Amino Acid-Fermenting Bacteria from the Rumen of Dairy Cattle Enrichment, Isolation, Characterization, and Interaction with Entodinium Caudatum

Home , Acetoanaerobium, Butyrivibrio, Clostridium aminophilum, Clostridium sticklandii, Rumen

Amino Acid-Fermenting Bacteria from the Rumen of Dairy Cattle Enrichment, Isolation, Characterization, and Interaction with Entodinium caudatum

THESIS

Presented in Partial Fulfillment of the Requirements for the Degree Master of Science in the Graduate School of The Ohio State University

Jacqueline M. Gano

Graduate Program in Animal Sciences

The Ohio State University

2013

Master's Examination Committee:

Dr. Zhongtang Yu, Advisor

Dr. Jeffrey Firkins

Dr. Macdonald Wick

Copyrighted by

Jacqueline M. Gano

2013

ABSTRACT

Excess ammonia emissions are a major concern for the dairy industry due to the detrimental impact ammonia emissions have on the environment and wastage of dietary nitrogen. Hyper-ammonia-producing bacteria (HAB) and protozoa in the rumen are the major contributors of excessive ammonia excretions from cattle.

Besides Clostridium aminophilum, C. sticklandii, and Peptostreptococcus anaerobius, little is known about the HAB present in the rumen. In addition, rumen protozoa prey on bacteria and other microbes, excreting considerable amounts of amino acids and/or peptides that could promote the growth of HAB. In addition, inhibition of HAB by plant secondary metabolites may ultimately reduce ammonia production by HAB, thereby lowering excess nitrogen emissions. The studies presented in this thesis investigate HAB characterizations and interactions. In the first study, mixed microbes were obtained from the rumen of three fistulated dairy cows and further enriched and isolated for amino acid-fermenting bacteria. As a result, new isolates displayed high rates of ammonia production, ranging from 0.87 to 2.45 mg N/dL, and identified through 16S rRNA gene sequencing as a Bacillus sp. and Proteus mirabilis. In the second study, HAB enrichment cultures were co-cultured with an Entodinium caudatum culture. The co- culturing experiment was conducted with or without a feed substrate for E. caudatum

ii and Micrococcus luteus to assess the impact of feeding the protozoan. Ammonia concentrations were higher in the E. caudatum alone treatments, both with and without the addition of the feed substrate compared with HAB alone or co-culture of HAB and E. caudatum, with 20.5 ± 0.8, 18.5 ± 0.2, 23.9 ± 0.3, and 23.2 ± 1.1 mg ammonia N/dL in treatment groups E. caudatum with feed substrate, E. caudatum without feed substrate,

E. caudatum with feed substrate and with M. luteus, and E. caudatum without feed substrate but with M. luteus, respectively. Ammonia concentration was also significantly (P < 0.05) increased by M. luteus addition in the E. caudatum alone treatment group without feed substrate but with M. luteus. In the third study, HAB in enrichment cultures were examined for inhibition by varying plant secondary metabolites in terms of ammonia concentration. Results indicate that carvacrol, origanum oil, clove oil, and vanillin all successfully (P < 0.01) reduced ammonia concentration 24 h post incubation both in the presence and absence of a feed substrate. Overall, results indicate the presence of additional amino acid-fermenting bacteria in the rumen, capable of rapid ammonia production. These amino acid- fermenting bacteria, when grown in co-cultures with E. caudatum may interact with protozoa present within the rumen. These amino acid-fermenting bacteria are also sensitive to certain plant secondary metabolites, therefore decrease in ruminal ammonia concentration and shifts in ruminal microbe populations may be achieved through dietary supplementation with some of these plant secondary metabolites.

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ACKNOWLEDGEMENTS

I would like to thank everyone who has helped me over the last year.

I would like to thank Dr. Yu for helping me and pushing me to succeed; Jill Stiverson, for

teaching me how to use everything in the lab as well as putting up with my late night

texts when something went wrong in the beginning; Josie Plank, for helping me figure

out how to work with protozoa and not kill them; Lingling Wang for helping me with sequencing; Deng Pan for helping me with SAS; and everyone else in the lab for dealing with my questions and helping me along the way. And, lastly, I would also like to thank

my parents for standing beside me no matter what.

VITA

June 2007………………………………….Wilmington High School, Wilmington, Ohio

June 2011………………………………….B. S., Animal Sciences, The Ohio State University

2011-2013…………………………………Graduate Research Assistant, Department of

Animal Sciences, The Ohio State University

Field of Study

Major Field: Animal Sciences

TABLE OF CONTENTS

Abstract …………………………………………………………………………………………………………….…………..ii

Acknowledgments …………………………………………………………………………………………………….…..iv

Vita…………………………………………………………………………………………………………………………………v

List of Tables ………………………………………………………………………………………………………….….....ix

List of Figures ………………………………………………………………………………………………………...... x

CHAPTER 1. Review of Literature ……………………………….……………………………………..……….1

Ammonia ….…………………………………………………………...... 1

Deposition of Ammonia and Ammonium …..……...……………………………………2

Ammonia Emissions in Animal Agriculture …..……………………………..……….…3

The Ruminant Forestomach …..…………………………………………………………..…..5

Microorganisms in the Reticulorumen …….…………………………………………..…6

Hyper-Ammonia-Producing Bacteria .……………………………………………………..9

Ammonia Assimilation ………………………………………………...... 11

Transamination ……………………………………………………………………………….…..13

Inefficient N Retention in Cattle …………………………………………………………...14

Efficiency of Amino Acid Deamination .…………………………………………………15

Dietary Modification of Rumen Fermentation Using Plant Secondary

Metabolites …………………………………………………………………………….……………16

CHAPTER 2. Enrichment, Isolation, and Characterization of Amino Acid-Fermenting

Bacteria ………..…………………………………………………………………………………………………..………..18

Abstract …………………………………………………………………….…………..…………….18

Introduction ………………………………………………………………….……………..……...19

Materials and Methods ………………………………………………….………………….…23

Results and Discussion ………………………………………………………………….….…..26

CHAPTER 3. Investigation into the Interactions of a Consortium of Hyper-Ammonia-

Producing Bacteria and Protozoa ….……………………………………………………………….….………..33

Abstract ………………………………………………………………………………….………..….33

Introduction ………………………………………………………………………….…...………..34

Materials and Methods ……………………………………………………….……...…….…36

Results and Discussion ……………………………………………………………………..…..39

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CHAPTER 4. Investigation into the Inhibition of Ammonia production by a Hyper-

Ammonia-Producing Bacterial Consortium by Plant Secondary Metabolites ..………..…...47

Abstract …………………………………………………………………………….………………...47

Introduction ………………………………………………………………………………….……..48

Materials and Methods …………………………………………………………………….….51

Results and Discussion …………………………………………………………………..……..53

Literature Cited ……………………………………………………………………………………………………..…….63

Appendix A: Extraneous Tables and Figures …………………………………………………………………92

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LIST OF TABLES

Table 2.1. Characterization of isolates ..………………………………………………….….…………….….28

Table 3.1. Treatment Groups ………………………………………………………………………………....…..43

Table A.1. Effects of HAB and E. caudatum co-cultures without M. luteus on pH,

ammonia production, and protozoal counts in vitro ……………………..…………….125

Table A.2. Effects of HAB and E. caudatum co-cultures with M. luteus on pH,

ammonia production, and protozoal counts in vitro …………………....………..……126

Table A.3. Effects of plant secondary metabolites on ammonia production

and pH in vitro …………………………….……………………………………………….….……..….127

Table A.4. Effects of plant secondary metabolites with feed substrate on ammonia

concentration and pH in vitro …………………………………………………………………..…128

LIST OF FIGURES

Figure 2.1. Growth curve of HAB3 when grown on Casamino Acids, casein,

xylose, glucose, maltose, cellobiose, or starch ……………………………………………29

Figure 2.2. Growth curve of HAB5 when grown on Casamino Acids, casein,

xylose, glucose, maltose, cellobiose, or starch ……………………………………………30

Figure 2.3. Growth curve of HAB7 when grown on Casamino Acids, casein,

xylose, glucose, maltose, cellobiose, or starch ……………………………………………31

Figure 2.4. Growth curve of HAB8 when grown on Casamino Acids, casein,

xylose, glucose, maltose, cellobiose, or starch ……………………………………………32

Figure 3.1. Ammonia concentration in co-cultures 12 h post incubation ..…….………..……44

Figure 3.2. Ammonia concentration in co-cultures 24 h post incubation .…………...…….…45

Figure 3.3. E. caudatum counts in co-cultures 24 h post incubation …………….……..…..….46

Figure 4.1. Ammonia concentration in plant secondary metabolite treatments

12 h post incubation …………………………………………………………………………...…………..59

Figure 4.2. Ammonia concentration in plant secondary metabolite treatments

24 h post incubation ……………………………………………………………………………………60

Figure 4.3. Ammonia concentration in plant secondary metabolite treatments

with feed substrate 12 h post incubation …………………….………………………………61

Figure 4.4. Ammonia concentration in plant secondary metabolite treatments

with feed substrate 24 h post incubation …………………….………………………………62

Figure A.1. Growth curve of HAB3 when grown on Casamino Acids ………………….…….…..93

Figure A.2. Growth curve of HAB3 when grown on Casein ……………..…………………………...94

Figure A.3. Growth curve of HAB3 when grown on Cellobiose …………………………………….95

Figure A.4. Growth curve of HAB3 when grown on Control …………..…………………………….96

Figure A.5. Growth curve of HAB3 when grown on Glucose …………..……………………………97

Figure A.6. Growth curve of HAB3 when grown on Maltose …………..……………………………98

Figure A.7. Growth curve of HAB3 when grown on Starch .……………..…………………………..99

Figure A.8. Growth curve of HAB3 when grown on Xylose ……………..…………………………100

Figure A.9. Growth curve of HAB5 when grown on Casamino Acids ……..….………….……101

Figure A.10. Growth curve of HAB5 when grown on Casein ……………..…….…………………102

Figure A.11. Growth curve of HAB5 when grown on Cellobiose ……………….………………..103

Figure A.12. Growth curve of HAB5 when grown on Control …………..……….………………..104

Figure A.13. Growth curve of HAB5 when grown on Glucose …………..…….………………….105

Figure A.14. Growth curve of HAB5 when grown on Maltose …………..……………….………106

Figure A.15. Growth curve of HAB5 when grown on Starch .……………..………….……………107

Figure A.16. Growth curve of HAB5 when grown on Xylose ……………..…………….…………108

Figure A.17. Growth curve of HAB7 when grown on Casamino Acids ……………….….……109

Figure A.18. Growth curve of HAB7 when grown on Casein ……………..……………….…..….110

Figure A.19. Growth curve of HAB7 when grown on Cellobiose …………………………...……111

Figure A.20. Growth curve of HAB7 when grown on Control …………..………………….……..112

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Figure A.21. Growth curve of HAB7 when grown on Glucose …………..………………………..113

Figure A.22. Growth curve of HAB7 when grown on Maltose …………..……………………….114

Figure A.23. Growth curve of HAB7 when grown on Starch .……………..……………………….115

Figure A.24. Growth curve of HAB7 when grown on Xylose ……………..……………………….116

Figure A.25. Growth curve of HAB8 when grown on Casamino Acids ………………….…….117

Figure A.26. Growth curve of HAB8 when grown on Casein ……………..…………………..…..118

Figure A.27. Growth curve of HAB8 when grown on Cellobiose …………………………..….…119

Figure A.28. Growth curve of HAB8 when grown on Control …………..…………………….…..120

Figure A.29. Growth curve of HAB8 when grown on Glucose …………..………………….…….121

Figure A.30. Growth curve of HAB8 when grown on Maltose …………..……………….………122

Figure A.31. Growth curve of HAB8 when grown on Starch .……………..…………….…………123

Figure A.32. Growth curve of HAB8 when grown on Xylose ……………..…………….…………124

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CHAPTER 1

REVIEW OF LITERATURE

Ammonia

When exposed to room temperature and atmospheric pressure, ammonia assumes the properties of a colorless gas and displays a typical pungent odor. However, when ammonia undergoes pressure and is cooled, it will become a colorless liquid.

Ammonia is the most abundant alkaline component in the atmosphere (Asman, 1995).

