1

Techniques of Rumen Fluid Collection

Nirali Pathak

niralipathak@ufl.edu

Fall 2020

CALS Honors Thesis

Department of Animal Science, University of Florida, Gainesville, 32611, FL, USA

College of Agricultural and Life Sciences

Thesis Advisor: Diwakar Vyas, 2250 Shealy Drive, Department of Animal Science, University

of Florida, Gainesville, FL 32611; 352-294-1079 (Phone); 352-392-7652 (Fax);

diwakarvyas@ufl.edu

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ABSTRACT

The objective of this study was to compare fermentation profiles and microbial diversity from

two different rumen sample collection methods. Three ruminally cannulated lactating dairy cows

were used as rumen fluid donors in a 3 × 3 Latin Square design. The treatments were rumen fluid

collected from stomach tube (ST) or from rumen cannula (C). The pH was measured after

collection, and samples were analyzed for volatile fatty acid (VFA), ammonia-N (NH3-N)

concentration, and microbiome composition. Data were analyzed using GLIMMIX procedure of

SAS. Significance was declared at P ≤ 0.05. Rumen pH was greater for ST compared to C (6.88

vs 6.25; P < 0.01). However, NH3-N (15.2 vs 10.6 mg/dL; P = 0.01) and total VFA (121.8 vs

95.5 mM; P < 0.01) was greater for C compared with ST. The rumen fluid collection methods

had no effects on molar proportion of individual volatile fatty acids. Microbiome analysis

indicated no differences due to sampling methods on dominant phyla, classes, families, and

genera; however, differences were observed in specific microbial groups. The collection methods

had no effects on Chao 1 (P = 0.14) and Shannon index (P = 0.21). In conclusion, the

fermentation parameters and microbiome analysis were affected by the rumen fluid collection

method.

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INTRODUCTION

Background

An accurate understanding of rumen fermentation (Geishauser and Gitzel, 1996; Duffield

et al., 2004; Jiang, 2020) and ruminal microbiome (Hook et al., 2009; Lodge-Ivey et al., 2009;

Terré et al., 2013) is crucial to develop dietary strategies for improving production efficiency and

reducing environmental impact of ruminant production systems. In depth analysis can be

performed using the ruminal fluid to better understand the nutritional effectiveness of the animal

as well as for the maintenance of animal health and performance (Jiang, 2020). Analysis of

ruminal fluid can be used to assess ruminal fermentation (Geishauser and Gitzel, 1996; Duffield

et al., 2004) as well as to analyze the ruminal microbial community (Hook et al., 2009; Lodge-

Ivey et al., 2009; Terré et al., 2013).

Differences in Sampling Methods

Prior to analysis, an important challenge to overcome is the determining the best method

of collection of the ruminal fluid. Rumen sampling techniques can affect both fermentation

parameters and microbiome analysis. Rumen cannulation is considered a reference method for

rumen fluid collection because of the ease of colleting representative samples (Beharka et al.;

1998; Lesmeister and Heinrichs, 2004); however, cannulation requires surgical alteration and is

considered an invasive method which may not be broadly applicable. Less invasive alternatives

like pumping ruminal fluid using an oral stomach tube has also been used for collecting rumen

fluid samples (Abdelgadir et al., 1996; Coverdale et al., 2004; Khan et al., 2008); however,

samples collected using stomach tube are more susceptible to saliva contamination (Terré et al.,

2013).

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Purpose of the Study

The objectives of this study were to compare rumen fermentation parameters and

microbial population of rumen fluid collecting using rumen cannula or stomach tube. The

hypothesis was that the rumen cannula will provide a more representative sample compared to

stomach tube; however, fermentation pattern and abundance of dominant microbes will be

comparable between both techniques.

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MATERIALS AND METHODS

The ruminal fluid collection protocol was approved by the University of Florida

Institutional Animal Care and Use Committee.

