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NOVEL STRATEGIES AND COMPOUNDSTO DECREASE RUMINAL
METHANOGENESIS IN VITRO

By

Emilio M. Ungerfeld

A DISSERTATION
Submitted to
Michigan State University
in partial fiilfillment of the requirements
for the degree of
DOCTOR OF PHILOSOPHY
Department of Animal Science

2003

ABSTRACT

NOVEL STRATEGIES AND COMPOUNDS TO DECREASE RUMINAL
METHANOGENESIS IN VITRO

By

Emilio M. Ungerfeld

Novel strategies and compounds to inhibit methane (CI-I4) formation in the rumen
were evaluated. Inhibition when attempting the inhibition of pyruvate oxidative
decarboxylation was small.

Propynoic acid and ethyl 2-butynoate decreased CH4 production, although they
decreased apparently fermented OM (F OM) and resulted in accumulation of H2, formate,
and ethanol. In contrast, B-hydroxybutyrate, crotonate, and 3-butenoic acid increased
FOM.

Aphidicolin and 3-bromopropanesulfonate did not affect CH4 formation.
Lumazine and a novel hexadecatrienoic acid.decreased CH4 production but decreased
FOM. Olive oil did not decrease CH4 production, but increased propionate production
without affecting FOM.

Combinations of lumazine, propynoic acid, and ethyl 2-butynoate with crotonic
acid or 3-butenoic acid were hypothesized to improve FOM and decrease accumulation
of H2, formate and ethanol. Crotonic acid and 3-butenoic acid were ineffective in
decreasing H2, formate, and ethanol and improving degradation. Lumazine, propynoic
acid, and ethyl 2-butynoate decreased N degradation and increased microbial OM and N
production and synthetic efficiencies. Propynoic acid and the highest concentration of

ethyl 2-butynoate also decreased OM and NDF degradation.

Differences in sensitivity to inhibitors were found among three ruminal
methanogens Methanobrevibacter ruminantium was the most sensitive to 2-
bromoethanesulfonate, propynoic acid, and ethyl 2-butynoate, Methanosarcina mazei
was the least sensitive to those chemicals, and Methanomicrobium mobile was
intermediate. M. ruminantium was the least sensitive to lumazine.

Lumazine caused mild and variable decreases in CH4 production. It did not
impair OM or NDF degradation, and increased microbial N and OM production. It
decreased proteolysis, which can increase N retention and decrease N release into the
envirOnment. Use of propynoic acid in ruminant diets could be problematic because of
potential toxicity. However, it is of interest to understand how it improved microbial
synthetic efliciencies. Inclusion of ethyl 2-butynoate in the diet to decrease CH4
production by about 50% might not affect OM and NDF degradation, decrease
proteolysis and increase microbial N flow. Ethyl 2-butynoate toxicity has not been
investigated, but it is advantageous that it disappeared after 24 h of incubation. The
greatest problem is to rechannel electrons away from H2, formate, and ethanol into

nutritionally useful sinks.

ACKNOWLEDGMENTS

To Steve Rust, for giving me an opportunity to study here, and fieedom to do research in .
my area of interest.

Committee members Mike Allen, Dave Beede, Mel Yokoyama and Rawle Hollingsworth
for their friendship and support.

Bob Burnett, for his help during all this time.

Dave Main, Dewey Longuski, Jackie Ying, Jenn Voelker, Seon-Woo Kim, Sue
Hengemuehle, Jim Liesman, Rob Tempelman, for their valuable help with lab work and
statistical analyses.

Jackie Christie and Nancy Perkins for their assistance.

Greg Zeilcus, Maris Laviekis, Claire Ville, Nathan Riddle, Jake McKinley and Dave
Finkelstein for their help with gas analyses.

David Boone and Yitai Liu for helping me to grow methanogens. Also Stephen
Ragsdale, C. A. Reddy, Joel Graber, Keith Joblin, Marcel Grapp for their useful advice.
Tom Herdt, Justin Zyskowski for their help with gas chromatography.

Family support throughout these years.

Friends: Department, Auld Hooligans Footie & Drinking Club, roomates Katie, Scott,
Chris, Vincent, Kamila, Gaston, Roger, Thor, Sidney, Tink, MSU Outing Club, other

MSU and US, Uruguay, UK, other countries

iv

TABLE OF CONTENTS

LIST OF TABLES .............................................................................. x
LIST OF FIGURES .............................................................................. xv
INTRODUCTION ................................................................................ 1
LITERATURE REVIEW ......................................................................... 6
Methane Production in the Rurnen ..................................................... 6
Methane Production as an Energy Loss .............................................. 13
Methane Emissions by Ruminants and Global Warming .......................... 15
Biochemistry and Energetics of Methanogenesis ................................... 20
Control of Methane Production in the Rurnen ....................................... 22
Dietary Manipulation .......................................................... 22
Chemical Additives ............................................................ 25
Microbial Additives ............................................................ 35
Others ............................................................................ 36
Conclusions ............................................................................... 37
References ................................................................................. 3 8
CHAPTER 2
Attempts to Decrease Ruminal Methanogenesis through the Inhibition of Pyruvate
Oxidative Decarboxylation ..................................................................... 46
Abstract ................................................................................... 46
Introduction .............................................................................. 47
Materials and Methods ................................................................. 48
Experiment 1 ................................................................... 48
Experiment 2 ................................................................... 53
Experiment 3 ................................................................... 54
Experiment 4 ................................................................... 55
Results and Discussion ................................................................. 56
Experiment 1 ................................................................... 56
Experiment 2 ................................................................... 62
Experiment 3 ................................................................... 65
Experiment 4 ................................................................... 67
Implications .............................................................................. 69
References ................................................................................ 7O

CHAPTER 3
Use of Some Novel Alternative Electron Sinks to Inhibit Ruminal Methanogenesis 73

Abstract .................................................................................... 73
Introduction ............................................................................... 74
Materials and Methods .................................................................. 75
Additives and Concentrations ................................................. 75
Ruminal Fluid Collection and Incubation .................................... 76
Analytical Procedures .......................................................... 77
Calculations ...................................................................... 78
Statistical Analysis .............................................................. 79
Results ...................................................................................... 80
Oxaloacetic Acid ................................................................ 80
Acetoacetate ..................................................................... 82
B-hydroxybutyrate ............................................................... 84
Crotonic Acid ..................................................................... 86
Propynoic Acid .................................................................. 88
3-Butenoic Acid ................................................................. 90
2-Butynoic Acid ................................................................. 92

Ethyl 2-butynoate ............................................................... 94
Discussion ................................................................................. 96
Oxaloacetate and Butyrate Enhancers ........................................ 96
Unsaturated Organic Acids and Esters ....................................... 98
Implications .............................................................................. 101
References ................................................................................ 103

CHAPTER 4

Some Miscellaneous Inhibitors of Ruminal Methanogenesis In Vitro .................... 106
Abstract ................................................................................... 106
Introduction .............................................................................. 106
Materials and Methods ................................................................. 108
Incubations ..................................................................... 108
Treatments ...................................................................... 108
Analysis ......................................................................... 109
Calculations ..................................................................... 1 10
Statistical Analysis ............................................................. 110
Results and Discussion ................................................................. 110
Aphidicolin ...................................................................... 110
Lumazine ........................................................................ 114

BPS ............................................................................... 117
Conclusions .............................................................................. 120
References ................................................................................ 121

vi

CHAPTER 5

Effects of Two Oils on In Vitro Ruminal Methane Production ............................ 124
Abstract ................................................................................... 124
Introduction .............................................................................. 125
Materials and Methods ................................................................. 126

Oils and Concentrations ...................................................... 126
Ruminal Fluid Collection and Incubation .................................. 126
Analytical Procedures ......................................................... 127
Calculations ..................................................................... 128
Statistical Analysis ............................................................. 128
Results and Discussion .................................................................. 128
Conclusions .............................................................................. 134
References ................................................................................ 136

CHAPTER 6

Effects of combinations of inhibitors of methanogenesis with crotonic acid or 3-butenoic

acid On in vitro ruminal fermentation and methane production ............................. 138
Abstract ................................................................................... 138
Introduction .............................................................................. 139
Materials and Methods ................................................................. 140

Experiment 1 ............................. , ...................................... 140
Experiment 2 ................................................................... 142
Experiment 3 ................................................................... 142
Experiment 4 ................................................................... 143
Experiment 5 ................................................................... 143
Experiment 6 ................................................................... 144
Results .................................................................................... 145
Experiment 1 ................................................................... 145
Experiment 2 ................................................................... 147
Experiment 3 ................................................................... 149
Experiment 4 ................................................................... 152
Experiment 5 ................................................................... 155
Experiment 6 ................................................................... 157
Discussion ............................................................................... 157
Lumazine ........................................................................ 157
Propynoic Acid .................................................................. 162
Ethyl 2-Butynoate .............................................................. 165
Crotonic Acid ................................................................... 166
3-Butenoic Acid ................................................................ 167
Conclusions .............................................................................. 169
References ............................................................................... 171

vii

CHAPTER 7
Effects of Combinations of Inhibitors of Methanogenesis with Crotonic Acid or 3-
Butenoic Acid on In Vitro Degradation and Microbial Biomass and N Synthesis 173

Abstract ................................................................................... 173
Introduction .............................................................................. 174
Materials and Methods ................................................................. 175
Experiment 1 ................................................................... 175
Experiment 2 ................................................................... 178
Experiment 3 ................................................................... 179
Experiment 4 ................................................................... 180
Experiment 5 ................................................................... 180
Experiment 6 ................................................................... 181
Results .................................................................................... 182
Experiment 1 .................................................................... 182
Experiment 2 .................................................................... 185
Experiment 3 .................................................................... 188
Experiment 4 .................................................................... 190
Experiment 5 .................................................................... 193
Experiment 6 .................................................................... 196
Discussion ................................................................................ 198
Conclusions .............................................................................. 205
References ............................................................................... 207
CHAPTER 8
Effects of Several Inhibitors on Pure Cultures of Ruminal Methanogens ................ 210
Abstract ................................................................................... 210
Introduction .............................................................................. 210
Materials and Methods .................................................................. 212
Cultures .......................................................................... 212
Medium Preparation ............................................................ 212
Chemicals ....................................................................... 213
Inoculation and Incubation ................................................... 214
Measurements and Calculations ............................................. 214
Statistical Analysis ............................................................ 214
Results and Discussion ................................................................ 215
BES .............................................................................. 215
BPS .............................................................................. 219
Lumazine ....................................................................... 220
Propynoic Acid ................................................................ 221
Ethyl 2-Butynoate .............................................................. 223
Conclusions .............................................................................. 224
References ................................................................................ 226
CONCLUSIONS ................................................................................ 230

viii

APPENDICES ................................................................................. 244

Unprocessed data ..................................................................... 245
Chemical structures and physico-chemical properties of compounds
studied ........................................................................ 304
References ..................................................................... 308

ix

LIST OF TABLES

Table 1-1. Ruminal methanogens ..................................................................... 10
Table 1-2 Sources of methane emissions ................................................... 17
Table 1-3 Estimates of global methane emissions by mammals .......................... 18
Table 1-4 Estimates of methane production by US cattle ................................... 19
Table 1-5 Partial reactions of methanogenesis and free energy changes .................. 21
Table 2-1. Ruminal fluid-free medium used in Experiment 1 .............................. 50