There are several beneficial uses for ammonia in both commercial and industrial applications, including incorporation into many household cleaners and pharmaceutical supplies. Ammonia can even serve as a component of plastics, fibers, and rocket fuels

(Kim et al., 2013). However, ammonia can also have several pernicious effects.

Ammonia, in excess, can lead to environmental concerns; affecting the atmosphere and soil conditions among other charges.

Oxidation of nitrogen oxides (NOx) to HNO3 and SO2 to H2SO4 leads to the concentration of a substantial portion of the acid generated in the atmosphere; annual

-3 mean concentrations of NO, NO2, and SO2 in an urban environment average 48.7 µg m ,

38.3 µg m-3, and 8.34 µg m-3 respectively (Barrero et al., 2006). Ammonia neutralizes

1 nitric acid (HNO3), forming ammonium nitrate, as well as sulfuric acid (H2SO4) forming ammonium sulfate and bisulfate, thereby lowering the acidity of precipitation, cloudwater, and aerosols (Galloway, 1995; Finlayson-Pitts and Pitts, 2000).

Deposition of Ammonia and Ammonium

While NH3 can serve to neutralize HNO3 and H2SO4 in the atmosphere, the

+ deposition of NH3 and NH4 into terrestrial ecosystems can have harmful effects on soil quality and plant species. Through interaction with H2SO4, NH3 is capable of causing N saturation; ultimately resulting in soil acidification (Galloway, 1995). High N loads in the soil may also shift plant species. When nitrogen enrichment occurs in the soil due in part to high ammonia and ammonium deposition levels, oligotrophic ecosystems may be propelled towards a predominately nitrophilic ecosystem (Bobbink et al., 1992).

Vegetation unequipped to handle the high nitrogen load are prone to fatality or

+ impaired growth (Nielsen et al., 1991). NH4 nitrification can also lead to the leaching of

+ + + K , Ca2 , and Mg2 causing nutrient imbalances that can impair plant growth and also

+ release harmful ions, including Al3 (Krupa, 2003).

Ammonia has also been shown to have a key role in air emission quality. PM2.5, particulate matter in the air with a particle size less than 2.5 micrometers, has been directly associated with an increased risk of suffering from pulmonary disease, reduced lung function, and even premature death (Burnett et al., 2000; Oberdorster et al., 2000;

Pope et al., 2002; Gauderman et al., 2004). Through the concentration of NH4+ and

2 control over the state of neutralization for H2SO4, ammonia increases PM2.5 accumulation in livestock facilities (Pinder et al., 2007).

Ammonia Emissions in Animal Agriculture

In the United States alone, animal agriculture is estimated to be responsible for

50 to 85% of national ammonia emissions, making it the primary contributor to atmospheric reactive nitrogen (Battye et al., 1994). Other countries report similar emissions. A study in Ireland holds Irish agriculture responsible for an estimated 98.8 to

107 kt NH3-N in 2012 (Hyde et al., 2003). Estimates for the UKare even higher, at an estimated 230.4 kt NH3-N in 2011 (Misselbrook et al., 2011).

Within animal agriculture, beef and dairy cattle play an important role in ammonia emission. With 89.3 million heads of cattle as of January 2013 in the United

States alone, the cattle industry is a leading cause of ammonia emissions within animal agriculture (USDA, 2013). The previously described study in Ireland even holds cattle farming responsible for over 75% of total emissions within animal agriculture (Hyde et al., 2003). Within the cattle industry, ammonia emissions attributable to cattle can be broken down and categorized in terms of: cow-calf systems, cattle kept on feedlots, and cattle for use in the dairy industry.

Pasture-based cow-calf systems, used for reproductive maintenance of mature cows and heifers, have not been widely studied in the United States in terms of ammonia emission. However, a study in Europe determined an emission factor of 4.7 g

-1 -1 NH3 animal day for grazing beef cattle, putting the annual emission factor at 1.7 kg

-1 -1 NH3 animal yr (Misselbrook et al., 2000). Other studies in Europe have estimated much higher yearly emission factors, placing emissions at anywhere between 9 to 39.5 kg NH3 hd-1 yr-1 (McInnes, 1996; Bouwman et al., 1997; Van der Hoek, 1998; Battye et al., 1994; Doorn et al., 2002; Strader et al., 2002).

In beef cattle feedyards, high-concentrate diets are fed to cattle during the

“finishing” phase of concentration. During such time, a high level of N loss occurs in the system in terms of volatilized ammonia (Todd et al., 2011). In Kansas, N loss has been reported at 38% (Baum and Ham, 2009). In Alberta, Canada, this loss was found to be even higher, at 63% (McGinn et al., 2007). Other studies have reported values ranging between these two: 51 to 65%, 63 to 68%, and 51 to 68% in studies conducted on feedlots in New Mexico, Texas, and Nebraska, respectively (Bierman et al., 1999;

Erickson et al., 2000; Erickson and Klopfenstein, 2001; Cole et al., 2006; Farran et al.,

2006; Flesch et al., 2007; Todd et al., 2008). As for a quantified estimated annual emission factor for feedlots, three studies in Texas found feedyards to have emission

-1 -1 factors ranging from 10.3 to 54.8 kg NH3 hd yr (Koziel et al., 2004; Baek et al., 2006;

Flesch et al., 2007). Feedlot cattle in Colorado were found to fall within this range, at

-1 -1 17.7 kg NH3 hd yr (Hutchinson et al., 1982). In several European studies, the emission factor is comparable to the range for cow-calf systems, ranging from 9 to 39.5 kg NH3 hd-1 yr-1 (Bouwman et al, 1997; Van der Hoek, 1998; McInnes, 1996; Battye et al., 1994;

Doorn et al., 2002; Strader et al., 2002).

Dairy cattle, often kept either in free stall barns or maintained on an open-lot, differ significantly in ammonia emissions from those found in beef cattle, emitting approximately 2.25 times more ammonia per head (Doorn et al., 2002). A study on

-1 -1 Texas open-lot dairy cattle found annual emissions to be 9.4± 5.7 kg NH3 hd yr

(Mukhtar et al., 2008). Authors of this study also noted seasonal variation, discovering a

47% decrease in winter emissions when considering open-lot corrals. In California, two

-1 -1 studies found emission rates of 6.9 to 52.2 and 40 kg NH3 hd yr (Cassel et al., 2005;

Rumburg et al., 2008). However, the latter failed to collect samples evenly distributed over season, only collecting a few winter samples, possibly overestimating their emission factor. The national ammonia emission from dairy cattle, as estimated by

-1 -1 Pinder et al. (2004), was found to be 23.9 kg NH3 hd yr . Studies in Europe report

-1 -1 similar findings, with emission factors ranging from 22 to 40 kg NH3 hd yr (Battye et al., 1994; Sutton et al., 1995; Bouwman et al., 1997; Van der Hoek, 1998; Misselbrook et al., 2000; Strader et al., 2002; Rumburg et al., 2008).

The Ruminant Forestomach

Deamination of amino acids within the stomach of cattle is one of the major contributors to national ammonia emission. Cattle contain a complex stomach consisting of four major compartments, the reticulum, rumen, omasum, and abomasum. The abomasum, commonly referred to as the true stomach, aids in the digestion of microbial proteins and modified lipids. The ruminant ‘forestomach,’ is the term collectively given to the reticulum, rumen, and omasum. 5

The ruminant stomach, as well as the abomasum, is located in the abdominal cavity and monopolizes a large area within the cavity. The reticulorumen alone occupies

65% of the abdominal cavity (Hofmann, 1988). As such, the contents of the forestomach can account for close to 15 to 20% of the total body weight of ruminants (Giesecke and

VanGylswyk, 1975). Due to sheer size and turnover time, ruminants have the capacity to store feeds in their forestomach, aiding in rumination, fermentation, and absorption.

Feed is ingested, chewed and mixed with saliva, to form a bolus. This bolus is then swallowed and projected into the reticulum. The reticulum is separated from the rumen on the ventral side by the rumino-reticular fold. However, there is no sphincter separating the two compartments, so to a large extent they function as a singular organ, causing the vicinity to be commonly referred to as the reticulorumen. The rumen is stratified into 3 major components; a gas dome, a fibre mat, and a fluid layer (Tschuor and Clauss, 2008). Rumination (the act of regurgitation, swallowing, and re-chewing) allows the digesta to be broken down, pass into the omasum, the abomasum, and subsequently the small intestine.

Microorganisms in the Reticulorumen

Once feed reaches the reticulorumen, it is fermentated by microorganisms, including protozoa, bacteria, and fungi. Ciliate protozoa are present in concentrations of

105 to 106 cells/mL of ruminal fluid (Sylvester et al., 2004). Protozoa are thought to constitute the top of the tropic chain in the rumen (Ozutsumi et al., 2005), in turn giving

6 them the ability to affect fermentation both directly, through their own mechanisms of fermentation, or indirectly, through interactions, such as predation (Williams and

Coleman, 1992). Protozoa have a limited ability to synthesize amino acids from ammonia. As a result, the need for bacterial amino acids by protozoa is very high.

Bacteria range in total numbers from 1010 to 1011 cells per gram of digesta in the forestomach (Pond and Bell, 2005). Often broken into at least 28 functioning groups, including cellulolytics, amylolytics, lipolytics, and proteolytics (Pond and Bell, 2005), ruminal bacteria are capable of many fermentative reactions.

Cellulolytic bacteria are present in relatively high numbers within the rumen, and as their name suggests, they are capable of degrading cellulose as well as hemicellulose, the main components of dietary fiber. Ruminococcus flavefaciens, R. albus, and

Fibrobacter succinogenes are often considered the principle cellulolytic bacteria in the rumen (Dehority, 2003). However, a more diverse group of bacteria have been found through 16S rRNA gene-based studies to utilize forage feed particles, including

Clostridium, Acetivibrio, and Lachnospiraceae taxa (Larue et al., 2005; Brulc et al., 2009;

Kim et al., 2011).

Amylolytic bacteria are those ruminal bacteria capable of utilizing starch.

Bacteria found to possess high amylase activity include Streptococcus bovis, Prevotella ruminicola, Fibrobacter succinogenes, Bifidobacterium pseudolongum, Bifidobacterium thermophilum, Bifidobacterium adolescentis, Borrelia sp., Butyrivibrio sp., Eubacterium

7 ruminantium, Ruminobacter amylophilus, Ruminococcus bromii, Succiniomonas amylolytica, and Lactobacillus sp. (Kotarski et al., 1992). Interestingly, these species attack starch differently and, as such, may need to work in a consortium to fully digest cereal grains (McAllister et al., 1990; McAllister and Cheng, 1996).

Lipolytic bacteria are bacteria that possess the ability, through microbial lipases, to hydrolyze fats. Anaerovibrio lipolytica and Butyrivibrio fibrisolvens are two well- known lypolytic bacteria found in the rumen (Hobson, 1988). Protozoa have also been linked to microbial lipolytic activity, however they do not contribute to biohydrogenation (Jenkins et al., 2008).

Proteolytic bacteria are able to utilize proteins, breaking them down into smaller peptides and amino acids. Overall proteolytic activity in the rumen is moderate when placed into comparison with the animal’s own pancreatic and gastric secretions.

However, due to retention time of feed particles in the rumen, proteolytic bacteria are able to ferment a large portion of dietary proteins (Ørskov and McDonald, 1979;

Broderick et al., 1991; Wallace, 1996). Known species of proteolytic bacteria include

Ruminobacter amylophilus, Prevotella ruminicola, Butyrivibrio fibrisolvens, Butyrivibrio alactacidigens, Selenomonas ruminantium, as well as strains of Eubacterium,

Fusobacterium, and Clostridium (Dehority, 2003). These species are also able to breakdown peptides and amino acids and, as such, were thought to be responsible for significant ammonia production in the rumen (Attwood et al., 1998; Bladen et al., 1961).

When amino acids are provided in excess to that required for cell synthesis, the excess 8 are metabolized to ammonia. When intracellular concentrations of ammonia increase, ammonia diffuses out of the cell. Further studies concluded that in vitro rates of ammonia production exceeded the activities of predominant bacterial species capable of utilizing protein or protein hydrolysates (Chen and Russel, 1988; Russel et al., 1988;

Chen and Russel, 1989; Chen and Russel 1990; Russel et al., 1991). Continued research led to the discovery of bacteria, less numerous in the rumen, that were able to utilize trypticase for the production of ammonia at a rapid rate as a result of their constrained ability to derive ATP from sugar fermentation (Russell et al., 1991). Bacteria capable of this activity became known as hyper-ammonia producing bacteria (HAB).