Experiment Design

The ruminal fluid collection protocol was approved by the University of Florida

Institutional Animal Care and Use Committee. Three ruminally cannulated Holstein cows from

the University of Florida Dairy Unit were used in a 3 × 3 Latin square design. The cows were fed

the same diet (Table 1). The rumen fluid samples were collected from the cows with an oral

stomach tube and through the rumen cannula. The samples were collected in the mornings at six

dates during a two-month period.

Measurements and Sampling Procedures

For samples collected by esophageal stomach tube, approximately 200 mL of ruminal

fluid was collected 4 hours after the morning feeding using an orally administered stomach tube

connected to a vacuum pump (Ruminator; profs-products.com, Wittibreut, Bayern, Germany).

About 200 mL of rumen fluid was taken after discarding the first 200 mL of rumen fluid to

reduce saliva contamination. For samples collected from the rumen cannula, the rumen contents

were strained through four layers of cheese cloth. Following collection, rumen fluid was filtered

through four layers of cheesecloth, and pH was measured with a pH meter (Accumet AB15,

Fisher Scientific, Hampton, NH). Approximately 40 mL of ruminal fluid from each sample was

stored at -80ºC for analysis of bacterial diversity and abundance. Exactly 400 μL of 50% H2SO4

was added to another set of 40 mL samples to use for analysis of VFA and NH3-N. These

samples were centrifuged at 11,500 x g for 20 minutes. The supernatant was stored at -80ºC until

the VFA and NH3-N analysis was completed.

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Chemical analysis

Concentrations of acetate, propionate, butyrate, isobutyrate, isovalerate, and 2-

methylbutyrate were measured using an HPLC (FL 7485, Hitachi, Tokyo, Japan) according to

the method of Muck and Dickerson (1988). The column (Aminex HPX-87H, Bio-Rad

Laboratories, Hercules, CA) used a 0.015 M H2SO4 mobile phase and a flow rate set at 0.7

mL/min at 45ºC and was connected to a UV detector (Sprctroflow 757, ABI Analytical Kratos

Division, Ramsey, NJ) set at 210 nm. Concentrations of NH3-N were measured using the phenol-

hypochlorite assay.

Microbiome Analysis

Ruminal fluids samples were thawed at room temperature (about 22ºC) and DNA was

extracted and purified using the PowerLyzer PowerSoil DNA isolation kit (MOBIO Laboratories

Inc., Carlsbad, CA) with bead beating, following the protocol provided by the manufacturer.

Bead beating (Bullet159 Blender Storm 24, Next Advance, Averill Park, NY) was used to

homogenize the suspension and mechanically disrupt the bacterial cells. It entailed 3 min of

beating using 0.1 mm beads, followed by 15 min at 70ºC without beating and then another 3 min

of bead beating using the same beads. The DNA concentration and purity were measured using a

Nanodrop ND-1000 (Thermo Fisher Scientific, Waltham, MA). The mean DNA concentration of

samples was 68.65 ng/μL, and the absorbance (A) ratio at 260 and 280 nm (A260/A280) ratio was

between 1.75 and 1.88. The DNA integrity was verified using agarose (0.7%) gel electrophoresis

and extracted DNA was stored at -80ºC until further analysis.

Raw sequencing reads were obtained from the Illumina BaseSpace website and analyzed

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with the Quantitative Insights into Microbial Ecology (QIIME) pipeline (version 1.9.0). Chao and

Shannon index produced as alpha diversity was analyzed with the script: alpha_diversity.py.

Weighted UniFrac distance produced as beta-diversity measures and then subjected to principal

coordinates analysis (PCoA) with the script: beta_diversity_through_plots.py. Analysis of

similarities (ANOSIM) was used to detect the statistical difference of UniFrac distance metric with

the script compare_categories.py. The relative abundance of bacterial taxa at six-level taxonomic

classification (phylum, class, order, family, genus and species) was obtained with the script:

summarize_taxa_through_plots.py.