Table 2-2. Effects of thiamin, amproliurn, adenine and adenosine on end products of
isolated ruminal bacteria fermentation (Exp. 1) ............................................... 57

Table 2-3. Effects of adenine, adenosine and ribose on end products of ruminal in vitro
fermentation (Exp. 2) ............................................................................. 63

Table 2-4. Effect of thiamin structural analogs on ruminal in vitro fermentation
(Exp. 3) ............................................................................................. 66

Table 2-5 Effect of pyruvate derivatives on end products of in vitro ruminal
fermentation (Exp. 4) ............................................................................. 68

Table 3-1. Effects of the addition of oxaloacetic acid on in vitro ruminal
fermentation ........................................................................................ 81

Table 3-2. Effects of the addition of acetoacetate on in vitro ruminal fermentation 83

Table 3-3. Effects of the addition of B-hydroxybutyrate on in vitro ruminal
fermentation ........................................................................................ 85

Table 34 Effects of the addition of crotonic acid on in vitro ruminal fermentation 87
Table 3-5. Effects of the addition of propynoic acid on in vitro ruminal fermentation 89
Table 3-6. Effects of the addition of 3-butenoic acid on in vitro ruminal fermentation .. 91
Table 3-7. Effects of the addition of 2-butynoic acid on in vitro ruminal fermentation .. 93

Table 3-8. Effects of the addition of ethyl 2-butynoate on in vitro ruminal
fermentation ........................................................................................ 95

Table 4-1. Effects of aphidicolin on ruminal fermentation in vitro .................... 112

Table 4-2. Effects of aphidicolin and DMSO on ammonia levels and FOM .......... 114
Table 4-3. Effects of the addition of lumazine on in vitro ruminal fermentation ...... 116
Table 4-4. Effects of the addition of BPS on in vitro ruminal fermentation ............ 118
Table 4-5. Effects of the addition of BES on in vitro ruminal fermentation ............. 119

Table 5-1. Effects of a hexadecatrienoic oil and olive oil on ruminal fermentation
in vitro ............................................................................................ 130

Table 6-1 . Effects of lumazine and crotonic acid on 24 h ruminal in vitro
fermentation (Experiment 1) .................................................................. 146

Table 6-2. Effects of lumazine and 3-butenoic acid on 24 h ruminal in vitro
fermentation (Experiment 2) .................................................................. 148

Table 6—3. Effects of propynoic acid and crotonic acid on 24 h ruminal in vitro
fermentation (Experiment 3) .................................................................. 150

Table 64. Effects of propynoic acid and 3-butenoic acid on 24 h ruminal in vitro
fermentation (Experiment 4) .................................................................. 153

Table 6-5. Effects of ethyl 2-butynoate and crotonic acid on 24 h ruminal in vitro
fermentation (Experiment 5) .................................................................. 156

Table 6-6. Effects of ethyl 2-butynoate and 3-butenoic acid on 24 h ruminal in vitro
fermentation (Experiment 6) .................................................................. 158

Table 6-7. Proportion of the decrease in CH4 formation accountable by the complete
reduction of propynoic acid triple bond (Experiments 3 and 4) .......................... 164

Table 7-1. Effects of lumazine and crotonic acid on 72 h in vitro degradation and
microbial biomass production Glxperiment 1) ............................................. 184

Table 7-2. Effects of lumazine and 3-butenoic acid on 72 h in vitro degradation and
microbial biomass production (Experiment 2) ............................................. 187

Table 7-3. Effects of propynoic acid and crotonic acid on 72 h in vitro degradation and
microbial biomass production (Experiment 3) ............................................. 189

Table 74. Effects of propynoic acid and 3-butenoic acid on 72 h in vitro degradation and
microbial biomass production (Experiment 4) .............................................. 191

Table 7-5. Effects of ethyl 2-butynoate and crotonic acid on 72 h in vitro degradation and
microbial biomass production (Experiment 5) .............................................. 194

Table 7-6. Effects of ethyl 2-butynoate and 3-butenoic acid on 72 h in vitro degradation
and microbial biomass production (Experiment 6) ......................................... 197

Table 8-1. Effects of 2-bromoethanesulfonate (BES) on CH4 production (umoles) of

three ruminal methanogens ..................................................................... 216
Table 8-2. Effects of 3-bromopropanesulfonate (BPS) on CH4 production (umoles) of
three ruminal methanogens ..................................................................... 219
Table 8-3. Effects of lumazine on CH4 production (umoles) of three ruminal
methanogens ...................................................................................... 220
Table 8-4. Effects of propynoic acid on CH4 production (umoles) of three ruminal
methanogens ...................................................................................... 222
Table 8-5. Effects of ethyl 2-butynoate on CH4 production (umoles) of three ruminal
methanogens ...................................................................................... 223
Table A-1 Unprocessed data for Chapter 2, Experiment 1: optical density ............. 245
Table A-2 Unprocessed data for Chapter 2, Experiment 1: gases, fermentation, and
ammonia .......................................................................................... 248
Table A-3 Unprocessed data for Chapter 2, Experiment 1: VFA production ........... 251
Table A—4 Unprocessed data for Chapter 2, Experiment 2: gases, fermentation and
ammonia ........................................................................................... 254
Table A-5 Unprocessed data for Chapter 2, Experiment 2: VF A production ........... 255
Table A-6 Unprocessed data for Chapter 2, Experiment 3: gases, fermentation, pH

and ammonia ..................................................................................... 256
Table A-7 Unprocessed data for Chapter 2, Experiment 3: VFA production ........... 257
Table A—8 Unprocessed data for Chapter 2, Experiment 4: gases, fermentation, pH

and ammonia ...................................................................................... 258
Table A-9 Unprocessed data for Chapter 2, Experiment 4: VFA production ........... 259

Table A-lO Unprocessed data for Chapter 3, Experiment 1, Run 1: gases,
fermentation, pH and ammonia ................................................................ 260

Table A-Il Unprocessed data for Chapter 3, Experiment 1, Run 1: VFA
production ......................................................................................... 261

Table A-12 Unprocessed data for Chapter 3, Experiment 1, Run 2: gases,
fermentation, pH and ammonia ............................................................... 262

Table A-13 Unprocessed data for Chapter 3, Experiment 1, Run 2: VFA
production ....................................................................................... 263

Table A-14 Unprocessed data for Chapter 3, Experiment 2, Run 1: gases,
fermentation, pH and ammonia ................................................................ 264

Table A-15 Unprocessed data for Chapter 3, Experiment 2, Run 1: VFA
production ........................................................................................ 265

Table A-16 Unprocessed data for Chapter 3, Experiment 2, Run 2: gases,
fermentation, pH and ammonia ............................................................... 266

Table A-17 Unprocessed data for Chapter 3, Experiment 2, Run 2: VFA
production ........................................................................................ 267

Table A-18 Unprocessed data for Chapter 4, aphidicolin: gases, fermentation, pH
and ammonia ..................................................................................... 268

Table A-19 Unprocessed data for aphidicolin: VFA production ......................... 269

Table A-20 Unprocessed data for Chapter 4, lumazine, BPS and BES: gases,
fermentation, pH and ammonia ............................................................... 270

Table A-21 Unprocessed data for Chapter 4, lumazine, BPS and BES: VFA

production ....................................................................................... 272
Table A-22 Unprocessed data for Chapter 5: gases, fermentation, pH and
animonia .......................................................................................... 274
Table A-23 Unprocessed data for Chapter 5: VFA production ........................... 275
Table A—24 Unprocessed data for Chapter 6, Experiment 1: gases, pH and
atlltnonia ........................................................................................... 276
Table A-25 Unprocessed data for Chapter 6, Experiment 1: VFA production ......... 277
Table A-26 Unprocessed data for Chapter 6, Experiment 2: gases, pH and
atlilzz-Jonia ........................................................................................... 278
Table A-27 Unprocessed data for Chapter 6, Experiment 2: VFA production ......... 279

xiii

Table A-28 Unprocessed data for Chapter 6, Experiment 3: gases, pH and

ammonia ........................................................................................... 280
Table A-29 Unprocessed data for Chapter 6, Experiment 3: VFA production ......... 281
Table A-30 Unprocessed data for Chapter 6, Experiment 4: gases, pH and
ammonia ........................................................................................... 282
Table A-31 Unprocessed data for Chapter 6, Experiment 4: VFA production ......... 283
Table A-32 Unprocessed data for Chapter 6, Experiment 5: gases, pH and
ammonia ........ ‘. ................................................................................. 284
Table A-33 Unprocessed data for Chapter 6, Experiment 5: VFA production ......... 285
Table A-34 Unprocessed data for Chapter 6, Experiment 6: gases, pH and
ammonia ........................................................................................... 286
Table A-35 Unprocessed data for Chapter 6, Experiment 6: VFA production ......... 287
Table A-36 Unprocessed data for Chapter 7, Experiment 1 ............................... 288
Table A-37 Unprocessed data for Chapter 7, Experiment 2 .............................. 289
Table A-38 Unprocessed data for Chapter 7, Experiment 3 .............................. 290
Table A-39 Unprocessed data for Chapter 7, Experiment 4 .............................. 292
Table A-40 Unprocessed data for Chapter 7, Experiment 5 .............................. 294
Table A-4l Unprocessed data for Chapter 7, Experiment 6 ............................... 296
Table A-42 Unprocessed data for Chapter 8 ................................................. 298
Table A-43 Chemical structures and physico-chemical properties of the compounds
Studied ............................................................................................ 304
xiv