Hyper-Ammonia-Producing Bacteria

HAB are bacteria that can produce ammonia at a rapid rate (>300 nmol NH3 mg cell protein-1 min-1) with a minimal ability to utilize sugars (Whitehead and Cotta, 2004;

Chen and Russel, 1989; Wallace, 1996; Flythe and Andries, 2009). The majority of HAB are considered to form an amino acid-utilizing niche; however a few HAB are capable of utilizing peptides. HAB have been isolated in cattle, sheep, and goats (Flythe and Kagan,

2010). Although only present in low numbers inside the rumen, the concentration of ammonia they are capable of producing makes them quantitatively important. The known HAB include Clostridium sticklandii, C. aminophilum, and Peptostreptococcus anaerobius (Paster et al., 1993). However, HAB species may vary between ruminants.

Krause and Russel (1996) were able to detect C. sticklandii, C. aminophilum, and P. anaerobius in cattle whereas Attwood et al. (1998) detected P. anaerobius, but not C. 9 aminophilum and C. sticklandii in New Zealand sheep. In other studies, several other novel bacteria have been identified with rapid rates of ammonia production that are taxonomically different from the three known isolates (McSweeney et al., 1999;

Eschenlauer et al., 2002), suggesting that many HAB remain uncultured.

In order for HAB to degrade amino acids and produce ammonia, they first need to deaminate the amino acids. Overall, there are five known pathways for deaminating amino acids. Four of these pathways are utilized to deaminate single amino acids, while the fifth, the Stickland reaction, utilizes pairs of amino acids. Overall, deamination can occur through oxidation, reduction, hydrolysis, or removal of elements of ammonia, resulting in the formation of an α-keto acid, a saturated fatty acid, an α-hydroxy fatty acid, or an unsaturated product, respectively (Smith and Macfarlane, 1997). C. sticklandii is well known for its ability to utilize the Stickland reaction, equipped with clostridial enzymes including glycine reductase and D-proline reductase which are capable of catalyzing an α-elimination. (Dürre, 2005). These enzymes ferment pairs of amino acids, in which one donates and one accepts an electron. The donor amino acid is deaminated to a 2-oxo acid and then used for ATP conservation when the acid is oxidatively decarboxylated to the corresponding acyl-CoA (Dürre, 2005). The acceptor amino acid is reduced to a fatty acid.

- + Alanine + 2 glycine + 2H2O →3 acetate + 3NH4 + CO2

Ammonia Assimilation

In the rumen, ammonia is considered the most important source of N for protein synthesis by many microbial species. Optimal ruminal ammonia concentration for maximal microbial growth, according to Wallace and Lahlou-Kassi (1995), is between 2 to 3 mM. However, as the authors admitted, this assumed no limiting nutrient, which likely oversimplifies and underestimates the concentration. A later study by Schwab et al. (2005) estimated the optimal concentration to be between 5 to 11 mM, depending on fermentation conditions. Most bacteria in the rumen can utilize ammonia as their main source of N, but in some isolates, ammonia utilization is even essential for growth

(Wallace and Lahlou-Kassi, 1995).

Ammonia assimilation first begins by diffusion of ammonia across the cell membrane and into the cell. Once ammonia is translocated into the cell, it is assimilated into amino acids by ruminal bacteria, most often through NAD-linked glutamate dehydrogenase (NAD-GDH) as evidenced by early studies on ruminal mixed populations

(Erfle et al., 1977; Wallace, 1979; Lenartova et al., 1987; Atasoglu et al., 1999). The NAD-

GAD pathway is as follows:

+ + 2 – Ketoglutarate + NH3 + NADH + H ↔ Glutamate + NAD

Other enzymes of assimilation include NADP-linked glutamate dehydrogenase (NADP-

GDH), the dual enzyme system consisting of glutamine synthetase (GS) and glutamate synthase (GOGAT), and alanine dehydrogenase (Wallace, 1979; Chalupa et al., 1983).

NAD-GDH enzymes have a high Km for ammonia, falling between 20-33 mM

(Erfle et al., 1977; Wallace, 1979), which allows the GDH pathway to function efficiently only in ammonia-rich, energy poor conditions (Villapakkam, 2008). Glutamate is the most abundant amino acid as a result of this mechanism (Atasoglu et al., 1999). When ammonia concentrations fall, however, NADP-GDH with a Km of 2-3 mM of ammonia, prevails (Chalupa et al., 1983; Erfle et al., 1977; Wallace, 1979; Lenartova et al., 1987).

However, even as ammonia concentrations fall, glutamate is still predominantly produced. Conditions with low levels of ammonia can also increase GS, though prevalence in vivo is uncertain (Schwab et al., 2005; Erfle et al., 1977). In GS and GOGAT coupled reactions, the enzyme GS first fixes ammonia in the amide of glutamine:

Glutamate + ATP + NH3 → Glutamine + ADP + P

GOGAT then catalyses transfer of the glutamine amide to 2-ketoglutarate (Meers et al.,

1970):

2 – Ketoglutarate + Glutamine + NADPH + H+ → 2 Glutamate + NADP+

In contrast to low ammonia concentrations alanine is the first amino acid present in high ammonia concentrations, as shown by Wallace (1979) and Blake et al. (1983). This suggests that the enzyme alanine dehydrogenase, with a Km of 70 mM (Wallace, 1979) is increased significantly in conditions of high ammonia concentrations.

Once ammonia is assimilated into glutamate, or in some cases alanine, N from

NH3 is distributed to other amino acids through the process known as transamination

(Schwab et al., 2005).

Transamination

The degradation of most standard amino acids occurs through transamination.

During this procedure, an α-amino group from an amino acid is transferred to an α-keto group using what is known as an aminotransferase. Aminotransferases, also commonly referred to as transaminases, are enzymes capable of interconverting amino acids and keto acids. Many different aminotransferases exist, showing high variability among substrate specificity. In bacteria the best known are glutamate-oxaloacetate and glutamate-pyruvate aminotransferases (Schwab et al., 2005). Some aminotransferases have specificity towards a particular amino acid or classes of amino acids, or an α-keto acid receptor. Ultimately, through the use of a transaminase, the α-amino group present on an amino acid is often transferred to α-ketogluterate, an α-keto acid, resulting in the concentration of an oxoacid and glutamate. The reaction is reversible, so both glutamate and α-ketogluterate serve as substrates.

Amino acid + α-ketogluterate ↔ oxoacid + glutamate

In the transamination reaction there is only a transfer of amino groups, meaning no free ammonia is liberated (Fursule, 2008). Following transamination, synthesized amino acids then continue on to be used for the biosynthesis of proteins.

Inefficient N Retention in Cattle

Although used by many rumen bacteria for synthesis of amino acids and proteins, as previously described, ammonia concentration in the rumen becomes a concern when it exceeds what can be utilized by the microbial population present. As a result, ammonia will be lost across the ruminal wall; serving as a main cause in ruminants of inefficient N retention (Leng and Nolan, 1984).

Diffusion across the rumen wall depends on the form of ammonia. NH3, the non- ionized form, is able to freely diffuse through the lipid bilayers present in the rumen wall

+ (Dinh, 2007). However, NH4 , the ionized form of ammonia, is not capable of this passive

+ transport. The current thinking is that NH4 uses some sort of electrogenic, facilitated transport, such as potassium channels, to gain access to the blood stream (Abdoun et

+ al., 2005; Dinh, 2007). However, it has recently been suggested that NH4 may cross

+ through the rumen wall through use of NH4 transport proteins (Abdoun et al., 2006).

Once ammonia has crossed the ruminal wall and entered the blood, detoxification occurs mainly in the periportal cells within the liver. However, recent studies have demonstrated that some detoxification of ammonia to urea can occur in rumen epithelial cells and duodenal mucosal cells (Oba et al., 2004). Requiring ATP, urea synthesis begins when one bicarbonate ion and one ammonia molecule synthesize carbamoyl-phosphate. From here, the ornithine-urea cycle begins, ultimately resulting in the concentration of urea (Dinh, 2007). Following synthesis, urea faces one of two passage routes. First, it may enter the gastrointestinal tract to be further utilized by the 14 ruminal microbial population. Reported values of urea-N entering the gastrointestinal tract differ between sex, age, diet, and species, with reports averaging 171 g N/day for dairy cattle and 28.1 g N/day for steers (Lapierre and Lobley, 2001; Archibeque et al.,

2002). Synthesized urea may also be excreted through the kidneys via urine. This urine, when excreted, can react with urease enzymes found in both feces and the soil, ultimately becoming hydrolyzed to ammonium (Faulkner and Shaw, 2008) and causing environmental concerns, as previously discussed.

Efficiency of Amino Acid Deamination

Obligate amino acid-fermenting bacteria have rapid rates of amino acid deamination. However, this anaerobic degradation of amino acids provides little energy to the bacteria (Dijkstra et al., 2005). As such, obligate amino acid-fermenting bacteria have been shown to degrade 10 to 25 times more amino acids than they were able to incorporate into microbial protein (Chen and Russell, 1988), indicating an inefficient use of amino acids.

Methods to better modulate efficient microbial protein synthesis and improve efficient use of amino acids within the rumen through reduced numbers and activity of

HAB are currently under review. While ammonia serves as an important source of nitrogen for ruminal microorganisms, some ruminal bacteria are also able to utilize amino acids as a nitrogen source. As such, certain bacteria, including carbohydrate- fermenting bacteria, may be capable of ultimately modulating HAB numbers (Rychlik

15 and Russell, 2000). These saccharolytic bacteria compete with HAB for the same amino acid pool, and due to the high abundance of saccharolytic bacteria they may be able to outcompete HAB, thereby lowering deamination of amino acids by HAB, especially when readily fermentable feed is fed. In particular, Streptococcus bovis has been shown to have an increased growth rate when provided access to preformed amino acids, reducing energy spilling (Russel, 2007), and thereby may show promise at inhibiting HAB within the rumen.

Dietary modification of rumen fermentation using plant secondary metabolites

Many methods have been proposed to lower ammonia emissions, including the use of plant secondary metabolites. Through secondary metabolism, many plant species produce a variety of organic compounds. These secondary metabolites are often placed into one of three groups: saponins, tannins, and essential oils, some exhibit antimicrobial activity against many bacteria, yeasts, and even molds (Gershenzon and

Croteau, 1991).

Saponins contain a high-molecular-weight glycoside that contains saccharide chain units linked to either a triterpene or steroidal aglycone moiety (Patra and Saxena,

2009). Antiprotozoal activity of saponins is generally attributed to their interaction with the sterol moiety of protozoa (Hostettmann and Marston, 1995), mainly present within the cell membrane. As a result, saponins are capable of destroying protozoal cell membranes, leading to leakage of cell contents (Patra and Saxena, 2009). Although

16 many studies have found antiprotozoal activity for saponins (Ivan et al., 2004; Baah,

2007; Singer et al., 2008), the effects of saponins on bacteria and fungi are less known and appear to be species-dependent (Wallace et al., 1994; Wang et al., 2000; Patra and

Saxena, 2009).

Essential oils are classified as secondary metabolites that are obtained by steam distillation from the plant volatile fraction (Gershenzon and Croteau, 1991) and contain two main chemical groups: terpenoids and phenylpropanoids. Both terpenoids and phenylpropanoids use a mechanism of action dependent on interaction with the cell membrane (Davidson and Naidu, 2000; Dorman and Deans, 2000). The hydrophobic cyclic hydrocarbons interact and accumulate within the bacterial lipid bilayer (Sikkema et al., 1994; Ultee et al., 1999), eventually leading to changes in the structure of the membrane, causing expansion and fluidification (Griffin et al., 1999). In the rumen environment, this process can lead to shifts in ruminal bacterial populations and ultimately mitigate energy and protein losses.

CHAPTER 2

ENRICHMENT, ISOLATION, AND CHARACTERIZATION OF AMINO ACID-FERMENTING

BACTERIA

ABSTRACT

Amino acid-fermenting bacteria, particularly hyper-ammonia-producing bacteria

(HAB) in the rumen contribute to excessive ammonia excretions from cattle operations.