Statistical Analysis

A 3 × 3 Latin square design was used with three ruminally cannulated dairy cows. The

data were analyzed using GLIMMIX procedure of SAS (version 9.1, SAS Institute Inc., Cary,

NC). Statistical model included fixed effects of treatment (method of rumen fluid collection),

period, and interaction (treatment × period). Cow was used as random factor in the model.

Statistical differences were declared significant at P ≤ 0.05 and tendencies at 0.05 < P < 0.10.

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RESULTS

The nutrient composition of the diet is presented in Table 1. Experimental animals were fed corn

silage-based diets providing 16.4% CP and 28.7 starch on DM basis. Ground shelled corn and

soybean meal were used as energy and protein source in the concentrates.

Fermentation parameters

The rumen fermentation parameters analyzed included: pH, ammonia-nitrogen, as well as

total and individual volatile fatty acids and are presented in Table 2. Ruminal pH was affected by

the method of rumen fluid collection as pH values observed from stomach tube samples were

greater than contents collected using cannula (6.88 vs 6.25, P < 0.01). Ammonia-N concentration

was lower for the stomach tube samples compared with cannula samples (10.6 vs. 15.2, P =

0.01). Similarly, VFA concentration was lower for stomach tube samples compared to values

observed with cannula samples (95.5 vs 121.8, P < 0.01). No effects were observed on individual

VFA concentration including acetate (P =0.20), propionate (P =0.16), butyrate (P =0.36),

isobutyrate (P =0.64), isovalerate (P =0.87), and valerate (P =0.98). Acetate-to-propionate ratio

tended to be greater with stomach tube samples compared with cannula samples (P =0.08),

Rumen microbiome

Microbial diversity as observed by Chao1 and Shannon index is presented in Table 3. No

differences were observed on microbial diversity, Chao 1 (554 vs 591; P = 0.14) and Shannon

index (8.62 vs 8.72; P =0.21) because of method of rumen content collection.

The three most dominant phyla from rumen contents, regardless of method of sample

collection, were Bacteroidetes, Firmicutes, and Spirochaetes (Figure 1A). The relative

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abundance of bacterial phyla is presented in Table 4. The relative abundance of Bacteroidetes

was greater in stomach tube samples than in cannula samples (60.5 vs. 53.8, P < 0.01). On the

contrary, the relative abundance of Firmicutes (25.0 vs 29.6, P < 0.01), Spirochaetes (4.72 vs

6.04, P > 0.04), and Actinobacteria (0.75 vs. 1.10; P = 0.02) was lower in stomach tube samples

than in cannula samples. No effects were observed on the relative abundance of other bacterial

phyla including Proteobacteria (P = 0.91), Cyanobacteria (P = 0.32) and Fibrobacteres (P =

0.88).

The relative abundance of bacterial classes in the rumen contents are presented in Table 5

and Figure 1 C. Regardless of the method of sample collection, most dominant classes were

Bacterioidia, Clostridia, and Negativicutes (Figure 1 C). The relative abundance of Bacteroidia

(60.5 vs. 53.7; P < 0.01) and Coriobacteria (0.24 vs. 0.54; P < 0.01) was greater with stomach

tube samples compared with cannula samples. However, the relative abundance of Clostridia

(14.9 vs. 19.5; P < 0.01), Spirochaetia (4.73 vs. 6.03; P = 0.04) and Erysipelotrichia (0.42 vs.

0.58; P = 0.02) was lower with stomach tube samples. No effects were observed on the relative

abundance of Negativicutes (P = 0.89) and other classes including Kiritimatiellae (P = 0.55),

Fibrobacteria (P = 0.81), Melainabacteria (P = 0.26), Actinobacteria (P = 0.60),

Saccharimonadia (P = 0.36), Gammaproteobacteria (P = 0.80), and Mollicutes (P = 0.17).

Analysis from both sampling methods indicated the three most abundant bacteria belong

to the family (Figure 1B; Table 6) Prevotellaceae, Ruminococcaceae, and Acidaminococcaceae.