LIST OF FIGURES

Figure 1-1 Carbohydrate fermentation by an anaerobic ruminal phycomycete in the
absence and presence of a methanogen ........................................................ 7

Figure 2-1. Effects of the addition adenine or adenosine on optical density of ruminal

bacterial cells ...................................................................................... 58
Figure 4-1 Effects of lumazine on gases production in ruminal fermentation .......... 115
Figure 7-1 Crotonic acid disappearance (Experiment 1) ................................... 183
Figure 7-2 3-Butenoic acid disappearance (Experiment 2) ................................ 186
Figure 7-3 Crotonic acid disappearance (Experiment 5) ................................... 195
Figure 7—4 3-Butenoic acid disappearance (Experiment 6) ................................ 196

Figure 7-5 Relationship between C disappeared from additives and the increase in
microbial C synthesis ............................................................. 203

XV

KEY TO SYMBOLS AND ABREVIATIONS

ADP — adenosine diphosphate
AN OVA — analysis of variance
ATP - adenosine triphosphate
BES -— 2-bromoethanesulfonate
BPS — 3-bromopropanesulfonate
C - carbon
CH4 ‘— methane
CO - carbon monoxide
CO2 — carbon dioxide
CoA — coenzyme A
CoM or HS-CoM — coenzyme M
CP - crude protein
DE - digestible energy
DM — dry matter
DM - dry matter intake
DMso - dimethylsulfoxide
ElVIN S — efficiency of microbial nitrogen synthesis
EMOMS — efficiency of microbial organic matter synthesis
F ON — apparently fermented organic matter
g § gravity acceleration (9.8 m/sz)

GC — gas chromatograph

xvi

GE — gross energy

GEI — gross energy intake

H - metabolic hydrogen

H2 — dihydrogen

H4MPT — tetrahydromethanopterin

HPLC - high precision liquid chromatography
LDso — lethal dose 50

LOI — level of intake

MEI ’— metabolizable energy intake

MFR - methanofuran

MN - microbial nitrogen

MOM -— microbial organic matter

N —- nitrogen

N2 - dinitrogen

NAD+ - oxidized nicotine adenine dinucleotide
NADH - reduced nicotine adenine dinucleotide

ND — nitrogen degradation
NDF — neutral detergent fiber
NDFD — neutral detergent fiber degradation
NH; - ammonium ion
Ni - nickel
03 - ozone

OD - optical density

xvii

 

OM - organic matter

OMD — organic matter degradation

PMOM -— prOportion of disappeared organic matter from substrate incorporated into
microbial biomass

R2 —- coefficient of determination

RNA — ribonucleotide acid

rRNA — ribosomal ribonucleotide acid

SSU rRNA — small subunit ribosomal ribonucleotide acid

VFA— volatile fatty acids

xviii

 

INTRODUCTION

Ruminants have evolved with a microbial pregastric digestive system that allows
the utilization of feed components that are unusable to other animals, such as cellulose,
hemicellulose, and non-protein-N (Van Nevel and Demeyer, 1996; Baker, 1997). This is
very advantageous, because structural are much more abundant than non-structural
polysaccharides (Hobson, I997). Pregastric microbial digestion allows humans to
harvest the photosynthetic potential of grasslands (Russell, 2002), and is the basis for the
exploitation of domestic ruminants and camelids for milk, meat, fiber, and draft power
(Baker, 1997).

The presence of microbial fermentation between the host animal and plant
material, however, constitutes an additional trophic level, and implies inevitable
inefiiciencies in the transfer of energy and matter (Baker, 1997). Products like CO2, CH4,
and heat, have no nutritional value for the host animal, and represent a loss of carbon,
energy, or both. Methane produced in the foregut and hindgut accounts for between 2

and 15% of the animals gross energy intake, although hindgut fermentation may account
for up to 12% of overall CH4 produced (Czerkawski, 1986; Baker, 1997).
Also, CH4 production is associated with the balance between glucogenic and non-
glucogenic VFA, which can be important for ruminants. Propionate is the main glucose
Pmcursor for ruminants, and there is an inverse relationship between propionate molar

Prolrortion and the proportion of fermented energy released as CH4 (Baker, 1997; Wolin
31: al., 1997).

 

 

There is also interest in CH4 production by domestic ruminants due to the role of
CH4 as a greenhouse gas involved in global warming (Baker, 1997). Methane emissions
are responsible for between 18 and 20% of global warming, and ruminal fermentation is a
major source of atmospheric CH4 (Moss, 1993).

For these reasons, considerable research on the inhibition of ruminal
methanogenesis has been carried out. Many chemical inhibitors have been investigated,
and have proved to be effective in decreasing CH4 to varying degrees. Problems
associated with their use have been decreased animal intake, H2 accumulation, microbial
adaptation, decreased digestibility, toxicity to the host animal, volatility, and inability to
improve energetic efficiency (Moss, 1993; Van Nevel and Demeyer, 1996).

Direct inhibition of the pathway of methanogenesis poses the problem of the
relocation of the electrons not used in CH4 formation. Inefficient relocation of these
electrons results in H2 accumulation and decreased re-oxidation of cofactors, which in
turn, inhibits fermentation (Van Nevel and Demeyer, 1996). It is proposed in this thesis
that alternative strategies, targeted towards inhibiting the production of precursors of CH4

formation or towards competing with methanogenesis for electrons, may decrease CH4
Production without causing some of those problems.
The present work examines novel strategies and compounds to inhibit ruminal
IIlviathanogenesis in vitro. Ruminal in vitro techniques allow for a fast, simple, and
inexpensive preliminary examination of the effects of chemicals on CH4 production and
feI‘l:rrentation. In Chapter 2, attempts to inhibit pyruvate oxidative decarboxylation, either
directly or through the inhibition of thiamin utilization, as a potential means to inhibit

CH4 production by decreasing the supplyof precursors, are described. In Chapter 3, the

use of oxaloacetate, butyrate enhancers, and unsaturated compounds as alternative
electron sinks to methanogenesis, were studied. In Chapter 4, three compounds that are
unrelated in their hypothesized mode of action were examined: aphidicolin, lumazine,
and 3-bromopropanesulfonate. In Chapter 5, the effects of a novel hexadecatrienoic fatty

acid extracted fiom a marine algae and of olive oil on CH4 production and ruminal

fermentation were studied.
From Chapters 2 through 5, three compounds were selected based on their ability

to decrease CH4 production: lumazine, propynoic acid, and ethyl 2-butynoate. However,
the inhibitors caused the formation of fermentation products without a nutritional value
and were estimated to have adverse effects on fermentation. In contrast, two organic
acids, crotonic acid and 3-butenoic acid, had minimal effects on CH4 production but
se ened to benefit fermentation. In Chapters 6 and 7, the effects of combinations of
l Innazine, propynoic acid, and ethyl 2-butynoate, with crotonic acid and 3-butenoic acid,
on CH4 formation, fermentation, substrate degradation, and microbial biomass production
Were examined. It was hypothesized that the combination of the inhibitors of CH4
fc)Ifilrttzration with the external electron sinks can relieve the constraints on fermentation
caused by the former, and re-channel electrons into butyrate formation.
Adaptation of ruminal rnicrobiota to chemical inhibitors of CH4 production can
a1 So be a problem for their utilization in vivo (Van Nevel and Demeyer, 1996). The
Dre Sence of methanogens resistant to inhibitors could lead to long term adaptation of the

t“—:l~)::.:._inal rnicrobiota (Ungerfeld, 1998). In Chapter 8, the effects of lumazine, propynoic
3% ‘
l d, ethyl 2-butynoate, 3-bromopropanesulfonate, and the classical methanogenesis

mi bitor 2-bromoethanesulfonate (N agaraja et al., 1997), on pure cultures of the ruminal

methanogens Methanobrevibacter ruminantium, Methanosarcina mazei, and

Methanomicrobium mobile were examined.

 

REFERENCES

Baker, S. K. 1997. Gut microbiology and its consequences for the ruminant. Proc. Nutr.
Soc. Aust. 21: 6—13.

Czerkawski, J. W. 1986. An Introduction to Rurnen Studies. Pergamon Press, Oxford,
UK.

Hobson, P. N. 1997. Introduction. In: P. N. Hobson and C. S. Stewart (eds.) The Rurnen
Microbial Ecosystem. p l-9. Blackie Academic and Professional, London.

Moss, A. R. 1993. Methane. Global Warming and Production by Animals. lst ed.
Chalcombe Publications, Kingston, Kent, UK.

N agaraja, T. G., C. J. Newbold, C. J. Van Nevel, and D. I. Demeyer. 1997. Manipulation
- of ruminal fermentation. In: P. N. Hobson and C. S. Stewart (eds) The Rurnen
Microbial Ecosystem. p 523-632. Blackie Academic and Professional, London.

Russell, J. B. 2002. Rurnen Microbiology and Its Role in Ruminant Nutrition. lst ed,
Ithaca, NY.

Ungerfeld, E. M. 1998. Characterisation of resistance to 2-bromoethanesulphonate (BES)
in sheep rumen fluid. MSc thesis, Aberdeen University, Aberdeen.

Van Nevel, C. J. and D. I. Demeyer. 1996. Control of rumen methanogenesis. Environ.
Monit. Asses. 42: 73-97.

Wolin, M. J ., T. L. Miller, and C. S. Stewart. 1997. Microbe-microbe interactions. In: P.
N. Hobson and C. S. Stewart (eds.) The Rurnen Microbial Ecosystem. p 467-491.
Blackie Academic and Professional, London.

 

 

CHAPTER 1

Literature Review

Methane production in the rumen

Ruminal fermentation can be considered an anaerobic oxidation of dietary
carbohydrates, proteins, and glycerol, to acetate, CO2, and NH4+, with concomitant
production of reduced end products, mainly CH4, propionate, and butyrate (Van Nevel
and Demeyer, 1996). Many of the carbohydrate-fermenting ruminal microorganisms
produce H2, CO2, and formate as fermentation products; however, none of these
organisms produces CH4. Methanogens use the H2, CO2, and formate generated from
carbohydrate fermentation to produce CH4 (Wolin et al., 1997).