Amino acid-fermenting bacteria are able to catabolize amino acids to ammonia, some of which ultimately is utilized by the ruminant. However, ammonia becomes a concern when concentrations in the rumen are in excess of that which can be utilized by the ruminal microbial population. In this study experiments were undertaken to enrich and isolate new strains of amino acid-fermenting bacteria and to characterize their ability to grow on several substrates. Fresh rumen fluid was collected from three ruminally fistulated dairy cows and combined as the source inoculum. Amino acid-fermenting bacteria were enriched in a medium containing Casamino Acids as sole substrates.

Following successive enrichment, the enrichment cultures were plated, and individual

18 colonies displaying rapid ammonia production concentration were isolated and characterized for ammonia production from casamino acids and the ability to grow in media containing starch, cellobiose, maltose, glucose, xylose, or casein. Strains displayed high rates of ammonia production (0.87 to 2.45 mg N/dL) when incubated at

39°C with an initial pH of 6.5 for 24 h. Strains were further identified through 16S sequencing as a Bacillus sp. and Proteus mirabilis. These results indicate that new and novel amino acid-fermenting bacterial species remain to be found in the rumen.

INTRODUCTION

Concerns over excess ammonia emissions in the dairy industry have recently come to light as a result of the detrimental impact ammonia emissions have been shown to have on the environment (Kohn, 2005). Research in recent years within the food animal industry has sought to improve nitrogen retention in an attempt to decrease these nitrogenous emissions. Currently, only 5 to 30% of nitrogen from animal feed is retained; the rest is excreted, leading to possible detrimental effects on the environment. One area within the focus of controlling ammonia emissions and nitrogen retention is the control of hyper-ammonia-producing bacteria (HAB).

Hyper-ammonia-producing bacteria are bacteria that express a rapid rate of ammonia production with a minimal ability to utilize sugars (Chen and Russel, 1989;

Wallace, 1996; Whitehead and Cotta, 2004; Flythe and Andries, 2009). These HAB are able to catabolize amino acids and sometimes short peptides to ammonia. This ammonia can undergo assimilation and further transamination by ruminant 19 microorganisms for the concentration of amino acids, the building blocks of proteins.

However, ammonia production in the rumen becomes a concern when it exceeds what can be utilized by the microbial population present. As a result, ammonia will be absorbed across the ruminal wall, serving as a main cause in ruminants of inefficient N retention (Leng and Nolan, 1984).

HAB have been discovered in sheep, goats, and cattle (Flythe and Kagan, 2010).

Although these bacteria are present in low levels inside the rumen, they are quantitatively important in terms of their ammonia production rate. The known ruminal

HAB include Clostridium sticklandii, Clostridium aminophilum, and Peptostreptococcus anaerobius (Paster et al., 1993).

Clostridium aminophilum, as studied by Paster et al. (1993), is an obligate Gram- positive anaerobe displaying an irregular rod morphology. These rods are 1.0 µm wide and 1.5 µm long. Considered nonmotile, growth is reported to occur between 25 to

45°C. Growth occurs on amino acids or peptides and glutamine, glutamate, serine, and histidine as the preferred carbon sources. With the ability to utilize amino acids, C. aminophilum has recently fallen under scrutiny as a potential inhibitor of growth of

Campylobacter jejuni, a bacterium capable of causing foodborne illness (Anderson et al.,

2010). Fermentative end products from fermentation of Casamino Acids include ammonia, acetate, and butyrate, with trace amounts of lactate and succinate.

Phylogenetic analysis places C. aminophilum in the genus Clostridium cluster XIVa within the family Lachnospiraceae.

Clostridium sticklandii, also an obligate Gram-positive bacterium, shows high sequence similarity of its 16S rRNA gene to those in the cluster XI of clostridia (Rainey et al., 2001). However, this species was assigned to the genus Acetoanaerobium in the family of Peptostreptococcaceae. Arginine, serine, threonine, cysteine, proline, and glycine appear to be the preferred carbon sources for C. sticklandii, disappearing rapidly when provided in the medium during the exponential growth phase (Fonknechten,

2010). C. sticklandii is also capable of utilizing pairs of amino acids, producing acetate, butyrate, and ammonia via the Stickland reaction (Balows et al., 1992).

Peptostreptococcus anaerobius is a Gram-positive, non-spore-forming obligate coccal anaerobe (Rainey et al., 2001). P. anaerobius is able to metabolize peptone as well as amino acids to acetic, butyric, isobutyric, caproic, and isocaproic acid. P. anaerobius is capable of utilizing leucine, serine, threonine, glycine, and phenylalanine

(Chen and Russel, 1988). When proline, glycine, alanine, valine, and isoleucine were provided as Stickland pairs, deamination was more rapid then when provided alone.

The occurrence of these species has been shown to vary between ruminants; C. sticklandii, C. aminophilum, and P. anaerobius were all detected in cattle (Krause and

Russel, 1996). However, P. anaerobius alone was isolated in New Zealand sheep

(Attwood et al., 1998). Several other new ruminant bacteria have been identified with

21 rapid rates of ammonia production as well as presenting taxonomical differences to the three known isolates (McSweeney et al., 1999; Eschenlauer et al., 2002). This information suggests that many amino acid-fermenting bacteria, and possibly HAB species, remain uncultured. As a result, before future methods of control of ammonia emissions are developed, more research needs to be performed on ruminal guilds and their relationships with one another. With this need in mind, the primary objective of this study was to isolate and characterize amino acid-fermenting bacteria from the rumen of dairy cattle.

Within these characteristics, the substrate utilization characteristics of amino acid-fermenting bacteria have proven to be of interest. Several studies have looked at varying enrichment media for HAB to see how changes in the media can affect growth as well as ammonia production. Eschenlauer et al. (2002) found an HAB isolate able to ferment glucose, sucrose, and maltose, trypticase and casein amino acids. As such, this raises another area of interest where amino acid-fermenting bacteria are concerned: the utilization of certain substrates by HAB and thereby ammonia production on varying substrates. Therefore, the secondary objective of this study was to test ability of amino acid-fermenting bacterial isolates to utilize starch, cellobiose, maltose, glucose, xylose, and casein when implemented into a basal medium.

MATERIALS AND METHODS

HAB Enrichment

Six hours after feeding, ruminal contents was collected from three ruminally fistulated dairy cows. Samples, including both the raft and liquid phase of rumen contents, were collected in plastic screw-cap containers that were filled to the brim and kept at 39°C prior to transfer to the laboratory. An equal volume of each of the 3 samples were combined and used as the inoculum, which was inoculated into a semi- defined medium containing Casamino Acids (30g/L) as the sole substrate. The semi- defined medium (pH 6.5 to 6.6) of Attwood et al. (1995) contained (per liter) K2HPO4

292 mg, KH2PO4 292 mg, Na2SO4 480 mg, NaCl 100 mg, CaCl2∙2H2O 64 mg, Na2CO3 4 g, yeast extract 0.5 g. Resazurin (0.01%) was used as the indicator of anaerobiosis of the broth. The enrichment cultures in Hungate tubes, which were set up in an anaerobic chamber and capped with rubber stoppers and aluminum seals, were incubated at 39

°C. The cultures were repeatedly transferred into the same type of fresh medium every

24 h.

Isolation of Amino Acid-Fermenting Bacteria

Once stabilization of the enrichment activity occurred, the enrichment cultures were anaerobically serially diluted and aliquots (0.1 mL) of the dilutions were plated on agar (2%) plates (pH 6.5 - 6.6) containing Casamino Acid as the sole substrate and phenol red (0.02% w/v) as indicator of ammonia production as defined by

Santoshkumar et al. (2010). The plates were anaerobically incubated at 39°C. Colonies that displayed a color change of the phenol red indicator dye from yellow-orange to red- pink were selected as positive for ammonia production. Select positive colonies were streaked on Casamino Acid plates until only one colony morphology was visible.

Colonies were then confirmed for ammonia production activity in Casamino Acid medium (pH 6.5 to 6.6) containing phenol red (0.02% w/v) and ammonia production was compared to a known HAB, P. anaerobius.

Characterization of HAB isolates

The cellular morphology of each isolate was determined following standard bacteriology method and Gram-staining; employing crystal violet, Gram’s iodine, 95% alcohol, and Safranin. The 16S rRNA genes of the new amino acid-fermenting bacterial isolates isolated above were PCR amplified using bacterial primers 27f and 2515r using the cells of each isolate as DNA template. The size of the PCR product was confirmed by agarose electrophoresis. The PCR product was then purified using a PCR Product

Purification kit (Qiagen) and then sequenced using 357f and 519r primers using a CEQ™

8800 Genetic Analysis System (Beckman Coulter). The two sequence reads for each isolate were assembled by sequence alignment using BioEdit. The isolates were classified and confirmed as pure cultures by comparing the sequences to the sequences in the RDP database (http://rdp.cme.msu.edu/classifier/classifier.jsp) using the Classifier program implemented in the RDP database.

The ability of the new amino acid-fermenting bacterial isolates to produce ammonia was determined by using Casamino Acid (30 g/L) medium in triplicate after 24 h for each isolate. The concentration of ammonia in the spent medium was determined using the phenol-hypochlorite reaction as described in Chaney and Marbach (1961).

The new isolates, HAB3 (a strain of Proteus mirabilis) and HAB5 (Bacillus sp.), together with two amino acid-fermenting bacterial strains isolated previously

(Phongthorn Kongmun, et al., unpublished data), HAB7 that was classified to

Fusobacterium ulcerans and HAB8 that was classified to P. anaerobius, were further characterized for their ability to use saccharides and protein utilization using a medium modified from the defined medium of Attwood et al. (1995). Substrates and 1% (v/v) inoculum were added to 10 ml medium (pH of 6.5 to 6.6) and incubated anaerobically for 24 h at 39°C. Substrates included 1.5% (w/v) starch, cellobiose, maltose, glucose, xylose, casein, and Casamino Acid. Optical density (600 nm) of each culture tube was then determined at 0 h, 2 h, 4 h, 8 h, 12 h, and 24 h.

Analysis

Optical density was analyzed as repeated measures of a completely randomized design using the MIXED procedure of SAS 9.2 (SAS Inst. Inc., Cary, NC).

RESULTS AND DISCUSSION

Five new isolates were found displaying high ammonia production in a Casamino

Acid medium, hereby designated as HAB1, HAB2, HAB3, HAB4 and HAB5. HAB1, HAB2,

HAB3, and HAB4 were identified through 16S sequencing as Proteus mirabilis (2.10 to

2.45 mg N/dL). HAB5, 0.87 ± 0.01 mg N/dL, was identified as a strain of Bacillus. Two previously isolated amino acid-fermenting bacteria, HAB7 and HAB8, were found to display high 16S sequence similarity to Fusobacterium ulcerans (2.86 ± 0.08 mg N/dL) and Peptostreptococcus anaerobius (4.97 ± 0.05 mg N/dL), respectively. Sequence similarity and morphology for each are displayed on Table 2.1.

Proteus mirabilis falls within the family Enterobacteriaceae and is a facultative anaerobe capable of both respiratory and fermentative metabolism (Brenner et al.,

2005). P. mirabilis is able to deaminate several amino acids, resulting in keto acids and ammonia (Stumpf and Green, 1944; Singer and Volcani, 1955; Baek et al., 2011), and as such, P. mirabilis has been isolated and found capable of high levels of ammonia production (Whitehead and Cotta, 2003). P. mirabilis has been reported to contain two separate amino acid deamination enzymes; one capable of deaminating a range of aliphatic and aromatic amino acids and the other only displaying activity against a lower range of basic amino acids (Pelmont et al., 1972; Duerre and Chakrabarty, 1975; Baek et al., 2011).

These isolates were tested for their ability to utilize select substrates relevant to the rumen (Figure 2.1-2.4, A.1-A.32). All four isolates showed the ability to degrade cellobiose (P < 0.05); however, previous studies have reported fermentation of cellobiose to be limited in F. ulcerans (Staley and Whitman, 2001). HAB3, HAB7, and

HAB8 displayed amylolytic activity, with significant optical density changes when grown on starch. The ability to ferment maltose is uncharacteristic of many P. mirabilis and F. ulcerans strains, which were thought to have a limited ability to utilize maltose (Brenner et al., 2005; Staley and Whitman, 2001). Isolates HAB3, HAB5, HAB7, and HAB8 all displayed an ability to ferment glucose, with a significant optical density change over time, supporting previous studies that have shown the ability of P. mirabilis, P. anaerobius, and F. ulcerans to utilize glucose (Park et al., 2012; Ezaki et al., 2006; Staley and Whitman, 2001). HAB3, HAB5, HAB7, and HAB8 were also found capable of xylose fermentation (P < 0.05) and some activity (P < 0.05) to degrade casein.

