139. https://doi.org/10.1111/j.1462-2920.2011.02543.x

Lodge-Ivey SL, Brownie-Silva J, Horvath MB (2009). Bacterial diversity and fermentation end products in rumen fluid samples collected via oral lavage or rumen cannula. J. Anim. Sci., 87: 2333-2337. https://doi.org/10.2527/jas.2008-1472

Madigan MT, Martinko JM, Parker J (2000). Biology of microorganism. Pentice-Hall International, London. pp. 681-685.

Matsuki T, Watanabe K, Fujimoto J, Takada T, Tanaka R (2004). Use of 16S rRNA gene-targeted group-specific primers for real-time PCR analysis of predominant bacteria in human feces. Appl. Environ. Microbiol., 70(12): 7220-7228. https://doi.org/10.1128/AEM.70.12.7220-7228.2004

Moran JB, Satoto KB, Dawson JE (1983). The utilization of rice straw fed to fed to zebu cattle and swamp buffalo as influenced by alkali treatment and Leucaena supplementation. Austral. J. Agric. Res., 34: 73-84. https://doi.org/10.1071/AR9830073

Morgavi DP, Kelly WJ, Janssen PH, Attwood GT (2013). Rumen microbial (meta) genomics and its application to ruminant production. Animal, 7: 184–201. https://doi.org/10.1017/S1751731112000419

Muyzer G (1999). DGGE/ TGGE a method for identifying genes from natural ecosystems. Curr. Opin. Microbiol., 2: 317-322. https://doi.org/10.1016/S1369-5274(99)80055-1

Muyzer G, de Waal EC, Uitterlinden AG (1993). Profiling of complex microbial populations by denaturing gradient gel electrophoresis analysis of polymerase chain reaction-amplified genes coding for 16S rRNA. Appl. Environ. Microbiol., 59(3): 695–700. https://doi.org/10.1128/aem.59.3.695-700.1993

Neumann LM, Dehority BA (2008). An investigation of the relationship between fecal and rumen bacterial concentrations in sheep. Zoo Biol., 27(2): 100–108. https://doi.org/10.1002/zoo.20166

Norton BW, Moran JB, Nolan JV (1979). Nitrogen metabolism in brahman cross, buffalo, banteng and shorthorn steers fed on low quality roughage. Austral. J. Agric. Res., 30: 341-351. https://doi.org/10.1071/AR9790341

Philippine Statistics Authority (2017). Carabao situation report, January- June 2017.

Romero-Pérez GA, Ominski KH, McAllister TA, Krause DO (2011). Effect of environmental factors and influence of rumen and hindgut biogeography on bacterial communities in steers. Appl. Environ. Microbiol., 77(1): 258–268. https://doi.org/10.1128/AEM.01289-09

Russell JB (2002). Rumen Microbiology and Its Role in Ruminant Nutrition. New York State College of Agriculture and Life Sciences. Department of Microbiology, Cornell University. 121p.

Sadet S, Martin C, Morgavi DP (2010). Bacterial diversity dynamics in rumen epithelium of wethers fed forage and mixed concentrate forage diets. Vet. Microbiol., 146: 98-104. https://doi.org/10.1016/j.vetmic.2010.04.029

Sarabia et al., 2009. The Dairy buffalo production handbook. Science City of Munoz, Nueva Ecija, Philippines: Philippine Carabao Center. 298p.

Tajima K, Aminov RI, Nagamine T, Matsui H, Nakamura M, Benno Y (2001). Diet-dependent shifts in the bacterial population of the rumen revealed with real-time PCR. Appl. Environ. Microbiol., 67: 2766–2774. https://doi.org/10.1128/AEM.67.6.2766-2774.2001

Wanapat M, Sommart K, Wachirapakorn C, Uriyapongson S, Wattanachant C (1994). Recent advances in swamp buffalo nutrition and feeding. In: Wanapat, M., Sommart K. (eds). Proceeding the 1st Asian Buffalo Association Congress. Khon Kaen, Thailand.

Wanapat M, Cherdthong A (2009). Use of real-time PCR technique in studying rumen cellulolytic bacteria population as affected by level of roughage in swamp buffalo. Curr. Microbiol., 58: 294–299. https://doi.org/10.1007/s00284-008-9322-6

Yoon SH, Ha SM, Kwon S, Lim J, Kim Y, Seo H, Chun J (2017). Introducing EzBioCloud: A taxonomically united database of 16S rRNA and whole genome assemblies. Int. J. Syst. Evol. Microbiol., 67: 1613-1617. https://doi.org/10.1099/ijsem.0.001755

Yu Z, Morrison M (2004). Comparisons of different hypervariable regions of rrs genes for use in fingerprinting of microbial communities by PCR-denaturing gradient gel electrophoresis. Appl. Environ. Microbiol., 70(8): 4800–4806. https://doi.org/10.1128/AEM.70.8.4800-4806.2004

Zeng Y, Zeng D, Zhang Y, Ni X, Tang Y, Zhu H, Jing B (2015). Characterization of the cellulolytic bacteria communities along the gastrointestinal tract of Chinese Mongolian sheep by using PCR-DGGE and real-time PCR analysis. World J. Microbiol. Biotechnol., 31(7): 1103–1113. https://doi.org/10.1007/s11274-015-1860-z