More than 90% of the glucose obtained from the degradation of carbohydrates in
the rumen is metabolized to pyruvate through glycolysis. As glycolysis releases reducing
equivalents, it is essential that pyruvate metabolism provides an electron sink for the
r e0).:idation of the reduced cofactors, so fermentation can continue (Russell and Martin,
1 9 84). Methane formation occurs from pyruvate breakdown products CO2, formate, and

H2 , and provides a route for the disposal of metabolic hydrogen in the absence of oxygen
(S tewart et al., 1997). Methanogenesis maintains a low partial pressure of H2, so that
rec Xidation of cofactors by hydrogenase is more favorable than by alcohol or lactate

<1 ehydrogenase (Van Nevel and Demeyer, 1996). Then, when methanogens are present,
B4 is the main electron sink. Pyruvate can then be metabolized to acetate, instead of

e
M01 or lactate, and the ATP yield of the hydrogen producer increases (Figure 1-1;

IL 1 :
Ssell and Wallace, 1997; Wolin et al., 1997):

‘ - "

 

 

 

HEXOSE

 

 

ACEWE

F i gme 1-1 Carbohydrate fermentation by an anaerobic ruminal phycomycete in the

absence and presence of a methanogen (Wolin et al., 1997).

In monoculture, lactate and ethanol are the electron sinks for NADH reoxidation.
In co-culture with a methanogen, NADH is oxidised to NADI by producing H2, which is
in tum used to reduce C02 to CH4 (Wolin et al., 1997).
Most ruminal CH4 is produced from H2 and CO2, although formate can also be

118 e<1 as a substrate (Russell and Wallace, 1997). Most formate, however, is converted to

Q2
hi

alld C02 prior to methanogenesis (Hungate etal., 1970). Rurnen outflow rates are too
$11 to allow significant methanogenesis from VFA, as methanogens using these

SR 1
b Strates grow slowly (Russell and Wallace, 1997).

 

ml

I!

 

Although the most important, CH4 is not the only electron sink in the rumen.

Propionate (Russell and Wallace, 1997), butyrate (Miller and Jenesel, 1979), and lactate

(Moss, 1993) formations imply the uptake of reducing equivalents. Other electron sinks
include sulfite, nitrate, and nitrite (N agaraja et al., 1997), and fatty acids

biohydrogenation and synthesis (Czerkawski, 1986).
Methanogens belong to the domain Archaea, and share with prokaryotes features

such as similar size, absence of organelles, and size of ribosomal subunits. Features in
common with eukaryotes include cell wall structure, insensitivity to vancomycin,

penincillin and kanamycin, absence of formyl-methionine in protein synthesis, and ADP-
ri bosylation of the peptide synthesis elongation factor EF2 by diphteria toxin. RNA
translation and ribosomal shape are distinctive from both prokaryotes and eukaryotes
(Moss, 1993; Wolin et al., 1997). All methanogens are fastidious anaerobes, have
relatively simple nutritional requirements, and, more importantly, use methanogenesis as
the only free energy source for ATP synthesis. Apart fiom this common physiological
features, there is considerable phylogenetic diversity among methanogens, reflected in
the macromolecules responsible for the sacculus rigidity, membrane lipid composition,
atld rRNA sequences. There is variation among methanogens in types and relative
Ei-tflilounts of cofactors such as coenzyme F420 analogues, vitamin Bl2-like corrinoids,

pterins, and methanofurans (White, 1988; Stewart et al., 1997; Wolin etal., 1997).
For their anabolism, methanogens take advantage of the first part of the C02

1-
% ductive pathway to de novo synthesize acetate from two molecules of CO2 and four H2.
Qetyl-COA is the central metabolite for synthetic reactions. Although the production of
1%
Q‘ltate from CO2 and H2 is thermodynamically favorable in reductive acetogenesis, this

 

; __

 

is only at higher partial pressures of H2 than those found in methanogenic habitats.
Furthermore, the formation of acetyl-CoA implies an additional energy cost. However,
there is no ATP requirement for acetyl-CoA biosynthesis in methanogens, and the free
energy is probably derived from a proton or sodium motive force. Pyruvate can be
synthesized from acetyl-CoA, and then converted to glucose by gluconeogenesis.
Hexoses are required as building blocks for cell wall components, although glycogen has
been found in some methanogens (Blaut, 1994).
Although methanogens as a group are able to use a variety of compounds as N
sources, individual species are relatively restricted in their choices. All methanogens can
use NH4+, many will fix N2 if deprived from NIH", some can deaminate amino acids,
some hydrolize urea, others metabolize methylamines, and some degrade purines or
pyrimidines (DeMoll, 1993).
Methanogens are the only archaea that have been found in the rumen. Archaeal
Small subunit ribosomal RNA (SSU rRNA) comprises between 0.6 and 2.4% of total
l"l-Tl-Ininal SSU rRNA (Lin et al., 1997). Compared to other ecosystems, there have been
3 ul‘prisingly few studies on the isolation and characterization of rumen methanogens.
This is partially explained by the difficulties with isolation or culture maintenance
C‘IIIa-‘l'vis et al., 2000). Methanogens that have been isolated from the rumen to date are

1 i Sted in Table 1-1.

 

 

 

 

 

 

 

 

 

 

 

 

 

 

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Species in Table 1-1 are not exclusive to the rumen, and have been isolated from
other habitats as well. Methanobacteriumformicicum, Methanosarcina barkeri, and M

mazei have been found in ditch muds and sewage plants (Stewart et al., 1997).

Methanogens cell envelopes are different from bacterial. The pseudomurein layer
of Methanobrevibacter and Methanobacterium species does not contain muramic acid,
di-aminopimelic acid, or teichoic/teichuronic acids, as does bacterial peptidoglycan
(Stewart et al., 1997). The methanogens pseudomurein contains L- instead of D-amino

acids, and N-acetyl-L-talosaminuronic acid instead of N-acetylmuramic acid. Also, the

linkage configuration in the glycan strands is B(1,3) rather than B(1,4) as in murein

(Sprott and Beveridge, 1993).
Methanogens that do not posses pseudomurein have S-layer proteins, both

glycosylated and non-glycosylated. Surrounding the S-layer, Methanosarcina mazei has

a methanochondroitin layer, composed of a non-sulfated polymer of N-

acetylgalactosamine, D-glucuronic (or D-galacturonic) acid, some D-glucose, and traces

Of D-mannose (Sprott and Beveridge, 1993).
The family Methanobacteriaceae, represented in the rumen by

Methanobrevibacter ruminantium, constituted most of the methanogenic population in a

Cow rumen (Sharp et al., 1998), and are probably the dominant H2- and C02-using

W methanogens (W olin et al., 1997; Baker et al., 1998). Coccobacilli that
morphologically resembled Methanobrevibacter spp. and bound antibodies from

Me 1‘ hanobrevibacter smithii PS and Methanobrevibacter arboriphilicus DHl and DC

were isolated from the rumen of a sheep fed silage (Baker et al., 1998).
:1 ethanomicrobium mobile is the only species of the order Methanomicrobiales so far

11

found in the rumen. Although it has been isolated on few occasions (Stewart et al.,
1997), it is regarded as an important species due to its high nrunbers (Paynter and
Hungate, 1968; Baker et al., 1998), and it was the dominant species in a sheep rumen
(Y anagita et al., 2000). The order Methanosarcinales includes in the rumen
Methanosarcina borkeri and Methanosarcina mazei. High numbers of Methanosarcina
have been found in sheep fed a diet rich in molasses (Rowe et al., 1979), although this
has not always been confirmed (V icini et al., 1987). SSU rRNA from unidentified
methanogens of the order Methanococcales has been found in small amounts in the
rumen of steers, cows, goats, and sheep (Lin et al., 1997). Other ruminal isolates have
not been unequivocally classified (Miller et al., 1986; Stewart et al., 1997; Tokura et al.,
1 999; Tajima et al., 2001; Takjima et al., 2001).
Some ruminal methanogens have been found to be ecto- (V ogels et al., 1980; Stumm
et al., 1982) and endosymbiotically (Finlay et al., 1994) associated with ruminal
protozoa. Between 10 and 20% of ruminal methanogens may be ectosymbiotically
associated with protozoa (Stumm et al., 1982). Endosymbionts might occupy between 1
and 2% of the protozoon volume (Finlay et al., 1994). Likely, the presence of
nilethanogenic endosymbionts in anaerobic protists is a consequence of the presence of
113"dr-ogenosomes. These are organelles engaged in cellular energy metabolism that
g<?J:uerate H2 and acetate from pyruvate or malate (Hackstein and Vogels, 1997). Close
physical proximity, but not direct contact, between hydrogenosomes and methanogenic
e1:)e<31<)symbionts, was found in ruminal protozoa (Wedam et al., 1999). Methanogenic
é‘1Q‘i‘1ity of protozoa-enriched fractions of ruminal fluid has been shown to be higher than

111 Strained ruminal fluid (Krumholz et al., 1983). Methane production is generally lower

in defaunated ruminal fluid (U shida et al., 1997), and in defaunated animals (Whitelaw et
al., 1984). Protozoal contribution to total CH4 of ruminal fluid was estimated to be
between 9 and 25% (Newbold et al., 1995), or even 37% in some estimations (Finlay et
al., 1994).

Partial sequences of SSU rDNA close to Methanobrevibacter smithii were dominant
among sequences retrieved from methanogens associated with ruminal ciliates. M
ruminantium, however, was absent (Tokura et al., 1999).

Ciliate protozoa affect methanogenesis in a genus-dependent manner. Calves that
were're-faunated with different protozoa] genus increased their CH4 production, except
for the Epidinium-monofaunation, which agrees with the low rate of attachment of
methanogens to Epidinium sp. (Itabashi, 2001). Polyplastron multivesiculatum appears

to be only weakly methanogenic, while Isotricha sp. could be the predominant CH4-

generating ciliate (U shida et al., 1997).

Methane production as an energy loss

Despite its profound implication for ruminal fermentation, CH4 formation is a loss
of energy for the host animal, as it is removed by eructation (Trei et al., 1972). Available
published data on CH4 losses by ruminants ranges from 2 to 12% of gross energy intake
(GED. Both extremes occurred in diets with 90% or more concentrates, which causes
difficulties with empirical prediction. Other than high grain diets, the majority of
praCtical ruminant diets results in methane losses of6 s 0.5% of GEI (Johnson et al.,
1 993). High grain diets are in general associated with higher amounts of CH4 as a result

13

of being more digestible, although the amount of CH4 produced per unit of OM digested
is lower than with roughages.