The relative abundance of Prevotellaceae (50.6 vs 39.2; P < 0.01) and Veillonellaceae (2.69 vs

1.93; P < 0.01) were greater in stomach tube compared with cannula samples. However, relative

abundance of Ruminococcaceae (6.88 vs. 9.03; P = 0.03), Rikenellaceae (3.23 vs. 3.89; P =

0.01), Lachnospiraceae (5.71 vs. 6.95; P = 0.04), and Spirochaetaceae (4.70 vs. 6.02; P = 0.04)

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was lower for stomach tube samples compared with cannula samples. No effects were observed

with the method of rumen content collection on the relative abundance of Succinivibrionaceae (P

= 0.71) and Acidaminococcaceae (P = 0.60).

The relative abundance of bacterial genus in rumen contents is presented in Table 7. The

relative abundance of Succiniclasticum, Treponema, and genus belonging to family

Prevotellaceae were among the most dominant genera observed in the rumen contents. The

relative abundance of Prevotella (36.7 vs. 28.1; P < 0.01) and unknown genus (3.66 vs. 2.0; P <

0.01) from Prevotellaceae were greater for stomach tube samples compared with cannula

samples. However, relative abundance of Treponema (36.7 vs. 28.1; P < 0.01), NK4A214 group

of Ruminococcaceae (2.22 vs. 3.35; P < 0.01), NK3A20 group of Lachnospiraceae (0.90 vs.

1.24; P = 0.01), Butyrivibrio (0.45 vs. 0.78; P = 0.05), and R-7 group of Christensenellaceae

(1.48 vs. 2.33; P < 0.01) were lower for the stomach tube samples compared with cannula

samples. The method used for rumen content collection had not effect on the relative abundance

of Succiniclasticum (P = 0.60), Ruminococcus (P = 0.51, Fibrobacter (P = 0.96), NK3B31 group

(P = 0.17), UCG-001 (P = 0.21), UCG-003 (P = 0.74) group of family Prevotellaceae, and

RF16_group of family Bacteroidales (P = 0.43).

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DISCUSSION

Rumen cannulation is the preferred method for collecting representative samples of

rumen digesta and is most commonly used for ruminant nutrition and microbiome research.

However, rumen cannulation is not feasible under most circumstances because of potential

animal welfare issues obliging us to depend on less invasive options like stomach tube sampling.

The present study was aimed at comparing fermentation characteristics and microbiome profile

from rumen samples collected using stomach tube and rumen cannula.

Several studies have used rumen cannulation and stomach tube samples for assessing

ruminal fermentation (Geishauser and Gitzel, 1996) and the structure of microbiome (Terre et al.,

2013) and have observed either similar or different effects depending on saliva contamination,

the type of sample collected, and the sampling site in rumen. The rumen fermentation parameters

analysis included pH, ammonia-nitrogen, and volatile fatty acids. The difference in pH is steady

with data presented by Morales (2014) and Duffield (2004). The higher pH values for the

stomach tube samples can be explained by potential saliva contamination (Duffield (2004);

Morales (2014)), even though during sample collection the first collection was always discarded

in effort to avoid this contamination. Some studies claim that salivary contamination is

minuscule (Lodge-Ivey, 2009), however our results showed highly significant difference.

The ammonia nitrogen results are different from the results of Lodge-Ivey (2009), where

it was reported that the data did not differ by sampling method. Total VFA and individual VFA

results are also different from those reported by Lodge-Ivey (2009), where it was reported that

total VFA and individual VFA proportions did not vary by collection method. The results from

our study do agree with the results of Terre et al. (2013), where it was reported that total VFA

concentrations were greater in rumen cannula samples than in stomach tube samples. Terre et al.

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(2013) attributes the difference in VFA concentrations to the saliva contamination of stomach

tube samples, which would decrease VFA concentration. The individual VFA parameters had

significant differences depending on the individual VFA being examined. The results from the

analysis of Terre et al. (2013) also reported that statistical significance varied depending on the

VFA. The data from Terre et al. (2013) showed that iso-butyric acid and valeric acid did not

have statistically significant differences between collection method.