Overall, these new isolates suggest additional amino acid-fermenting bacterial species beyond the known C. sticklandii, C. aminophilum, and P. anaerobius that display high capacity of ammonia production and unique saccharide and protein utilization.

Through further isolation and understanding of amino acid-fermenting bacterial species, including their growth characteristics on varying energy substrates, new interventions can hopefully be found for the dairy industry to reduce nitrogen outputs from dairy facilities.

Table 2.1. Characterization of Isolates.

% Ammonia Sequence Concentration Isolate Nearest Taxon Similarity Morphology (mg N/dL) Rod, Gram HAB1 Proteus mirabilis 97 negative 2.10 ± 0.02

Rod, Gram HAB2 Proteus mirabilis 99 negative 2.18 ± 0.02

Rod, Gram HAB3 Proteus mirabilis 99 negative 2.45 ± 0.05

Rod, Gram HAB4 Proteus mirabilis 99 negative 2.22 ± 0.02

Rod, Gram HAB5 Bacillus sp. 98 positive 0.87 ± 0.01

Fusobacterium Coccal, Gram HAB7 ulcerans 98 negative 2.86 ± 0.08

Peptostreptococcus Coccal, Gram HAB8 anaerobius 99 positive 4.97 ± 0.05

Each isolate was tested in a Casamino Acid medium in triplicate. All incubations occurred at 39°C with a pH of 6.5 to 6.6 in Hungate tubes capped with rubber stoppers and aluminum seals. Ammonia data represent the mean values ± SEM.

Figure 2.1. Growth curve of HAB3 when grown on Casamino Acids, casein, xylose, glucose, maltose, cellobiose, or starch. Each substrate was used at 1.5% (w/v) concentration. All incubations occurred at 39°C with a pH of 6.5 to 6.6 in Hungate tubes capped with rubber stoppers and aluminum seals. Each treatment was done in triplicate and the data represent the mean values.

Figure 2.2. Growth curve of HAB5 when grown on Casamino Acids, casein, xylose, glucose, maltose, cellobiose, or starch. Each substrate was used at 1.5% (w/v) concentration. All incubations occurred at 39°C with a pH of 6.5 to 6.6 in Hungate tubes capped with rubber stoppers and aluminum seals. Each treatment was done in triplicate and the data represent the mean values.

Figure 2.3. Growth curve of HAB7 when grown on Casamino Acids, casein, xylose, glucose, maltose, cellobiose, or starch. Each substrate was used at 1.5% (w/v) concentration. All incubations occurred at 39°C with a pH of 6.5 to 6.6 in Hungate tubes capped with rubber stoppers and aluminum seals. Each treatment was done in triplicate and the data represent the mean values.

Figure 2.4. Growth curve of HAB8 when grown on Casamino Acids, casein, xylose, glucose, maltose, cellobiose, or starch. Each substrate was used at 1.5% (w/v) concentration. All incubations occurred at 39°C with a pH of 6.5 to 6.6 in Hungate tubes capped with rubber stoppers and aluminum seals. Each treatment was done in triplicate and the data represent the mean values.

CHAPTER 3

INVESTIGATION INTO THE INTERACTIONS OF A CONSORTIUM OF HYPER-AMMONIA-

PRODUCING BACTERIA AND PROTOZOA

ABSTRACT

Hyper-ammonia-producing bacteria (HAB) and protozoa in the rumen contribute to excessive ammonia production from cattle. In this study, the relationship between enriched HAB and protozoa was examined. Fresh rumen fluid was collected from three ruminally fistulated dairy cows and combined as the source inoculum of an HAB enrichment culture. Following successive enrichment in a Casamino Acid medium, co- cultures were performed for the HAB enrichment culture and a culture of the ciliated protozoan, Entodinium caudatum, that contained both mixed bacteria and archaea. The co-culturing experiment was conducted with or without the feed substrate for protozoa and Micrococcus luteus, a strictly aerobic bacterium, to assess the impact of feeding the protozoan on ammonia production. Unexpectedly, ammonia concentrations were higher in E. caudatum alone treatments than in the co-cultures of HAB enrichment and protozoa irrespective of addition of a feed substrate for protozoa, with 20.5 ± 0.8, 18.5 ±

0.2, 23.9 ± 0.3, and 23.2 ± 1.1 mg ammonia N/dL in treatment groups E. caudatum with feed substrate, E. caudatum without feed substrate, E. caudatum with feed substrate and with M. luteus, and E. caudatum without feed substrate but with M. luteus, respectively. Ammonia concentration was also increased (P < 0.05) by M. luteus addition in the E. caudatum alone treatment group when no feed substrate was added.

However, the addition of feed substrate had no significant effect on ammonia concentration. E. caudatum counts were greater (P < 0.05) in all cultures receiving the feed substrate except for E. caudatum alone treatments receiving M. luteus addition; however, no significant differences in counts occurred with the addition of HAB to E. caudatum cultures. This study did not support a positive collaboration in ammoniagenesis between the HAB enrichment culture and the E. caudatum culture that carried contamination with bacteria and possibly archaea. A bacteria- and archaea-free

Entodinium caudatum culture is needed to investigate the interaction between protozoa and HAB.

INTRODUCTION

Animal agriculture is estimated to be responsible for 50 to 85% of the national ammonia emissions in the USA (Battye et al., 1994). A major contributor to this problem is the deamination of amino acids within the ruminal environment by microorganisms.

One such niche of microorganisms currently under review for their contribution to ruminal ammonia emissions is occupied by hyper-ammonia-producing bacteria (HAB).

HAB are bacteria that express a rapid rate of ammonia production with a minimal ability

34 to utilize sugars (Chen and Russel, 1989; Wallace, 1996; Whitehead and Cotta, 2004;

Flythe and Andries, 2009). HAB have been isolated from the rumen of cattle, sheep, and goats. Although HAB typically exist at low abundance inside the rumen, the amount of ammonia they are capable of producing make them quantitatively important. The known HAB species include Clostridium sticklandii, Clostridium aminophilum, and

Peptostreptococcus anaerobius.

Protozoa play an important role in ruminal function through interaction with other rumen microorganisms, and as such are capable of indirectly impacting ammoniagenesis. Protozoa, often considered the simplest eukaryotic life-form, consist of a mouth, a single digestive cavity, a cytoproct, rectum, macronucleus, micronucleus, and one or more contractile vacuoles all bounded by either a cuticle or pellicle

(Dehority, 2003).

Protozoa, in general, constitute the top of the tropic chain in the rumen and are present in large numbers. Ciliate protozoa have an abundance of 105 to 106 cells/mL of ruminal fluid (Ozutsumi et al., 2005; Sylvester et al., 2004). Due to their prominence, protozoa can directly modify functions of the guilds of ruminal microbes and, therefore, overall ruminal functions, including ammonia production. Protozoa are able to stimulate ammonia production by providing short peptides, which can be utilized by HAB.

Protozoa also ingest bacteria, fungi, and small feed particles and use them as a source of protein. These proteins are then degraded, yielding peptides and free amino acids.

Some of the proteolytic products are further utilized to synthesize protozoal protein,

35 but many peptides and amino acids also find their way back into ruminal fluid due to secretory processes as well as autolysis and death of the protozoa (Coleman, 1985;

Dijkstra, 1994).

Conceivably, protozoa and HAB can have two different types of interactions: 1) commensalism in which protozoa supply HAB with short peptides and amino acids as substrates for ammonia production and 2) predation in which protozoa prey on HAB.

The above interactions will have two different effects on ammoniagenesis in the rumen, with the former enhancing ammonia production by HAB, while the latter reduces ammonia production. However, how the interaction between protozoa and HAB affect ammonia production remains unexplored, particularly at the microbial community level.

As such, the primary objective of this study was to examine the interactions between

HAB as an enriched consortium and E. caudatum, in terms of ammonia production and protozoal counts. E. caudatum has demonstrated a lack of specificity in bacterial engulfment and as such was utilized for this study (Coleman, 1964; Coleman, 1972).

MATERIALS AND METHODS

HAB Enrichment

Briefly, 6 h after feeding, ruminal contents was collected from three ruminally fistulated dairy cows and enriched anaerobically in a Casamino Acid (30 g/L) medium and incubated at 39°C. Refer to chapter 2 Materials and Methods for more details on

HAB enrichment.

Micrococcus luteus Culture

M. luteus was grown in Tryptic Soy Broth medium aerobically at 35°C for 24 h.

The cell biomass was harvested by concentration and then resuspended in Casein

(15g/L) medium.

Entodinium caudatum Culture

The protozoan Entodinium caudatum was previously provided by Dr. Burk

Dehority. The culture was maintained in protozoal medium SP (Dehority, 1993). The culture was transferred twice a week. At each transfer, the culture was divided into two, and half of the culture was placed into fresh SP medium and continued as the stock culture. The other half was then either discarded or doubled in a similar fashion and also used as a stock culture if a larger protozoal culture volume was required. Substrate for the E. caudatum culture was a suspension containing 1.5% (w/v) starch and 1% (w/v) orchardgrass hay that was prepared in distilled water and gassed with O2-free CO2

(Dehority, 2003). Starch was included as ground wheat and rice flower 50:50, while orchardgrass was provided as powder. The E. caudatum culture contained no other protozoal species, but it contained bacteria and presumably archaea that propagate together with E. caudatum.

Co-cultures

Seventy-two hours prior to the experimental trial, 1% (v/v) of the enriched HAB culture was inoculated into the previously described Casamino Acid (30g/L) medium (pH of 6.5 to 6.6) in a Hungate tube capped with a rubber stopper and aluminum seal. The culture was then incubated at 39°C for 24 h. Following the 24 h incubation, 6 mL of the

24 h culture was further transferred into 54 mL of a less concentrated Casamino Acid

(15g/L) medium (pH of 6.5 to 6.6) in a serum bottle capped with a rubber stopper and aluminum seal and incubated at 39°C for 48 h. Series of cultures and co-cultures of HAB and E. caudatum were set up in three replicates as shown in Table 3.1. The spent SP medium and spent casein medium consisted of culture supernatant from a protozoal culture and a HAB culture, respectively, through centrifugation. The spent media were used to equalize the medium composition of all the treatments. All treatments were done in triplicate in Hungate tubes capped with rubber stoppers and aluminum seals. All the treatments were incubated anaerobically at 39 °C. After 12 h, each treatment was subsampled for ammonia analysis and then incubated for another 12 h.

Analysis

All the cultures were analyzed for total protozoa counts, pH, and ammonia concentration. The protozoa were microscopically enumerated as previously described by Dehority (1993). If samples were not analyzed for ammonia concentration immediately, they were stored at -20°C until further processing was possible. Ammonia

38 production was analyzed as repeated measures of a completely randomized design using the MIXED procedure of SAS 9.2 (SAS Inst. Inc., Cary, NC). Covariance structure selection was based on the Bayesian Information Criterion. Protozoal counts were log- transformed to improve normality and analyzed as a completely randomized design using the MIXED procedure of SAS 9.2 (SAS Inst. Inc., Cary, NC).

RESULTS AND DISCUSSION

Protozoal counts

At 24 h post incubation, E. caudatum numbers were significantly higher (p <

0.05) in all cultures receiving the feed substrate (Table A.1 and A.2). However, the addition of M. luteus to the cultures had no significant effect on protozoal counts compared to treatment groups that did not receive M. luteus (Figure 3.3). These results indicate that the presence of feed in the form of plant materials, but not necessarily M. luteus biomass, is capable of modifying protozoal activity and ultimately population size.

While M. luteus appeared to have little to no effect on E. caudatum numbers, this is not necessarily true of all microbial biomass. E. caudatum has been shown to have some preference over ingestion of certain bacteria, though preference appears limited when compared to other ruminal protozoa (Coleman and Sandford, 1979; Coleman, 1964;

Coleman, 1972). Transfer period may also have affected predation on M. luteus.