Among dairy (lactating, pregnant, and dry Holstein cows) and beef cattle
(pregnant and dry cows, and steers), sheep, and goats, lactating cows had the highest DM
intake (DMI), and the lowest CH4 production per unit of DMI. Although total CH4
produced increased with increasing DMI, CH4 produced per unit of DMI decreased. A
quadratic response of CH4 to DMI was established (Kurihara et al., 1997).

Published values of CH4 production as a percentage of GEI were used to built
multivariate regression models to predict CH4 energy losses based on level of intake
(LOI) and the percentage of digestible energy (DB) in gross energy (GE). Increasing LOI
resulted in decreasing CH4 losses as a percentage of GEI, which was attributed to an
increased rate of passage, resulting in decreased ruminal fermentation (Jarosz and
Johnson, 1999). As the maintenance subsidy is decreased, the proportion of CH4 emitted
as a consequence of covering maintenance requirements (“non-productive methane

losses”) decreases as a proportion of total CH4 losses (Johnson et al., 1993; Kurihara et
a1 - , 1997). The relationship of CH4 losses to DB in GE was much more variable than its
relationship to LOI. Digestible energy in GE did little to improve the predictability of
CH4 losses when the diet contained less than 80% roughage. The R2 values for high
forage diets were three times greater than for high grain diets (Hill et al., 1992; Johnson
et a1 ., 1993).
Fine grinding and/or pelleting of forage diets can reduce CH4 losses by 20 to

00/0. Also, as the ether extract content increases, CH4 production decreases (Jarosz and

J
thson, 1999).

14

Energy lost as CH4 depends on: 1) the fraction of the dietary carbohydrate that is
fermented; 2) the VFA profile of the fermentation, as the propionate pathway is an
alternative H sink to methanogenesis; and 3) the amount of C and H captured by

microbial growth (Johnson et al., 1991; Wolin et al., 1997).

Methane emissions by ruminants and global warming
The solar energy reaching the earth’s surface warms it and is radiated back in the
infrared region of the spectrum. Approximately 30% of this radiation is absorbed in the
tropOSphere by greenhouse gases: CO2, CH4, nitrous oxide, and chlorofluorocarbons.
Numerous measurements have shown a recent increase of CH4 concentrations in the
atmosphere. This is well correlated with the increase in human world population,
indicating an anthropogenic origin (Moss, 1993).
At present, CH4 emissions account for about 18% of total global warming. While
C 02 has a greater effect on global warming than CH4 at the present time, the latter is
increasing at a faster rate (1.1 vs. 0.5%/year). Furthermore, as a greenhouse gas, CH4 is
50 times more potent than CO2 on a mass basis (Moss, 1993; Young, 2001).
Being a reduced gas in an oxidizing atmosphere, CH4 must be constantly emitted
by some source to be present at a steady state concentration. The chemical reactions that
omiidize the atmospheric CH4 afiect the chemical state of the atmosphere through the
products of reactions, and the consumption of reactant species. Atmospheric CH4 exerts
an influence on the earth climate both directly and indirectly. The former role involves
ale absorption of infrared radiation, warming the earth surface, and the near-surface

atrrlo sphere, and cooling the stratosphere. Indirectly, CH4 is ultimately oxidized in the

15

atmosphere to C02. The atmospheric production of CO2 from CH4 can be equivalent to
about 6% of the direct annual release of CO2 from anthropogenic sources (Moss, 1993).

Also, the oxidation of CH4 in the presence of nitrogen oxides produces ozone
(03), which is a particularly effective greenhouse gas in the upper troposphere.
Additionally, the oxidation of CH4 in the stratosphere produces water vapor, which can
cause a temperature increase. Increase in the global average temperature can result in a
higher CH4 production from soil methanogens, accelerating the process (Moss, 1993).

The largest single source of CH4 (about 21%) appears to be wetlands. About 70%

of CH4 emissions would be anthropogenic (Table 1-2).

Domestic animals appear to rank second among agricultural sources of CH4, with
another 2% possibly contributed by anaerobic decomposition of manure. However, there
is uncertainty in these estimates, and enteric emissions from livestock have been ranked
as the largest anthropogenic source of CH4 (McCrabb, 2001).

There are potential sources of CH4 not listed in Table 1-2. Additional CH4 in

hYdIates could be released if melting of the permafrost of the artic tundra begins, or if
there is a warming of ocean bottoms. Potential CH4 release from these sources is very

large, and it could become of major importance (Johnson et al., 1991; Moss, 1993).

16

 

Table 1-2 Sources of methane emissionsl

 

 

 

 

 

 

 

 

 

 

 

 

Source Estimated amount Range (million ton) Approximate
(million ton) %
Natural wetlands 115 100 — 200 21
Rice paddies 110 60 — 170 20
Domestic animals 75 60 — 95 14
Biomass burning 55 50 - 100 10
Oil and gas drill 45 3O - 50 8
Termites and other 40 10 - 100 7
wild animals
Landfills 40 30 - 70 7
Coal mining 35 20 — 45 6
Animal waste 10 5 — 30 2
Oceans and lakes 10 6 - 45 2
Hydrates2 5 6 — 1003 l

 

 

 

 

 

 

lAdapted fi'om Johnson et a]. (1991); based on Cicerone and Oremland (1988).

2Methane molecules surrounded by rigid cages of water; they are prevalent under the
permafrost and beneath the sea in continental margins (Moss, 1993).

3Methane in hydrates could be released if global temperature continues to increase (Moss,
1 993 ).

Estimates of CH4 emissions by different species of mammals are shown in Table

l ‘3 - Due to their size and numbers, cattle are major contributors, accounting for about

740/0 of total domestic animal emissions. Methane production by wild ruminants is very

8111811 when compared with domestic animals (Moss, 1993).

17

Table 1-3 Estimates of global methane emissions by marnmals'

 

 

 

 

 

 

 

 

 

 

 

 

Species Population2 Typical methane Total amount Fraction
(million) production (l/d) (million ton) (%)
Cattle, developed 603 210 33.1 41
countries
Cattle, developing 688 134 24.1 30
countries
Sheep 1,150 19 6.9 9
Water buffalo 126 192 6.3 8
Wild mammals 237 variable 4.0 5
Goats 460 19 2.3 3
Horses and mules 117 74 1.7 2
Camels 17 223 1.0 l
Pigs 800 5 1.0 l
Humans 4,726 0.2 0.3 0.4
1 Total 80.7 100.0

 

 

 

 

 

 

 

fiAdapted from (Johnson et al., 1991).

2Based on 1984 estimates.

Beef cattle are responsible for about two thirds of total cattle emissions in the US.
Within beef cattle, beef cows account for about 60% of the emissions (Table 1-4;

JQhJ-ison, 1993).

18

 

 

Table 1-4 Estimates of methane production by US cattle‘

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

Population Days Energy in CH4 Total CH4 Fraction
Class (million) fed methane (L/head (million (%)
(% of GE) /d) ton/year)
Beef cows 33.7 365 6.2 262 2.3 39
Dairy cows 10.7 365 5.8 492 1.4 23
Stockers 38.1 150 6.5 202 0.8 13
Replacements 9.9 365 6.5 220 0.6 10
Feedlot 26.8 140 3.5 153 0.4 6.7
Calves 38.9 210 6.0 53 0.3 5
Bulls 2.2 365 6.0 380 0.2 3.3
Total 160.3 6.0 100

 

‘Adapted from Johnson et al. (1991).

Potential CH4 production from animal manure is large. The amount depends on

the amount of manure in lagoons, and on the activity of methane oxidizing

microorganisms on the lagoon surface. Limited observations suggests that ruminant

manure produce only 2% of its potential level (Johnson et al., 1991). However, anaerobic

lagoons are a major target for greenhouse gases mitigation, not only because of CH4, but

also because of nitrous oxide emissions (Johnson et al., 2001). Due to much slower

Qlltflow rates than the gastrointestinal tract, acetate is the main precursor for

l'lle‘lhanogenesis in anaerobic fermentation of animal effluent (Takahashi, 2001).

19

 

Biochemistry and energetics of methanogenesis

Most methanogens can grow on CO2 and H2 as sole energy sources (Thauer et al.,
1993y

CO2 + 4H2 -> CH4 + 2H2O

The reduction of CO2 to CH4 proceeds via coenzyme bound Ct-intermediates.
Methanofuran (MFR), tetrahydromethanopterin (H4MPT), and coenzyme M (HS-COM)
are the three Cl-unit carriers found in all methanogens analyzed to date. Methanogenesis
also involves several electron carriers: coenzyme F420, N-7-mercaptoheptanoyl-0-
phospho-L-threonine, ferredoxin, a polyferredoxin with 12 [4Fe-48] clusters and other
ion-sulfur proteins with unknown functions. Methanogens capable of oxidizing methyl
groups also contain cytochromes (Thauer et al., 1993).

The following coenzyme-bound Ct-intermediates in methanogenesis from carbon
dioxide and hydrogen have been identified (Thauer et al., 1993): N-formyl-MF R (CHO-
MFR), NS-formyl-H4MPT (CHO-H4MPT), N5, N'o-methenyl-H4MPT (CHEH4MPTT), N5,
Nm-methylene-H4MPT (CH2=H4MPT), NS-methyl-l-I4MPT (CH3-H4MPT), and
methylcoenzyme M (CH3-S-CoM). The partial reactions and their free energy changes

are shown in Table 1-5.

20

Table 1-5 Partial reactions of methanogenesis and fiee energy changes

co2 + MFR + H2 —2 CHO-MFR + H2O + H“; no“: 16 kJ/mol
CHO-MFR + H4MPT —> CHO-H4MPT + MFR; AGO’ = 4.4 kJ/mol
CHO-H4MPT + H* —> CHEH4MPT+ + H2O; AGO' =-4.6 kJ/mol
CHEH4MPT’ + H2 —2 CH2=H4MPT + 11*; Ao°'=-5.5 kJ/mol
CH2=H4MPT + H2 -—> CH3-H4MPT; AG°’=-17.2ltJ/tnol

CH3-H4MPT + HS-CoM —> CH3-S-CoM + H4MPT ; no” = -29.7 kJ/mol

CH3-S-CoM + H2 —9 CH4 + HS-CoM ; AG°’=-85 kJ/mol

 

These reactions account for a total free energy change of —1 30.4 kJ/mol, which
differs only by 0.6 kJ/mol from the free energy change calculated from the standard free
energies of formation from the elements (Thauer et al., 1993). There is evidence that the
last two reactions are coupled with energy conservation by transmembrane proton and
sodium ion gradients (Blaut, 1994), and that the first, endergonic reaction, is driven by
reversed electron transport (Thauer et al., 1993).