Along with the rumen fermentation parameters, we also assessed fluctuation in microbial

population from rumen contents samples using stomach tube and rumen cannula. Recently, some

studies have observed overall resemblance in the ruminal microbiome between rumen contents

collected via stomach tube and rumen cannula; however, relative abundance of certain microbial

groups were reported to be different depending on the sampling methods (Lodge-Ivey et al.,

2009; Henderson et al., 2013). From the microbial analysis, it can be understood that both

sampling methods yield representative results with regards to the diversity indices and

population presence. Chao 1 provides an estimation of diversity based on abundance of microbes

belonging to certain class while Shannon index is commonly used to characterize species

diversity in a microbial community. The diversity indices show that both treatments indicate

highly diverse bacterial communities. Morales et al. (2014) observed diversity indices from

samples collected using stomach tube and rumen cannula in different species (goat and sheep)

fed different diets. While the indices were different in both sheep and goats; not difference was

observed because of the differences in the diet composition (Morales et al., 2014). Similarly,

both Chao1 and Shannon indices were comparable between stomach tube and rumen cannula

samples, agreeing with the results observed in the present study.

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We anticipated significant differences in the relative abundance of microbial population

due to methods used for sample collection since negligible amounts of solid material is collected

via stomach tube while cannula samples collect both solid and liquid fractions of the rumen

digesta. Our hypothesis was also based on the fact that stomach tubing will only allow small

highly degraded fiber fractions to be sampled and the primary colonizers may be under-

represented (Henderson et al., 2013). In addition, significant differences in the structure of

microbial community exists depending on the sampling site in rumen and this is due to the

presence of several micro-niches in the rumen. Sampling through rumen cannula allows for more

consistent sampling while stomach tube samples may be influences by sample collected from

specific rumen location (Shen et al., 2012).

While the overall community structure observed in this study was similar we observed

differences in the relative abundance of some microbial groups between the two sampling

methods. The relative abundance of family Prevotellaceae was increased 1.3-fold; however, the

abundance of family Lachnospiraceae was lower with sampling via stomach tube and the results

are in agreement with findings from previous study (Henderson et al., 2013). Despite the overall

resemblance of microbial community structure, we observed differences in the relative

abundance of phyla, families, class, and genera of microbial community. Based on these results,

samples collected from stomach tube and rumen cannula may give an valid quantitative

representation of microbial community structure; however, we should be cautious interpreting

data on the relative abundance of microbial groups from both methods considering differences

observed in the present study.

In conclusion, rumen fermentation characteristics including rumen pH was greater while

ammonia-N, and total VFA concentration were lower in rumen contents collected by stomach

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tube when compared with cannula samples; however, no differences were observed on individual

VFA concentration. Similarly, no differences were observed on the diversity indices and

dominant phyla, classes, families and genera of the microbial community; however, significant

differences of sampling type were observed in microbial groups. Rumen cannulation is

considered the reference method of collecting representative digesta samples for studying effects

on rumen fermentation and microbial composition; however, cannulation is invasive and may not

be most common and practical alternative for rumen sample collection. Stomach tubing is non-

invasive and is more practical and feasible alternative for collecting rumen digesta samples. This

study supports that stomach tubing is feasible alternative to rumen cannulation for collecting

rumen digesta samples. However, researchers should be cautious interpreting rumen

fermentation parameters from stomach tube samples as it tends to overestimate pH probably due

to salivary contamination and underestimate ammonia-N as well as result in differences in

abundance of bacterial communities. Further studies are required to better comprehend the

differences between both sampling methods and to validate if the differences are consistent

across studies. In the latter case, a mathematical correction factor may be estimated to account

for differences in parameters between the two collection methods to make esophageal stomach

tube sampling an appropriate alternative to rumen cannulation.