No significant differences in protozoal counts were seen when the HAB was added to E. caudatum cultures (Figure 3.3). This would suggest that the addition of HAB

39 do not directly impact the growth of E. caudatum. Even in co-cultures of HAB and E. caudatum receiving no feed substrate, protozoal numbers were not significantly different from E. caudatum cultures alone with no substrate, suggesting that E. caudatum did not utilize HAB for growth in the absence of the feed substrate. However, this assumption would need to be further supported by quantifying the population of the HAB cultures in each treatment.

Ammonia production

Several results became apparent from the HAB enrichment and E. caudatum cultures (Tables A.1 and A.2). This data was further analyzed in terms of ammonia production differences between 0 and 12 h (Figure 3.1) and 0 and 24 h (Figure 3.2), negating any initial differences in ammonia readings per tube.

Results at 12 h post incubation indicate (Figure 3.1) an increase (P < 0.05) in ammonia concentration in all treatments containing the E. caudatum culture except for the co-culture of HAB and E. caudatum without the protozoal feed added, in comparison to HAB alone treatment groups (with and without M. luteus addition). E. caudatum alone treatment groups with M. luteus addition had the highest ammonia concentration in the cultures with and without the feed substrate addition in comparison to all other treatment groups. Similarly, results at 24 h post incubation

(Figure 3.2) indicated no additive effect when E. caudatum and HAB were cultured together. Instead, ammonia concentration was higher (P < 0.05) in the E. caudatum

40 alone cultures when compared to the HAB alone groups. The E. caudatum cultures contained no other protozoan, but it contained bacteria and archaea. Although removal of these contaminants has been attempted in previous studies (Onodera and

Henderson, 1980; Morgavi et al., 1994; Fondevila and Dehority, 2001), no fully efficient or effective method has been available up to this point to remove the bacteria or archaea from the protozoal cultures. In this study, these bacterial populations could have deamination properties and lead to some, if not all, of the ammonia production seen in the E. caudatum treatments. However, protozoa are capable of ammonia production, so results could also reflect the deamination properties of E. caudatum.

Although it is of interest that the protozoal alone cultures resulted in high ammonia concentration, it is also of interest that co-cultures of enriched HAB and E. caudatum did not display an additive effect.

The M. luteus addition to the cultures also provided interesting results. By 12 h post incubation E. caudatum alone treatment receiving no feed substrate showed a difference (P < 0.05) in ammonia concentration between with and without M. luteus addition, with 5.3 ± 0.6 and 12.2 ± 0.4 mg N/dL respectively. Again, at 24 h, the E. caudatum alone treatment group receiving no feed substrate showed a difference (P <

0.05) in ammonia concentration between with and without M. luteus addition, with 18.5

± 0.2 and 23.2 ± 1.0 mg N/dL respectively. Axenic ruminal protozoal cultures free of both bacteria and archaea are unable, at the current time, to be successfully maintained for long periods of time due to both difficulties in removal bacteria, even with the use of

41 antibiotics (Coleman, 1962; Onodera and Henderson, 1980; Hino and Kametaka, 1977;

Bonhomme et al., 1982), and the need for bacteria to serve as an energy source or provide growth factors (Quinn et al., 1962; Rahman et al, 1964; Coleman et al., 1977).

These results would suggest that the addition of M. luteus affected the protozoal culture in some way and may provide a method for maintaining protozoal cultures in the absence of anaerobic rumen bacteria. However, the addition of feed substrate had no significant effect on ammonia concentration at either 12 or 24 h post incubation. These results indicate that the interactions among HAB and protozoa may be affected by the addition of M. luteus or microbial biomass, but not necessarily plant materials.

Overall, this study did not support a positive collaboration in ammoniagenesis between the HAB enrichment culture and the E. caudatum culture that carried contamination with bacteria and possibly archaea. As such, further studies need to be undertaken to remove contaminants in the E. caudatum cultures in order to examine if increased ammonia concentration in the protozoal alone cultures was due in fact to the

E. caudatum culture, or instead bacterial contaminants residing in the culture. Once this occurs, a better understanding can be obtained as to the interactions between HAB and protozoa, hopefully aiding in new interventions for the dairy industry to reduce nitrogen outputs from dairy facilities through these interactions.

Table 3.1. Treatment Groups.

Treatment Without M. luteus With M. luteus HAB alone 1.25 ml HAB 1.25 ml HAB 0.05 ml H2O 0.05 ml H2O 1.225 ml casein medium 1.025 ml casein medium 1.225 ml SP medium 1.225 ml SP medium 1.25 ml spent SP medium 1.25 ml spent SP medium 0.20 ml M. luteus HAB + E. caudatum 1.25 ml HAB 1.25 ml HAB with protozoal feed 1.25 ml E. caudatum 1.25 ml E. caudatum 0.05 ml feed substrate 0.05 ml feed substrate 1.225 ml casein medium 1.025 ml casein medium 1.225 ml SP medium 1.225 ml SP medium 0.20 ml M. luteus HAB + E. caudatum 1.25 ml HAB 1.25 ml HAB without protozoal 1.25 ml E. caudatum 1.25 ml E. caudatum feed 0.05 ml H2O 0.05 ml H2O 1.225 ml casein medium 1.025 ml casein medium 1.225 ml SP medium 1.225 ml SP medium 0.20 ml M. luteus E. caudatum alone 1.25 ml E. caudatum 1.25 ml E. caudatum with protozoal feed 0.05 ml feed substrate 0.05 ml feed substrate 1.225 ml casein medium 1.025 ml casein medium 1.225 ml SP medium 1.225 ml SP medium 1.25 ml spent Casein 1.25 ml spent casein medium medium 0.20 ml M. luteus E. caudatum alone 1.25 ml E. caudatum 1.25 E. caudatum without protozoal 0.05 ml H2O 0.05 ml H2O feed 1.225 ml casein medium 1.025 ml casein medium 1.225 ml SP medium 1.225 ml SP medium 1.25 ml spent casein 1.25 ml spent casein medium medium 0.20 ml M. luteus Experimental groups and inoculum for each as part of co-culture experiment. Each treatment was done in triplicate and all incubations occurred at 39°C with a pH of 6.5 to

6.6 in Hungate tubes capped with rubber stoppers and aluminum seals.

a ab bc bc bc cde c cde de e

Figure 3.1. Ammonia concentration in co-cultures 12 h post incubation. Each treatment was done in triplicate and all incubations occurred at 39°C with a pH of 6.5 to 6.6 in

Hungate tubes capped with rubber stoppers and aluminum seals. The data represent the mean values ± SEM; different letters denote significant differences (P < 0.05).

a a ab bc cd cd d d

e e

Figure 3.2. Ammonia concentration in co-cultures 24 h post incubation. Each treatment was done in triplicate and all incubations occurred at 39°C with a pH of 6.5 to 6.6 in

Hungate tubes capped with rubber stoppers and aluminum seals. The data represent the mean values ± SEM; different letters denote significant differences (P < 0.05).

a a ab acd a bc cd cd d

Figure 3.3. E. caudatum counts in co-cultures 24 h post incubation. Each treatment was done in triplicate and all incubations occurred at 39°C with a pH of 6.5 to 6.6 in Hungate tubes capped with rubber stoppers and aluminum seals. The data represent the mean values ± SEM; different letters denote significant differences (P < 0.05).

CHAPTER 4

INVESTIGATION INTO THE INHIBITION OF AMMONIA PRODUCTION BY A HYPER-

AMMONIA PRODUCING BACTERIAL CONSORTIUM BY PLANT SECONDARY

METABOLITES

ABSTRACT

Ammoniagenesis in the rumen is a wasteful process. Amino acid-fermenting bacteria, particularly hyper-ammonia-producing bacteria (HAB), are able to deaminate amino acids within the rumen, causing excess ammonia production. A recent source to modulate rumen function, and therefore ammonia production, are plant secondary metabolites. These secondary metabolites primarily interact with the bacterial cell membrane and have been explored for their ability to mitigate ammoniagenesis in the rumen using either single species of HAB or rumen microbial communities. The aim of this study was to evaluate their effectiveness at inhibiting ammonia production from amino acids by a consortium of enriched ammonia producers. In Experiment 1, plant secondary metabolites fenugreek and vanillin were used in doses of 1.0 g/L and carvacrol, clove oil, and origanum oil were used at 0.5 g/L in a medium containing

Casamino Acids (30 g/L) as sole substrate. In Experiment 2, fenugreek, vanillin, carvacrol, clove oil, and origanum oil were used in doses similar to those in experiment

1, but a feed substrate was added to evaluate its interaction with the secondary plant metabolites in affecting ammoniagenesis. The results indicate that fenugreek is least potent in inhibiting ammoniagenesis. Carvacrol, origanum oil, and clove oil significantly reduced (P < 0.01) ammonia concentration post incubation, regardless of the presence or absence of a feed substrate. While the feed addition did not significantly attenuate the inhibition of ammoniagenesis on origanum oil and carvacrol, feed addition showed some inhibition on the effectiveness of clove oil at both 12 h and 24 h. Also, when no feed substrate was present, vanillin significantly (P < 0.01) lowered ammonia at both 12 h and 24 h. However, when a feed substrate is supplied, vanillin significantly reduced (P

< 0.01) ammonia concentration at only 24 h post incubation.

INTRODUCTION

Excess ammonia emissions, of which animal agriculture is estimated to be responsible for 50 to 85% of ammonia emissions nationwide (Battye et al., 1994), are a major concern for the dairy industry due to the detrimental impact of ammonia emissions on the environment. Hyper-ammonia-producing bacteria, present within the rumen, are a major contributor to ammonia emissions emitted from dairy cattle. HAB are able to catabolize amino acids as well as short peptides, possibly provided by protozoa and other proteolytic bacteria, to produce ammonia. This ammonia is then able to undergo assimilation followed by further transamination for synthesis of cellular

48 protein by ruminant microorganisms. While this process is beneficial, concern arises when ammonia production in the rumen are in excess of that which can be utilized by the microbial population present in the host. When this situation occurs, ammonia is absorbed across the ruminal wall and converted to urea, which is excreted resulting in inefficient N retention (Leng and Nolan, 1984).

Many methods have been proposed to lower ammonia emissions, including the use of antibiotics, defaunation, and plant secondary metabolites. Monensin, an antibiotic classified as a carboxylic ionophore polyether, has been shown to have antimicrobial effects (Pressman and Fahim, 1982; Russell, 1996) as well as capabilities to lower ammoniagenesis within the rumen both in vitro and in vivo (Chen and Russell,

1989; Yang and Russell, 1993). However, studies have shown monensin is ineffective at reducing ammonia in cattle predominantly fed alfalfa hay (Lana and Russel, 1997).

Limited sensitivity by Gram-negative bacteria has also been shown (Russell, 1996), though certain Gram-negative species have still been found susceptible (Callaway and

Russell, 2000).

Defaunation reduced NH3-N concentrations (Jouany and Ushida, 1998; Eugène et al., 2004), thereby increasing protein-derived ammonia conversion to microbial protein and improving net protein availability to the animal (Ivan et al., 1992; Eugène et al.,

2004). However, the full extent as to the role protozoa play has yet to be completely elucidated and concerns have arisen as to the residual effects defaunating agents may

49 have on beneficial effects of protozoa on the rumen and overall microbial community

(Hristov and Jouany, 2005).

Recent studies have focused on alternative sources to those previously described to modulate rumen function, one of which is through the use of plant secondary metabolites. Plant species, through secondary metabolism, produce a variety of organic compounds, some of which exhibit antimicrobial activity against many bacteria, yeasts, and even molds (Gershenzon and Croteau, 1991). These secondary metabolites are often placed into one of three groups: saponins, tannins, and essential oils.

Saponins are a high-molecular-weight glycoside that contain saccharide chain units linked to either a triterpene or steroidal aglycone moiety (Patra and Saxena, 2009).

Although many studies have reported on the ability of saponins to reduce ammonia production in mixed ruminal cultures (Busquet et al., 2006; Patra and Yu, 2013), reduced concentrations are thought to be due to defaunation (Ivan et al., 2004; Baah, 2007;

Singer et al., 2008). Saponins interact with the sterol moiety present within the protozoal cell membrane (Hostettmann and Marston, 1995), ultimately causing leakage of cell contents (Patra and Saxena, 2009).