Methanogens contain several hydrogenases for the activation of H2, which is a
substrate in four of the seven partial reactions (Table 1-5). There are two (N iFe)

hydrogenases: a coenzyme F42o-reducing hydrogenase, and a coenzyme F42o-non-
reducing hydrogenase, with an unknown electron acceptor. In addition of the two (N iF e)
h.Ydrogenases, most methanogens contain a third, very active hydrogenase, H2-forming

methylenetetrahydromethanopterin dehydrogenase, which differs from other

21

L

 

hydrogenases known to date in that it does not contain nickel or ion/sulfur clusters

(Thauer et al., 1993).

Control of methane production in the rumen

Strategies to decrease CH4 production in the rumen include dietary manipulation,

chemical additives, microbial additives, and others.

Dietary manipulation

Increasing the proportion of concentrates in the diet usually decreases the
proportion of GEI lost as CH4 (Johnson et al., 1993; Van Nevel and Demeyer, 1996),
although no difi'erences were observed in goats fed at maintenance when the percentage
of hay in the diet was decreased from 90 to 30% (Kurihara et al., 1997). In lactating
dairy cows, however, CH4 energy losses were decreased from 14.3 to 10.5% of
metabolizable energy intake (MEI) when hay in the diet was decreased from 70 to 30%
(Kurihara et al., 1997).

Elevation of dietary crude protein fi'om 4 to 9% in goats fed at maintenance
resulted in an increase in CH4 produced per unit of DMI and the ratio of CH4 energy
losses to MEI (Kurihara et al., 1997).

A modified mathematical model of rumen digestion was used to simulate the

effect of different nutritional strategies on CH4 production (Benchaar et al., 2001). It was
fOurrd that diet changes would allow decreases in CH4 emission between 10 and 40%.
Increasing DM intake and the proportion of concentrate in the diet decreased CH4 losses

as a proportion of GEI by 7 and 40%, respectively. The replacement of fibrous

22

concentrate with starchy concentrate, and the utilization of less ruminally degradable
starch decreased CH4 losses by 22 and 17%, respectively. The use of more digestible
forage resulted in decreases between 15 and 21%. The replacement of legumes for
grasses reduced CH4 losses by 28%, while replacing silage for hay decreased it by 20%.
In general, decreasing methanogenesis is accompanied by a decreased acetate to
propionate ratio, in agreement with competing interspecies H transfer reactions between
the formation of CH4 and propionate (Van Nevel and Demeyer, 1996). Means of
decreasing the acetate to propionate ratio via the diet include grinding and pelleting of
roughages, heat treatment of grain, increasing feeding frequency, increasing intake of
mixed diets, and chemical treatments of straws (Van Nevel and Demeyer, 1996).
Supplementation of dry cows with 35 g of ZnSO4 per day decreased CH4
production per unit of DMI by 62%, and the ratio of CH4 energy losses to MEI by 61%,
although DM digestibility was decreased by 4 percentage units. The reduction in CH4
production was thought to be related to the repression of rumen fermentation. Although
protozoal numbers tended to decrease, methanogens were not affected (Kurihara et al.,
1 997). It is possible, though, that the most probable number determination used to
estimate methanogens numbers only accounted for free methanogens.
Biohydrogenation of fatty acids was proposed as an alternative electron sink to
methanogenesis (Czerkawski, 1986). The extent of the inhibition depends on the nature
and amount of the lipid fed (Van Nevel and Demeyer, 1996). The extent of the decrease
is greater with unsaturated fatty acids (Czerkawski, 1986), and free fatty acids are more
POtent than triacylglycerols (Van Nevel and Demeyer, 1996). However, the reduction in

methanogenesis cannot be explained, at least entirely, by biohydrogentation of fatty

23

 

acids, or by negative effects of fatty acids on protozoa (Nagaraja et al., 1997). Studies

with pure cultures revealed that ruminal methanogens are very sensitive to long chain,
unsaturated fatty acids (Prins et al., 1972; Henderson, 1973). Also, Ruminococcus albus
and Ruminococcusflavefaciens, which have a Gram positive cell wall structure and
produce CH4 precursors, are more inhibited by long chain, unsaturated fatty acids than
Gram positive bacteria important in propionate formation (Henderson, 1973; Maczulak et
al., 1981). The toxic action of long chain fatty acids is due to adsorption onto the cell
wall, which alters nuuients passage (Henderson, 1973).

‘ Methane emissions are also decreased by high amounts of lipids in the diet
because of lower ruminal digestion of OM and fiber. Although there can be some
compensation by a shift of digestion to the lower tract, the amount of CH4 formed per
mole of substrate fermented in the hindgut is much lower than in the rumen (Van Nevel
and Demeyer, 1996).

Supplementation with coconut oil decreased CH4 production in the chemostat
(Machmiiller et al., 1998) and in sheep (Machmi’rller and Kreuzer, 1998), although there
was a tendency to decrease fiber digestibility in vivo. Canola and cod liver oils decreased
CH4 production in the chemostat without detrimental effect on DM or NDF
disappearance. Methanogen numbers were decreased, but cellulolytic and amylolytic
bacteria, or endoglucanase activity were not affected, or were increased, by canola and

c0d liver oils (Dong et al., 1997). The addition of 3.5% soybean oil to dairy cow diets

did not change CH4 production, although the CO2 to CH4 ratio increased from 10.4 to

1 l -3 (Sauer et al., 1998).

24

 

 

_- -_-—--—-~

 

Supplementation of sheep with 5% myristic acid (C140) decreased CH4 production
by about 50% with a 1:1.5 hayzconcentrate diet. The extent of the decrease was lower
with a 1:0.5 hayzconcentrate diet. With this diet, an increase in dietary calcium released

the inhibition of CH4 production, presumably due to the formation of inactive soaps in

the rumen (Machmi'rller et al., 2001)

Laurie acid (C120) , but not myristic acid (C140) or stearic acid (C134,) , decreased
CH4 formation in mixed ruminal batch fermentations. Methanogen numbers were
decreased by lauric acid and myristic acid, although the latter did not affect methanogenic

activity. Possibly, methanogenic activity per cell increased due the excess of H2

available. Myristic acid in addition to lauric acid enhanced the inhibitory effect of the

latter on both methanogenesis and methanogen numbers, showing synergism with respect

to CH4 formation (Soliva et al., 2001).

Chemical additives
Compounds that inhibit ruminal methanogenesis have three different modes of

action: 1) inhibition of microorganisms that produce the precursors for CH4 production
(H2, CO2, and/or formate); 2) direct inhibition of methanogens; and 3) alternative electron
acceptors that compete with methanogenesis.
In the first category are ionophores (N agaraja et al., 1997) and defaunating
(Itabashi, 2001; Takahashi, 2001) agents, which inhibit bacteria with a Gram-positive cell
Wall structure, and protozoa, respectively. An indirect consequence is a reduction in CH4

DrOduction (McSweeney and McCrabb, 2001), but they are not specific inhibitors of

“1111mm methanogenesis.

25

¥— __

 

The extent of methanogenesis inhibition caused by ionophores depends on the
dose administered, and substrate incubated or ration fed (Van Nevel and Demeyer, 1996).
Monensin has been shown to decrease CH4 production in vitro (Stanier and Davies, 1981;
Sauer and Teather, 1987) and in vivo (Sauer et al., 1998). The decrease is mainly
mediated by the inhibition that monensin exherts on H2 -producing bacteria with a Gram-
positive cell wall structure, although there might be some direct effects of monensin on
methanogens related to an inhibition of Ni uptake. Conversely, gram negatives are
protected by their outer membrane. The inhibition of H2 -producing bacteria depletes the
precursors for CH4 formation. There is a shift in the microbial population towards less
sensitive species that produce more propionate (Van Nevel and Demeyer, 1996; Nagaraja
et al., 1997). With this approach, H2 does not accumulate, and propionate increases at the
expense of acetate and butyrate (Garcia-Lopez et al., 1996). These changes decrease the
energy lost in CH4, improving the energy retained in VFA (N agaraja et al., 1997).
However, the inhibition of methanogenesis caused by ionophores in vivo is rather modest
(between 10 and 30%). Another problem is that partial adaptation of ruminal rnicrobiota
for the decrease in methanogenesis has been found after two weeks of feeding monensin.
On the contrary, changes in VF A molar proportions were maintained at the end of long
term trials with cattle and sheep. This uncoupling of the long term effect of monensin on
methanogenesis and the VFA pattern is not in agreement with the stoichiometry of
nuninal fermentation (Johnson et al., 1994; Van Nevel and Demeyer, 1996).

Defaunation reduces CH4 production in vivo between 20 and 50% (Van Nevel
and Demeyer, 1996). Although it increases total bacterial numbers, it decreases

methanogens (Itabashi, 2001). Defaunation can decrease methanogenesis by acting at

26

 

 

 

 

different levels: less fiber digestion, loss of methanogens attached to protozoa, and loss of
protozoa themselves, as they are strong producers of H2 and formate. At present, no
satisfactory defaunation method is available to apply on a practical scale (Van Nevel and
Demeyer, 1996).

Because of its selective inhibition of bacterial hydrogenases, CO decreased the
availability of H2, and therefore decreased CH4 production by 89% (Russell and Martin,
1984). Acetate to propionate ratio was decreased. However, the fact that the inhibition
was not alleviated by the addition of H2, suggests that CO could have also affected
methanogens directly.