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LITERATURE CITED

Abdelgadir, I., Morrill, J., & Higgins, J. (1996). Effect of Roasted Soybeans and Corn on

Performance and Ruminal and Blood Metabolites of Dairy Calves. Journal of Dairy

Science, 79(3), 465-474. doi:10.3168/jds.s0022-0302(96)76387-7

Beharka, A., Nagaraja, T., Morrill, J., Kennedy, G., & Klemm, R. (1998). Effects of Form of the

Diet on Anatomical, Microbial, and Fermentative Development of the Rumen of

Neonatal Calves. Journal of Dairy Science, 81(7), 1946-1955. doi:10.3168/jds.s0022-

0302(98)75768-6

Coverdale, J., Tyler, H., Quigley, J., & Brumm, J. (2004). Effect of Various Levels of Forage

and Form of Diet on Rumen Development and Growth in Calves. Journal of Dairy

Science, 87(8), 2554-2562. doi:10.3168/jds.s0022-0302(04)73380-9

Duffield, T., Plaizier, J., Fairfield, A., Bagg, R., Vessie, G., Dick, P., . . . Mcbride, B. (2004).

Comparison of Techniques for Measurement of Rumen pH in Lactating Dairy Cows.

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Geishauser, T., and Gitzel, A. (1996). A comparison of rumen fluid sampled by oro-ruminal

probe versus rumen fistula. Small Ruminant Research, 21(1), 63-69. doi:10.1016/0921-

4488(95)00810-1

Henderson, G., Cox, F., Kittelmann, S., Miri, V.H., Zethof, M., Noel, S.J., Waghorn, G.C.,

Janssen, P.H., 2013. Effect of DNA extraction methods and sampling techniques on the

apparent structure of cow and sheep rumen microbial communities. PLOS ONE 8 (9),

e74787.

Hook S. E., Northwood K. S., Wright A.-D. G., and McBride B. W. 2009. Long-term monensin

supplementation does not significantly affect the quantity or diversity of methanogens in

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the rumen of the lactating dairy cow. Appl. Environ. Microbiol. 75:374–380.

doi:10.1128/AEM.01672-08.

Jiang, Y., Ogunade, I., Pech-Cervantes, A., Fan, P., Li, X., Kim, D., . . . Adesogan, A. (2020).

Effect of sequestering agents based on a Saccharomyces cerevisiae fermentation product

and clay on the ruminal bacterial community of lactating dairy cows challenged with

dietary aflatoxin B1. Journal of Dairy Science, 103(2), 1431-1447. doi:10.3168/jds.2019-

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Lesmeister, K., & Heinrichs, A. (2004). Effects of Corn Processing on Growth Characteristics,

Rumen Development, and Rumen Parameters in Neonatal Dairy Calves. Journal of Dairy

Science, 87(10), 3439-3450. doi:10.3168/jds.s0022-0302(04)73479-7

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(2014). Use of stomach tubing as an alternative to rumen cannulation to study ruminal

fermentation and microbiota in sheep and goats. Animal Feed Science and Technology,

198, 57-66. doi:10.1016/j.anifeedsci.2014.09.016

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Shen, J. S., Chai, Z., Song, L. J., Liu, J. X., Wu, Y. M. (2012) Insertion depth of oral stomach

tubes may affect the fermentation parameters of ruminal fluid collected in dairy cows.

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22921624.

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doi:10.3168/jds.2012-5921

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SUPPORTING FIGURES/TABLES

Table 1. Ingredient and chemical composition of the experimental diet

Ingredients % of diet DM

Corn silage 47.07

Corn grain, ground shelled 18.82

Soybean meal, 44% 15.14

Citrus pulp 4.71

Whole cottonseed 7.37

Palmit-801 1.68

Mineral and vitamin mix2 5.21

Nutrient composition (DM basis)

Crude protein, % 16.44

RDP (% CP) 59.26

RUP (% CP) 40.74

aNDFom, % 28.04

ADF, % 17.29

Starch, % 28.67

Sugar, % 3.97

NFC, % 42.51

Macro-minerals

Calcium, %3 0.65

Phosphorous, % 3 0.41

Magnesium, % 0.30

Potassium, % 1.06

Sulphur, % 0.19

Sodium, % 0.44

Chloride, % 0.35 1Global Agri Trade Corporation, Long Beach, CA.