Essential oils are classified as secondary metabolites that are obtained by steam distillation from the plant volatile fraction (Gershenzon and Croteau, 1991). Essential oils contain two chemical groups, terpenoids and phenylpropanoids, which contain many key active compounds. Both terpenoids and phenylpropanoids use a mechanism

50 of action dependent on interaction with the cell membrane (Davidson and Naidu, 2000;

Dorman and Deans, 2000; Benchaar et al., 2008). The hydrophobic cyclic hydrocarbons interact with the cell membrane, resulting in their accumulation within the bacterial lipid bilayer (Sikkema et al., 1994; Ultee et al., 1999). Eventually, this accumulation will disrupt the structure of the membrane, causing expansion and fluidification (Griffin et al., 1999; Calsamiglia et al., 2010) and ultimately slowing down bacterial growth due to increased energy requirements for homeostasis (Cox et al., 2001). In the rumen environment, this process has decreased ammonia production both in vitro and in vivo

(Wallace et al., 2002; McIntosh et al., 2003; Giannenas et al., 2011; Patra and Yu, 2013).

As such, the primary objective of this study was to determine the effectiveness of select plant secondary metabolites at inhibiting our HAB enrichment culture and in turn ammonia production.

MATERIALS AND METHODS

HAB Enrichment

Briefly, 6 h after feeding, ruminal contents was collected from three ruminally fistulated dairy cows and enriched anaerobically in a Casamino Acid (30 g/L) medium and incubated at 39°C. Refer to the Materials and Methods of Chapter 2 for more details on HAB enrichment.

Plant secondary metabolites

Five different plant secondary metabolites, including fenugreek, carvacrol, clove oil, origanum oil, and vanillin, were purchased from Sigma-Aldrich (Sigma-Aldrich, St.

Louis, MO) and further utilized in this study. Each of the secondary metabolites was used at one of two doses depending on effectiveness determined in a previous study

(data not shown): fenugreek and vanillin were added to achieve a final concentration of

1.0 g/L and carvacrol, clove oil, and origanum oil were added to a final concentration of

0.5 g/L. A control containing no secondary metabolites was used in parallel.

In vitro Incubations: Experiment 1

One ml of the enriched HAB culture was added to 9.0 ml of a Casamino Acid (30 g/L) medium. In addition, fenugreek and vanillin were added at 1.0 g/L and carvacrol, clove oil, and origanum oil were added at 0.5 g/L. Treatments were incubated in triplicate in Hungate tubes capped with rubber stoppers and aluminum seals. All incubations occurred anaerobically for 24 h at 39°C. Each culture was subsampled for ammonia analysis at 0 h, 12 h, and 24 h.

In vitro Incubations: Experiment 2

Incubations were carried out anaerobically in Hungate tubes capped with rubber stoppers and aluminum seals in triplicate for each dose of each secondary metabolite and the control. Substrate consisted of a ground alfalfa hay and a dairy concentrate

52 mixture (consisting mainly of ground corn [33.2%], soybean meal [14.2%]), AminoPlus

(Ag Processing Inc., USA) [15.5%], distillers’ grains [19.8%], wheat middlings [11.3%]), in a ratio of 50:50 were used as the substrate. 1% (v/v) of the enriched HAB culture was added to 9.0 ml of a Casamino Acid (30 g/L) medium. Fenugreek and vanillin were added at 1.0 g/L and carvacrol, clove oil, and origanum were added at 0.5 g/L. 1.0 g/L of the feed substrate was added to each culture. Each culture was subsampled for ammonia analysis at 0 h, 12 h, and 24 h.

Analysis

Ammonia concentrations were analyzed using the phenol-hypochlorite reaction as defined by Chaney and Marbach (1961). The pH in each culture at the end of 24 h incubation was also determined using a pH meter. If samples were not analyzed immediately, they were stored at -20°C until further processing was possible.

The ability of each secondary metabolite to limit ammonia production was analyzed as repeated measures of a completely randomized design using the MIXED procedure of SAS 9.2 (SAS Inst. Inc., Cary, NC).

RESULTS AND DISCUSSION

Experiment 1

Table A.3 displays total ammonia concentration 12 h and 24 h post incubation.

This data was further analyzed in terms of ammonia concentration differences between

0 and 12 h (Figure 4.1) and 0 and 24 h (Figure 4.2), subtracting the initial ammonia concentration readings per tube.

Carvacrol, clove oil, and origanum oil, when used at a dose of 0.5 g/L, decreased ammonia concentrations in the enriched HAB culture at both 12 h and 24 h post incubation. By 24 h post incubations, ammonia readings increased in carvacrol and origanum treatments, but remained fairly steady in the clove oil treatment, with 1.45 ±

0.39, 0.18 ± 0.07, and 0.71 ± 0.09 mg ammonia N/dL for carvacrol, clove oil, and origanum oil, respectively.

Carvacrol is a phenolic monoterpenoid found in oregano oil and several other essential oils in high quantities. Previous studies have shown that carvacrol is a broad spectrum antimicrobial (Friedman et al., 2002). Delocalized electrons on the benzene ring and the presence of a hydroxyl group are thought to give carvacrol its antimicrobial activity, allowing it to target the bacterial membrane and ultimately dissipate proton motive force and electron flow across the bacterial membrane (Helander et al., 1998;

Ultee et al., 1999). Carvacrol decreases ammonia concentrations in vitro (Macheboeuf et al., 2008; Busquet et al., 2006) and may indeed be due to the inhibition of HAB as the results from this study suggest.

Origanum oil contains a high content level of carvacrol and thymol and inhibits a broad spectrum of bacterial species (Barrata et al., 1998; Lis-Balchin and Deans, 1997).

Thymol, similar to other essential oils, is capable of altering cell membrane permeability.

Origanum oil, as a result, also has been shown to retain the ability to decrease ammonia concentrations in vitro (Macheboeuf et al., 2008; Patra and Yu, 2013).

Eugenol contributes to the antimicrobial capabilities of clove oil. As a phenylpropanoid, eugenol contains an aromatic ring of 6 carbons. Similar to terpenes, eugenol is capable of penetrating the lipopolysaccharide layer, altering cell structure and causing leakage of the cell membrane. Clove oil has inhibited ammonia from mixed ruminal cultures in vitro (Patra and Yu, 2013; Busquet et al., 2006).

Vanillin, used at a higher dose of 1.0 g/L due to limited inhibition during an initial test, still significantly reduced (P < 0.01) ammonia concentration at both 12 h and 24 h.

Vanillin is the primary extract found in vanilla beans (Vanilla planifola, Vanilla pompon, and Vanilla tahitensis; Davidson and Naidu, 2000). Although resulting in almost a 90% reduction in ammonia at 12 h post incubation, by 24 h ammonia concentration by the vanillin treatment was only reduced by no more than 60% as compared to the control.

Similarly, a previous study on vanillin in mixed ruminal cultures reported a 40% reduction in ammonia 24 h post incubation (Patra and Yu, 2013). Interestingly, in that study, the abundance of C. sticklandii was found unaffected while the abundance of C. aminophilum actually increased, despite reduction in ammonia. This may suggest bacteriostatic activity (Fitzgerald et al., 2004).

Fenugreek had no significant effect on ammonia production by 24 h post incubation. In mixed ruminal cultures however, fenugreek significantly reduced

55 ammonia concentration 24 h post incubation (Busquet et al., 2006). However, the effectiveness of fenugreek has often been attributed to its antiprotozoal tendencies.

Fenugreek has high steroidal saponin content and as such is able to interact with the sterol moiety found within the protozoal membrane, destroying the membrane and leading to leakage of cell contents (Patra and Saxena, 2009). As such, because no protozoa were present within the enriched HAB culture, fenugreek may not have displayed a direct antimicrobial effect on the HAB such as might occur indirectly in a mixed ruminal culture through inhibition of protozoa.

These results indicate that varying plant secondary metabolites are capable of affecting ammoniagenesis. A significant decrease in concentrations of ammonia (p <

0.01), as indicated in cultures of carvacrol, clove oil, and origanum oil at both 12 h and

24 h post incubation in HAB enriched cultures suggests that the inhibition of ammonia as seen in previous in vitro studies on mixed ruminal cultures is probably due to susceptibility of HAB to plant secondary metabolites. This premise is supported by the work of McIntosh et al. (2003) who found C. sticklandii and P. anaerobius, two known

HAB, are highly susceptible to essential oils.

Experiment 2

Previous studies have drawn attention to the concept that not all plant secondary metabolites that prove affective in vitro are necessarily still affective in vivo.

One reason currently thought to limit a plant secondary metabolite’s effectiveness in

56 vivo is the presence of feed particles within the rumen. Recent researchers have hypothesized that components within essential oils, such as carvacrol, thymol, and eugenal, which lend antimicrobial qualities to essential oils may bind to fats as well as other hydrophobic materials present in feeds, preventing the components from binding to target bacteria (Si et al., 2006). As such, it is important to understand if in vitro studies on secondary metabolites as antimicrobials are applicable within the rumen.

Results from experiment 2 provided evidence for the effectiveness of certain secondary metabolites in the presence of feed particles.

When the feed substrate was provided to HAB enriched cultures that received the different plant secondary metabolites, several results became apparent (Table A.4).

The data was further analyzed in terms of ammonia production differences between 0 and 12 h (Figure 4.3) and 0 and 24 h (Figure 4.4), subtracting the initial ammonia concentration readings per tube.

Carvacrol, clove oil, and origanum oil, when used at a dose of 0.5 g/L significantly

(p < 0.01) inhibited ammonia by the enriched HAB culture containing a feed substrate at both 12 h and 24 h post incubation. Vanillin, used at a higher dose of 1.0 g/L reduced (P

< 0.01) ammonia concentration at only 24 h. However, fenugreek, used at 1.0 g/L, still had no significant effect on ammonia concentration.

Overall, although the feed addition did not significantly attenuate the inhibition of ammoniagenesis on origanum oil and carvacrol, feed addition showed some

57 inhibition on the effectiveness of clove oil at both 12 h and 24 h post incubation. Also, when no feed substrate was present, vanillin significantly (P < 0.01) lowered ammonia at both 12 h and 24 h. However, when the feed substrate was supplied, vanillin significantly reduced (P < 0.01) ammonia concentration only at 24 h post incubation.

These results lend some credibility to the application of in vitro studies on plant secondary metabolites to the actual ruminant environment. Depending on the plant secondary metabolite to be used, it may be possible to reduce nitrogen outputs from dairy facilities.

b b b b

Figure 4.1. Ammonia concentration in plant secondary metabolite treatments 12 h post incubation. Each treatment was done in triplicate and all incubations occurred at 39°C with a pH of 6.5 to 6.6 in Hungate tubes capped with rubber stoppers and aluminum seals. Doses consisted of 1.0 g/L for fenugreek and vanillin and 0.5 g/L for carvacrol, clove oil, and origanum. The data represent the mean values ± SEM; different letters denote significant differences (P < 0.01).

a b a b

c c c

Figure 4.2. Ammonia concentration in plant secondary metabolite treatments 24 h post incubation. Each treatment was done in triplicate and all incubations occurred at 39°C with a pH of 6.5 to 6.6 in Hungate tubes capped with rubber stoppers and aluminum seals. Doses consisted of 1.0 g/L for fenugreek and vanillin and 0.5 g/L for carvacrol, clove oil, and origanum. The data represent the mean values ± SEM; different letters denote significant differences (P < 0.01).

ab a

d d

Figure 4.3. Ammonia concentration in plant secondary metabolite treatments with feed substrate 12 h post incubation. Each treatment was done in triplicate and all incubations occurred at 39°C with a pH of 6.5 to 6.6 in Hungate tubes capped with rubber stoppers and aluminum seals. Doses consisted of 1.0 g/L for fenugreek and vanillin and 0.5 g/L for carvacrol, clove oil, and origanum. 1.0 g/L of the feed substrate was added to each culture. The data represent the mean values ± SEM; different letters denote significant differences (P < 0.01).

a b a b

b b b

c c b b

Figure 4.4. Ammonia concentration in plant secondary metabolite treatments with feed substrate 24 h post incubation. Each treatment was done in triplicate and all incubations occurred at 39°C with a pH of 6.5 to 6.6 in Hungate tubes capped with rubber stoppers and aluminum seals. Doses consisted of 1.0 g/L for fenugreek and vanillin and 0.5 g/L for carvacrol, clove oil, and origanum. 1.0 g/L of the feed substrate was added to each culture. The data represent the mean values ± SEM; different letters denote significant differences (P < 0.01).