A second strategy is to use chemicals that are directly toxic to methanogens.
Halogenated CH4 analogues such as chloroform, or bromochloromethane, and related
compounds such as chloral hydrate or amichloral (a hemiacetyl of chloral and starch), are
potent inhibitors of methanogenesis. Chloral hydrate is converted to chloroform in the
rumen, and the latter inhibits methanogenesis by blocking the methyl transfer from B12 to
coenzyme M. This prevents the formation of methylcoenzyme M, which is necessary for
the last step of methanogenesis. Likely, carbon tetrachloride (CCl4) and
bromochloromethane inactivate the same enzyme complex (Garcia-Lepez et al., 1996).
Chloroform and chloral hydrate can have acute toxic effects on the animal. When
bromomethane and amichloral were fed to animals, intake was reduced between 0 and
13%, but feed conversion was improved between 0 and 11% in seven out of eight studies
(McSweeney and McCrabb, 2001). Because of intake reductions, weight gains decreased
between 0 and 5%, although one study found a 10% improvement. A recent study with

cyclodextrin—coated bromochloromethane, a process that makes it less volatile, showed

27

 

some reduction in feed intake. However, as weight gain was not affected, conversion
efficiency was improved by 4 and 11% in low and medium quality diets, respectively
(McCrabb et al., 1997).

Pyromellitic diimide and some of its derivatives were shown to be potent
inhibitors of CH4 production in mixed ruminal cultures in vitro (Linn et al., 1982).
Pyromellitic diimide caused a 97% decrease in CH4 production in mixed batch cultures,
with a 30-fold increase in H2 accumulation (Martin and Macy, 1985). The acetate to
propionate ratio was decreased when the substrate was hay or soluble carbohydrates, but
not with a mixture of hay and concentrate. Even though diimide was a potent inhibitor of
methanogenesis in vitro, it did not decrease the number of methanogens in vivo. As the
increase in H2 accumulation would indicate that methanogenesis was directly inhibited, it
is possible that CH4 production per methanogen cell was decreased without affecting the
total number of methanogens. Methane formation was found when diimide was added to
the rumen fluid of animals being fed the chemical, suggesting adaptation of the mixed
ruminal rnicrobiota (Martin and Macy, 1985). Partial adaptation to inhibitors of
methanogenesis has also been shown for trichloroacetarnide (Clapperton, 1974) and
trichloroethyl adipate (Clapperton, 1977). This compound caused a transient inhibition
of methanogenesis in vivo, but weight gain and feed conversion efficiency of lambs were
worsened by feeding the chemical (Clapperton, 1977).

2-Trichloromethyl-4-dichloromethylene benzo[1,3] dioxin-6-carboxylic acid was
shown to decrease CH4 production by 91% in continuous culture. Hydrogen production
increased progressively. Net production of acetate, propionate and butyrate was

increased by 15, 119, and 6%, respectively. The efficiency of microbial protein synthesis

28

 

 

was decreased by the inhibitor. There was no adaptation of microbial activity to the
inhibitor (Stanier and Davies, 1981).

Several other compounds containing trichloromethyl groups were screened for
their ability to inhibit CH4 production in ruminal mixed cultures in vitro (Davies et al.,
1982). Some 6-substituted derivatives of 2,4-bis(trichloromethyl)-benzo[1 ,3]dioxin
inhibited methanogenesis when present at low concentrations. Basic substitutions had
good inhibitory activity, but large lipOphilic groups reduced it. The authors found
sustained effects in vivo with the carboxylic acid and carboxamide derivatives. 2,4-
Bis(tr'ichloromethyl)-benzo[1,3]dioxin-6-carboxylic acid was efi‘ective in reducing CH4
production in sheep when administered intrarurninally over a 5-week period (Davies et
al., 1982). In cattle, CH4 production was decreased throughout the 23 days of the study.
There was a trend for lower intakes and weight gains of cattle fed 2,4-
bis(trichloromethyl)-benzo[1,3]dioxin-6-carboxylic acid in the first half of a 28-week
performance trial. In the last 14 weeks, there was an improvement in the weight gain and
feed efliciency compared to the control at the highest dose of the chemical. The
conversion of dietary energy into energy retained in the animal was improved (Davies et
al., 1982).

Although CH4 analogues and other halogenated compounds can be severe
inhibitors of CH4 production, there are several difficulties for their use in animal
production: H2 accumulates, still representing an energy loss, digestibility and microbial
growth can be impaired, feed intake can be depressed, the inhibition caused by some
compounds can be transient, and some inhibitors can have toxic effects on the animal

(Van Nevel and Demeyer, 1996).

29

 

Forages contain phenolic monomers such as p-coumaric acid and ferulic acid. p-
Coumaric acid, phenolic acid, and cinarnmic acid, but not their hydrogenated analogs,
decreased all the measured end products, including CH4, in ruminal fermentation in vitro
(U shida et al., 1989). These compounds also decreased digestibility (Martin, 1988);
therefore, they do not appear to be specific inhibitors of CH4 production. Rather, they
likely decrease metanogenesis through a reduction in the availability of precursors.

A variety of organotin compounds has been shown to be toxic to several
methanogens, including a non-ruminal strain of Methanosarcina barkeri. The
mechanism of toxicity is unknown (Boopathy and Daniels, 1991).

Analogues of some unique cofactors involved in methanogenesis have been
assessed for their inhibitory activity on various enzymes of methanogenesis. Methyl-S-
coenzyrne M reductase catalizes the last two-electron reductive step of the overall eight-
electron reduction of carbon dioxide to methane (Wackett et al., 1987 ):

CHg-S-CoM + 2 e' + 2 I-I+ —+ CH4 + HS-CoM

2-Bromoethanesulfonate is a structural analog of coenzyme M, and inhibits the
reductive demethylation of methyl-S-coenzyme M (Miiller et al., 1993). It is a very
specific inhibitor of methanogenesis, and non-toxic to almost all other microorganisms
(Sparling and Daniels, 1987). Unfortunately, the inhibition has been transient in vivo
(Nagaraja et al., 1997).

The H donor in the reduction of methyl-S-coenzyme M to methane is N—(7-
mercaptoheptanoyl)threonineO-3-phosphate (HS-HTP), which forms a heterodisulfide
with coenzyme M afier releasing one electron (Sauer, 1991):

CH3-S-CoM + HS-HTP —> CH4 + CoM-S-S-HTP

30

 

 

The heterodisulfide is subsequently reduced by one pair of electrons and recycled.
HS-HTP is bound to a UDP-disaccharide through a carboxylic-phosphoric
anhydride linkage. There is a UDP-N-acetylglucosamine (UDP-GlcNAc) binding site in
methyl-S—coenzyme M reductase. It was found that a periodate cleaved derivative of
UDP-GlcNAc inhibits the formation of the heterodisulfide in a reaction with purified
components (Sauer, 1991). To date, the effects of UDP-GlcNAc derivatives have not

been studied in liVe methanogens.

Although cyanocobalamin is required for the activation of methyl-S-coenzyme M
reductase, high concentrations were found to be inhibitory. The inhibition by
cyanocobalamin and other corrins appeared to be a direct effect on the ATP-dependent
activation of the methylreductase. The reduction of CO2 to formylmethanofuran was also
inhibited (Whitman and Wolfe, 1987), as this endergonic reaction is energetically
coupled to the reduction of the CoM-S-S-HTP heterodisulfide.

9, IO-Anthraquinone at 5.0 ppm decreased CH4 production by 78, 95, and 83% in
in vitro ruminal batch cultures with hay, a mixed, and a high-concentrate substrate,
respectively. There was an increase in H2 accumulation. Acetate molar percentage was
decreased and propionate increased. Total VFA concentrations were decreased with hay,
but not with the mixed or high-concentrate substrates. Results with continuous culture
showed no adaptation of the ruminal rnicrobiota to 9, 10-anthraquinone. However,
prolonged feeding in vitro unexpectedly lowered propionate molar percentage. It is
reasonable to think that anthraquinone uncouples the electron transfer from cytochrome-
linked or membrane-bound ATP synthesis, thus preventing the reduction of methyl

coenzyme M to CH4 (Garcia-Lepez et al., 1996).

31

 

In the formation of methylcoenzyme M, a corrinoid prostetic group, typically 5-
hydroxy-benzimidazolyl-cobamide, transfers a methyl group to coenzyme M.
Iodopropane is a corrinoid inhibitor, and it inhibited CH4 production in pure cultures of
Methanobacterium thermoautotrophicum, Methanobacterium formicicum, and
Methanosarcina barkeri. 2-Iodopropane coated with or-cyclodextrin at 0.2 or 0.4 mM
initial concentration inhibited methanogenesis in in vitro ruminal batch cultures by 48%
and 97%, respectively, increasing H2 accumulation (Moharnrned et al., 2001).

p—Aminobenzoate (pABA) is a natural substrate for 4-(B-D-ribofuranosyl)
aminbbenzene 5’-phosphate syntlretase. Three analogs of pABA inhibited CH4 synthesis
in mixed ruminal cultures (DeMontigny et al., 2002).

A consequence of the direct inhibition of ruminal methanogenesis is an increase
in H2 accumulation. This, in turn, can interfere with the interspecies H transfer,
inhibiting the reoxidation of cofactors (N agaraja et al., 1997), and causing the
accumulation of unusual fermentation end products, like ethanol (McCrabb et al., 1997;
Wolin et al., 1997). These fermentation pathways are associated with a reduced
efficiency of microbial growth (McSweeney and McCrabb, 2001). The adaptive changes
of the ruminal microbial community to the inhibition of methanogens are relatively
unknown, although a shift of the VFA pattern towards propionate is a consistent
response. This is a consequence of the disruption of the interspecies H transfer, and the
relocation of part of the reducing equivalents spared from methanogenesis into
propionate formation (N agaraja et al., 1997; McSweeney and McCrabb, 2001).

However, the effects of methanogenesis inhibition and the resulting elevated H2 partial

32

pressures on cellulolytic numbers have not been studied (McSweeney and McCrabb,
2001)

Reducing reactions can withdraw reducing equivalents from methanogenesis.
Alternative electron sinks as nitrate or sulfate have higher reducing potentials than CO2
(Itabashi, 2001). Ruminal reduction of nitrate present in plants decreases CH4 production
in the rumen, although this benefit is counterbalanced by the formation of nitrite, which
can become toxic if it accumulates (Takahashi, 2001).

Compounds in the fermentation pathways that leads to propionate, and other
organic acids, have been used as alternative electron sinks to methanogenesis. Aspartate,
fumarate, and malate, each at 0, 4, 8, and 12 mM initial concentration, did not inhibit CH4
production in 24 h in vitro batch cultures (Callaway and Martin, 1996). Malate did not
decrease CH4 production in vitro in the absence of added substrates and with cracked
corn, but it inhibited methanogenesis by 28% with soluble starch as a substrate (Martin
and Streeter, 1995). Dihydrogen did not accumulate in these experiments. Methane
production in vitro was decreased by pyruvate, acry late, fumarate, and a—ketoglutarate by
8, 14, 8, and 13%, respectively (Lopez et al., 1999). The increase observed in propionate
production with added fumarate and acrylate stoichiometrically agreed with the decrease
in CH4 production.