2Vitamin mineral mixture (DM basis): Ca, 7.44%; P, 1.60%; Mg, 2.52%; K, 0.21%, S, 0.44%; Na,

8.13%; Cl, 3.30%; Biotin, 2.17 ppm; Fe 1221.5 ppm; Zn 1450.35 ppm; Cu, 220.32 ppm; Mn,

1180.14 ppm; Se, 7.33 ppm; Co, 22.51 ppm; I, 12.43 ppm; Vitamin A, 273.22 KIU/kg; Vitamin

D, 63.95 KIU/kg; Vitamin E, 546.44 IU/kg.

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Table 2. Rumen fermentation characteristics in samples collected via stomach tube or rumen

cannula

Stomach

Tube

Cannula SE Method Period

pH 6.88 6.25 0.16 <0.01 0.88

NH3-N, mg/dL 10.6 15.2 1.16 0.01 <0.01

Total VFA 95.5 121.8 5.31 <0.01 <0.01

Individual VFA,

mol/100 mol

Acetic acid 58.7 57.8 1.02 0.20 0.09

Propionic acid 19.6 20.6 1.03 0.16 0.80

Isobutyric acid 3.37 3.80 0.79 0.64 0.28

Butyric acid 13.9 13.4 0.44 0.36 0.11

Isovaleric acid 2.83 2.79 0.22 0.87 0.64

Valeric acid 1.64 1.63 0.21 0.98 0.18

Acetate-to-propionate 3.01 2.85 0.18 0.08 0.77

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Table 3. Microbial diversity determined in samples obtained via stomach tube or rumen cannula

Stomach

Tube

Cannula SEM Method Period

Diversity Indices Chao 1 554 591 18.6 0.14 0.23

Shannon 8.62 8.72 0.06 0.21 0.53

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Table 4. Effects of method of rumen content collection on the relative abundance of dominant

bacterial phyla in the rumen

Treatment

Stomach Tube Cannula SEM Treatment

Bacteroidetes 60.5 53.8 1.11 <0.01

Firmicutes 25.0 29.6 1.04 <0.01

Proteobacteria 1.07 1.04 0.22 0.91

Spirochaetes 4.72 6.04 0.87 0.04

Cyanobacteria 3.31 2.83 0.84 0.32

Actinobacteria 0.75 1.10 0.26 0.02

Fibrobacteres 0.77 0.81 0.18 0.88

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Table 5. Effects of method of rumen content collection on the relative abundance of bacterial

class in the rumen

Treatment

Stomach Tube Cannula SEM Treatment

Clostridia 14.9 19.5 1.35 <0.01

Mollicutes 0.43 0.53 0.05 0.17

Spirochaetia 4.73 6.03 0.90 0.04

Gammaproteobacteria 0.92 0.86 0.17 0.80

Saccharimonadia 0.63 0.72 0.12 0.36

Kiritimatiellae 1.44 1.56 0.25 0.55

Negativicutes 9.63 9.44 1.11 0.89

Erysipelotrichia 0.42 0.58 0.08 0.02

Fibrobacteria 0.76 0.82 0.18 0.81

Melainabacteria 3.31 2.76 0.85 0.26

Actinobacteria 0.49 0.55 0.15 0.60

Coriobacteria 0.24 0.54 0.11 <0.01

Bacteroidia 60.5 53.7 1.14 <0.01

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Table 6. Effects of method of rumen content collection on the relative abundance of bacteria

families in the rumen

Treatment

Stomach Tube Cannula SEM Treatment

Prevotellaceae 50.6 39.2 1.14 <0.01

Ruminococcaceae 6.88 9.03 0.75 0.03

Veillonellaceae 2.69 1.93 0.30 0.02

Rikenellaceae 3.23 3.89 0.48 0.01

Lachnospiraceae 5.71 6.95 0.86 0.04

Succinivibrionaceae 0.90 0.81 0.17 0.71

Spirochaetaceae 4.70 6.02 0.90 0.04

Acidaminococcaceae 6.88 7.54 0.98 0.60

24

Table 7. Effects of method of rumen content collection on the relative abundance of bacterial