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APPENDIX A: EXTRANEOUS TABLES AND FIGURES

e d de c b a a a a

Figure A.1 Growth curve of HAB3 when grown on Casamino Acids. Substrate was used at

1.5% (w/v) concentration. Incubations occurred at 39°C with a pH of 6.5 to 6.6 in

Hungate tubes capped with rubber stoppers and aluminum seals. Treatment was done in triplicate and the data represents the mean values; different letters denote significant differences (P < 0.05).

c c bc b a a a

Figure A.2. Growth curve of HAB3 when grown on Casein. Substrate was used at 1.5%

(w/v) concentration. Incubations occurred at 39°C with a pH of 6.5 to 6.6 in Hungate tubes capped with rubber stoppers and aluminum seals. Treatment was done in triplicate and the data represents the mean values; different letters denote significant differences (P < 0.05).

e de d

b a a

Figure A.3. Growth curve of HAB3 when grown on Cellobiose. Substrate was used at

1.5% (w/v) concentration. Incubations occurred at 39°C with a pH of 6.5 to 6.6 in

Hungate tubes capped with rubber stoppers and aluminum seals. Treatment was done in triplicate and the data represents the mean values; different letters denote significant differences (P < 0.05).

c bc bc

abc

ab a a

Figure A.4. Growth curve of HAB3 when grown on Control. Substrate was used at 1.5%

d d d c

a a a

Figure A.5. Growth curve of HAB3 when grown on Glucose. Substrate was used at 1.5%

e d d

a a a

Figure A.6. Growth curve of HAB3 when grown on Maltose. Substrate was used at 1.5%

c b b

a a a a a

Figure A.7. Growth curve of HAB3 when grown on Starch. Substrate was used at 1.5%

e d

c b a a a

Figure A.8. Growth curve of HAB3 when grown on Xylose. Substrate was used at 1.5%

100

e d d

c a b

a a

Figure A.9. Growth curve of HAB5 when grown on Casamino Acids. Substrate was used at 1.5% (w/v) concentration. Incubations occurred at 39°C with a pH of 6.5 to 6.6 in

Hungate tubes capped with rubber stoppers and aluminum seals. Treatment was done in triplicate and the data represents the mean values; different letters denote significant differences (P < 0.05).

101

c c d b b a a a

Figure A.10. Growth curve of HAB5 when grown on Casein. Substrate was used at 1.5%

102

c c b ab a a a

Figure A.11. Growth curve of HAB5 when grown on Cellobiose. Substrate was used at

1.5% (w/v) concentration. Incubations occurred at 39°C with a pH of 6.5 to 6.6 in

Hungate tubes capped with rubber stoppers and aluminum seals. Treatment was done in triplicate and the data represents the mean values; different letters denote significant differences (P < 0.05).

103

c c bc bc b

a a

Figure A.12. Growth curve of HAB5 when grown on Control. Substrate was used at 1.5%

104

e d c b a a a

Figure A.13. Growth curve of HAB5 when grown on Glucose. Substrate was used at 1.5%

105

d cd c b b a a a

Figure A.14. Growth curve of HAB5 when grown on Maltose. Substrate was used at 1.5%

106

a a a a a a a

Figure A.15. Growth curve of HAB5 when grown on Starch. Substrate was used at 1.5%

107

c b

b a a a a

Figure A.16. Growth curve of HAB5 when grown on Xylose. Substrate was used at 1.5%

108

d d c c a

b a a

Figure A.17. Growth curve of HAB7 when grown on Casamino Acids. Substrate was used at 1.5% (w/v) concentration. Incubations occurred at 39°C with a pH of 6.5 to 6.6 in

Hungate tubes capped with rubber stoppers and aluminum seals. Treatment was done in triplicate and the data represents the mean values; different letters denote significant differences (P < 0.05).

109

c c c bc a b a a a

Figure A.18. Growth curve of HAB7 when grown on Casein. Substrate was used at 1.5%

110

d d

b b a a

Figure A.19. Growth curve of HAB7 when grown on Cellobiose. Substrate was used at

1.5% (w/v) concentration. Incubations occurred at 39°C with a pH of 6.5 to 6.6 in

Hungate tubes capped with rubber stoppers and aluminum seals. Treatment was done in triplicate and the data represents the mean values; different letters denote significant differences (P < 0.05).

111

b b b b ab a a a

Figure A.20. Growth curve of HAB7 when grown on Control. Substrate was used at 1.5%

112

e d d

b a a

Figure A.21. Growth curve of HAB7 when grown on Glucose. Substrate was used at 1.5%

113

d c bc ab a a

Figure A.22. Growth curve of HAB7 when grown on Maltose. Substrate was used at 1.5%

114

b a ab a a a

Figure A.23. Growth curve of HAB7 when grown on Starch. Substrate was used at 1.5%

115

d c bc ab a a a

Figure A.24. Growth curve of HAB7 when grown on Xylose. Substrate was used at 1.5%

116

d cd c c a

a a

Figure A.25. Growth curve of HAB8 when grown on Casamino Acids. Substrate was used at 1.5% (w/v) concentration. Incubations occurred at 39°C with a pH of 6.5 to 6.6 in

Hungate tubes capped with rubber stoppers and aluminum seals. Treatment was done in triplicate and the data represents the mean values; different letters denote significant differences (P < 0.05).

117

c cd d b b a a a

Figure A.26. Growth curve of HAB8 when grown on Casein. Substrate was used at 1.5%

118

de d e

b a a

Figure A.27. Growth curve of HAB8 when grown on Cellobiose. Substrate was used at

1.5% (w/v) concentration. Incubations occurred at 39°C with a pH of 6.5 to 6.6 in

Hungate tubes capped with rubber stoppers and aluminum seals. Treatment was done in triplicate and the data represents the mean values; different letters denote significant differences (P < 0.05).

119

b b b b

a a

Figure A.28. Growth curve of HAB8 when grown on Control. Substrate was used at 1.5%

120

d d d c

a a

Figure A.29. Growth curve of HAB8 when grown on Glucose. Substrate was used at 1.5%

121

e d d

c b a

Figure A.30. Growth curve of HAB8 when grown on Maltose. Substrate was used at 1.5%

122

d c b a a a a

Figure A.31. Growth curve of HAB8 when grown on Starch. Substrate was used at 1.5%

123

e e

c b a

Figure A.32. Growth curve of HAB8 when grown on Xylose. Substrate was used at 1.5%

124

Table A.1. Effects of HAB and E. caudatum co-cultures without M. luteus on pH, ammonia production, and protozoal counts in vitro.

Ammonia concentration 24 h Treatment 24 h pH Protozoal O h 12 h 24 h Counts HAB 12.22 13.18 15.24 7.50 0 11.96 14.60 15.59 7.47 0 11.19 15.09 15.76 7.53 0

HAB + E. 10.88 18.16 22.86 7.50 1.18 x 104 caudatum 11.74 17.61 23.87 7.44 1.26 x 104 with feed 11.17 17.68 22.29 7.48 1.23 x 104

HAB + E. 7.80 13.83 23.97 7.43 6.72 x 103 caudatum 8.90 10.36 24.90 7.47 7.2 x 103 without feed 7.77 12.27 23.25 7.45 5.87 x 103

E. caudatum 8.18 15.65 29.43 7.45 1.09 x 104 with feed 8.51 16.99 29.71 7.46 1.06 x 104 8.78 14.71 27.71 7.50 9.76 x 103

E. caudatum 9.06 14.55 28. 02 7.49 7.36 x 103 without feed 9.52 13.70 27.85 7.48 7.04 x 103 9.38 15.65 27.68 7.45 6.96 x 103

Each treatment was done in triplicate and all incubations occurred at 39°C with a pH of

6.5 to 6.6 in Hungate tubes capped with rubber stoppers and aluminum seals. See Table

3.1 for treatments.

125

Table A.2. Effects of HAB and E. caudatum co-cultures with M. luteus on pH,

ammonia production, and protozoal counts in vitro.

Ammonia concentration 24 h Treatment 24 h pH Protozoal O h 12 h 24 h Counts HAB 13.37 13.70 19.40 7.44 0 12.96 14.07 17.03 7.49 0 13.08 13.90 15.89 7.54 0

HAB + E. 7.41 16.20 22.53 7.53 1.22 x 104 caudatum 8.35 14.69 23.69 7.46 1.14 x 104 with feed 9.73 14.31 26.06 7.58 1.17 x 104

HAB + E. 8.59 14.25 23.42 7.56 8.64 x 103 caudatum 9.57 14.93 25.10 7.50 8.40 x 103 without feed 13.59 15.38 25.72 7.58 9.28 x 103

E. caudatum 7.51 18.51 31.97 7.58 1.20 x 104 with feed 9.44 18.75 33.00 7.33 1.12 x 104 8.85 19.64 32.59 7.40 1.04 x 103

E. caudatum 9.01 20.64 30.94 7.41 8.64 x 103 without feed 8.46 21.36 30.85 7.41 7.68 x 103 9.06 21.12 34.34 7.41 7.68 x 103

Each treatment was done in triplicate and all incubations occurred at 39°C with a pH of

6.5 to 6.6 in Hungate tubes capped with rubber stoppers and aluminum seals. See Table

3.1 for treatments.

126

TABLE A.3. Effects of plant secondary metabolites on ammonia production and pH in vitro.

0 h NH3-N 12 h NH3-N 24 h NH3-N Treatment Concentration Concentration Concentration 24 h pH (mg N/dL) (mg N/dL) (mg N/dL) Fenugreek 0.53 4.64 14.37 6.85 0.70 5.60 12.57 6.75 0.43 6.64 14.66 6.80

Carvacrol 0 .29 0 .84 1.49 6.67 0.57 0.84 2.79 6.64 0.34 0.43 1.28 6.65

Clove Oil 0.33 0 .33 0.38 6.59 0.29 0.44 0.50 6.60 0.16 0.41 0.45 6.62

Origanum 0.33 0 .45 1.22 6.64 0.31 0.53 0.94 6.65 0.33 0.53 0.94 6.63

Vanillin 0.48 1.04 6.04 6.77 0.55 1.35 8.62 6.76 0.45 0.91 5.29 6.72

Control 0.50 5.25 14.93 6.90 0.43 6.21 16.39 6.95 0.46 7.94 16.05 6.97

Each treatment was done in triplicate and all incubations occurred at 39°C with a pH of 6.5 to 6.6 in Hungate tubes capped with rubber stoppers and aluminum seals.

Doses consisted of 1.0 g/L for fenugreek and vanillin and 0.5 g/L for carvacrol, clove bud oil, and origanum. 1.0 g/L of the feed substrate was added to each culture.

127

TABLE A.4. Effects of plant secondary metabolites with feed substrate on ammonia production and pH in vitro.

0 h NH3-N 12 h NH3-N 24 h NH3-N Treatment Concentration Concentration Concentration 24 h pH (mg N/dL) (mg N/dL) (mg N/dL) Fenugreek 0.38 6.03 11.60 6.80 0.35 6.61 10.69 6.81 0.19 6.15 13.13 6.84

Carvacrol 0.05 1.06 1.79 6.64 0.03 0.86 0.85 6.64 0.17 0.65 0.75 6.62

Clove Oil 0.33 2.48 5.95 6.78 0.15 2.60 6.90 6.81 0.27 3.66 4.76 6.73

Origanum 0.12 0.42 1.39 6.20 0.03 0.15 0.95 6.61 0.15 0.45 0.92 6.59

Vanillin 0.42 4.64 6.54 6.77 0.55 4.47 6.72 6.78 0.35 3.68 4.91 6.71

Control 0.35 5.68 15.09 6.91 0.21 4.47 15.17 6.95 0.20 5.62 12.59 6.88

Each treatment was done in triplicate and all incubations occurred at 39°C with a pH of 6.5 to 6.6 in Hungate tubes capped with rubber stoppers and aluminum seals.

Doses consisted of 1.0 g/L for fenugreek and vanillin and 0.5 g/L for carvacrol, clove bud oil, and origanum. 1.0 g/L of the feed substrate was added to each culture.

128