genus in the rumen

Treatment

Stomach

Tube

Cannula SEM Treatment

Bacteroidales_RF16_group 0.55 0.49 0.05 0.43

F082_D_5__uncultured_rumen_bacteRIA 1.18 2.29 0.32 <0.01

Muribaculaceae D_5__uncultured rumen

bacterium

1.91 3.93 0.81 0.02

Genus (Prevotellaceae family)

Prevotella 36.7 28.1 1.14 <0.01

Prevotellaceae D_5__Prevotellaceae

NK3B31 group

0.80 0.65 0.20 0.17

Prevotellaceae UCG-001 4.67 4.16 0.35 0.21

Prevotellaceae UCG-003 2.76 2.69 0.35 0.74

Unknown genus 3.66 2.0 0.41 <0.01

Rikenellaceae family Alistipes 3.11 3.54 0.51 0.09

Fibrobacter 0.79 0.80 0.18 0.96

Christensenellaceae R-7 group 1.48 2.33 0.21 <0.01

Butyrivibrio 0.45 0.78 0.16 0.05

Lachnospiraceae NK3A20 group 0.90 1.24 0.20 0.01

Ruminococcaceae NK4A214 group 2.22 3.35 0.26 <0.01

Ruminococcus 0.58 0.72 0.25 0.51

Saccharofermentans 0.61 0.82 0.12 0.09

Succiniclasticum 6.87 7.53 1.00 0.60

Treponema 4.60 5.85 0.84 <0.01

25

Figure 1. Relative Abundance of Bacterial community in samples obtained via stomach tube or

rumen cannula. (A) at Phylum level; (B) at Family level; (C) at Class level.

(A)

0%

10%

20%

30%

40%

50%

60%

70%

80%

90%

100%

Stomach Tube Cannula

Bac

teri

al R

elat

ive

Ab

un

dan

ce

Phylum

Actinobacteria Bacteroidetes Chloroflexi Cyanobacteria

Elusimicrobia Fibrobacteres Firmicutes Kiritimatiellaeota

Lentisphaerae Patescibacteria Planctomycetes Proteobacteria

Spirochaetes Synergistetes Tenericutes Others

26

(B)

0%

10%

20%

30%

40%

50%

60%

70%

80%

90%

100%

Stomach Tube Cannula

Bac

teri

al R

elat

ive

Ab

un

dan

ce

Family

Prevotellaceae Ruminococcaceae

Acidaminococcaceae Lachnospiraceae

Spirochaetaceae Rikenellaceae

Veillonellaceae Muribaculaceae

Bacteroidales__F082 Gastranaerophilales__uncultured rumen bacterium

Christensenellaceae Kiritimatiellae__uncultured rumen bacterium

Gastranaerophilales Succinivibrionaceae

Fibrobacteraceae Bacteroidales RF16 group

Bacteroidales__uncultured Clostridiales__Family XIII

Saccharimonadaceae Erysipelotrichaceae

Bacteroidales__p-251-o5 others

27

(C)

0%

10%

20%

30%

40%

50%

60%

70%

80%

90%

100%

Stomach Tube Cannula

Bac

teri

al R

elat

ive

Ab

un

dan

ce

Class

Actinobacteria Alphaproteobacteria Anaerolineae

Bacilli Bacteroidia Clostridia

Coriobacteriia Deltaproteobacteria Elusimicrobia

Erysipelotrichia Fibrobacteria Gammaproteobacteria

Gracilibacteria Kiritimatiellae Lentisphaeria

Melainabacteria Mollicutes Negativicutes

Oxyphotobacteria Planctomycetacia Saccharimonadia

Spirochaetia Synergistia Uncultured Rumen Bacterium

Others