In continuous culture, 33 and 44% of added acrylate and firmarate, respectively,
were recovered in propionate (Newbold et al., 2001). Methane formation was decreased
by 14 and 28%, respectively. Fumaric acid was added to sheep diets at 0, 20, 40, and 80
g/kg DM (Newbold et al., 2001). Intake was stimulated and digestibility was not

affected. Methane production was decreased by 3, 4, and 12% at 20, 40, and 80 g of

33

fumaric acid/kg DM, respectively. The addition of fumaric acid resulted in lower molar
proportions of acetate and butyrate, and higher propionate.

Furnarate metabolism of several ruminal species was studied with pure cultures
(Asanuma et al., 1999). F ibrobacter succinogenes, Selenomonas ruminantium,
Veillonella parvula, Selenomonas lactilytica, and Wollinella succinogenes utilized most
of the fumarate added. to the medium. There was a corresponding increase in succinate
and/or propionate production, and a slight increase in acetate and butyrate. Except for
Selenomonas spp., utilization was similar with H2 or formate as electron donors,
indicating the presence of formate dehydrogenases. Other ruminal bacteria utilized
smaller amounts of fumarate.

The apparent Km of methanogens for H2 was lower than for the fumarate-utilizing
bacteria (Asanuma et al., 1999). However, methanogens had a higher Km when formate
was the electron donor. Coculture of methanogens with fumarate-utilizers showed that
the addition of fumarate decreased methanogenesis, especially when formate was the
electron donor. Among the firmarate-utilizers, W. succinogenes was the most effective in
decreasing methanogenesis, which agrees with the fact that it had the lowest Km among
fumarate-utilizers for both H2 and formate.

Malate at 10 or 20 mM initial concentration decreased methanogenesis in ruminal
batch cultures by 15 and 20%, respectively (Mohammed et al., 2001). There was a
decrease in the acetate to propionate ratio. The addition of malate to ruminal cultures

where methane production was inhibited by 2-iodopropane coated with or-cyclodextrin

decreased CH4 production further, and also decreased H2 accumulation (Mohammed et

al., 2001).

34

Failure of an inhibitor of ruminal methanogenesis to improve productivity may
result fi'om a number of causes: diversion of metabolic H into products unusable by the
host animal, adverse effects of the compound on diet palatability, toxicity to ruminal
microorganisms or the host animal, the length of time that an effective concentration of
the compound is sustained in the rumen may be too short, or microbial populations may
adapt to the compound (Van Nevel and Demeyer, 1996; Nagaraja et al., 1997; Baker,

1 999).

Microbial additives

Addition of chemical additives is not the only means to rechannel the substrates
for CH4 production into alternative products. Acetogenic bacteria, which are found in the
hindgut of mammals and termites, produce acetate from the reduction of CO2 with H2
(Nagaraja et al., 1997):

2 CO2 + 4 H2 -> CH3COOH + 2 H2O

Reductive acetogenesis is an important H sink in hindgut fermentation. Reductive
acetogens were also the main H utilizers in newborn lambs, but seemed to be
outcompeted by methanogens thereafter (Morvan et al., 1994). Reductive acetogenesis
has been suggested as a possible alternative electron sink to ruminal methanogenesis
(Mackie and Bryant, 1994; Garcia-Lopez et al., 1996). However, methanogenesis
predominates over reductive acetogenesis as an electron sink in the rumen. As
methanogenesis is thermodynamically more favorable (Kohn and Boston, 2000), ruminal
methanogens have lower thresholds for utilizing H2. Also, reductive acetogens are not

obligative hydrogenotrophs and can use other compounds as energy substrates. In the

35

 

 

 

hindgut, however, acetogenesis can be an important electron sink. The reasons for these
ecological differences between compartments are unknown (McSweeney and McCrabb,
2001)

The inhibition of CH4 production, with its resultant increase in H2 partial pressure,
could make reductive acetogenesis more thermodynamically favorable, and eliminate
methanogens competitive advantage due to their lower H2 thresholds. The addition of the
reductive acetogen Peptostreptococcus productus greatly decreased H2 partial pressure
when methanogenesis was inhibited by 2-bromoethanesulfonate. The addition of the
acetogen also resulted in an increase in acetate production (N ollet et al., 1997).

Other microbial additives have been studied regarding their effects on ruminal
methanogenesis. Addition to mixed ruminal cultures of the lactic producers Leuconostoc
mesenteroides subsp. Mesenteroides, Leuconostoc lactis, or Lactococcus lactis subsp.

lactis, or the yeasts Trichosporon sericeum, or Candida kefyr, were shown to decrease

CH4 production (Gamo et al., 2001).

Others

Genetic selection to improve feed conversion efficiency could decrease CH4
emissions (Hegarty, 2001), if the level of production was kept constant.

Circulating antibodies against several ruminal microorganisms, including
methanogens, were found in Australian sheep The use of a vaccine against ruminal
methanogens is currently being investigated as a possible strategy for reducing CH4

emissions. (McSweeney and McCrabb, 2001).

36

A brush that mechanically stimulates ruminal motility decreased CH4 production
between 63 and 71%. Apparently, the physical stimulation increased the rate of passage,
which decreased CH4 production. Methane production was not affected by the size of the

stimulating brush (Matsuyarna et al., 2001).

Conclusions

Although CH4 production in the men has been inhibited by several additives,
there have been shortcomings such as transient effects, toxicity, decreased intake and/or
digestion. Ruminal fermentation is an interactive network of chemical reactions, and the
inhibition of methanogenesis should not be considered as an isolated, specific
intervention. Rather, it will have consequences on general microbial activities in the
rumen and animal metabolism. An ideal inhibitor must be extremely specific with a
persistent action, harmless to the animal and the environment, and without residues in
edible products (Van Nevel and Demeyer, 1996). Ultimately, the management of H

dynamics in the rumen is the most important factor to be considered when developing

strategies to control ruminant CH4 emissions (Joblin, 1999).

 

 

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45

CHAPTER 2
Attempts to decrease ruminal methanogenesis through the inhibition of pyruvate

oxidative decarboxylation

Abstract

The inhibition of pyruvate oxidative decarboxylation was studied as a means of
decreasing ruminal methanogenesis in vitro. In Experiment 1, the addition of thiamin (10
mM), amprolium (10 mM), adenine (10 mM), or adenosine (10 mM) decreased
methanogenesis by 22, 42, 47, and 76%, respectively. However, microbial growth was
almost non-existent, and the ratio of CO2 to CH, in the control was unusually high.
Organic matter fermentation was low in all treatments. Most likely, using an isolated cell
pellet instead of ruminal fluid, biased the conditions to make them not typical of ruminal
fermentation. In Experiment 2, the addition of adenosine (10 mM) and adenine (10 mM),
with and without ribose (10 mM), to mixed batch cultures including ruminal fluid did not
decrease methanogenesis. In Experiment 3, the addition of oxythiarnin (5 mM) decreased
methanogenesis by 23%. In Experiment 4, three pyruvate derivatives (2 mM) did not
inhibit methanogenesis, although hydroxypyruvate improved OM fermentation by 11%.
The strategies employed did not seem to inhibit pyruvate oxidative decarboxylation, and
when methanogenesis decreased in Experiment 1, this seemed to be due to the technique

used rather than to the treatments imposed.

46

Introduction

Methane production by ruminants is a carbon and energy loss, and contributes to
global warming. The inhibition of CH4 production in the rumen would have significant
economic and environmental benefits (Moss, 1993).

Pyruvate oxidative decarboxylation is the first step in the conversion of pyruvate
to acetate and butyrate in the rumen (Russell and Wallace, 1997). This reaction produces
acetyl-CoA, C02, and reducing equivalents, when catalyzed by pyruvate oxidoreductases
(U yeda and Rabinowitz, 1971), and in acetyl-CoA and formate, when catalyzed by
formate lyases (Gottschalk and Andressen, 1979). Reducing equivalents generated by
pyruvate oxidative decarboxylation and by glycolysis can be used by methanogens to
reduce CO2 and formate to CH4 (Moss, 1993).

Thiamin pyrophosphate is a cofactor of pyruvate oxidoreductases (Williams et al.,
1990). Thiamin is synthesized in the rumen (McDonald et al., 1995) and is required by or
stimulative for some ruminal microorganisms (Wolin et al., 1997). The inhibition of
thiamin utilization by ruminal microorganisms could block pyruvate oxidative
decarboxylation, diverting pyruvate to propionate formation, decreasing the availability
of CO2 and reducing equivalents for methanogenesis.

Thiamin structural analogs can inhibit bacterial growth (Koser, 1968) and impair
thiamin uptake in protozoa (James, 1980; Shigeoka et al., 1987) and animal hepatocytes
(Lmneng et al., 1979), erythrocytes, and ghosts (Casirola et al., 1990). Adenine and
adenosine impaired thiamin synthesis in Escherichia coli (Iwashima et al., 1968) by

lowering the hydroxymethylpyrimidine moiety synthesis (Kawasaki et al., 1969).

47

The objective of this series of experiments was to decrease CH, production by
mixed ruminal cultures by blocking pyruvate oxidative decarboxylation. In a first
experiment, it was hypothesized that the combination of thiamin structural analog
amprolium, with adenine or adenosine, would decrease methanogenesis by
simultaneously blocking thiamin intracellular synthesis and its extracellular uptake. In
the second experiment, adenine and adenosine were hypothesized to decrease CH4
production by inhibiting thiamin synthesis. In the third experiment, thiamin structural
analogs amprolium and oxythiamin were hypothesized to decrease CH, production by
impairing thiamin utilization. In the fourth experiment, it was hypothesized that the
direct inhibition of pyruvate oxidoreductases through the use of pyruvate derivatives

(F loumoy and Frey, 1989; Williams etal., 1990) would decrease CH4 production.

Material and Methods
Experiment 1

Arrangement of treatments. Amprolium, a structural analog of thiamin, was used
in conjunction with adenine or adenosine to attempt the simultaneous inhibition of
thiamin intracellular synthesis and external thiamin utilization. A basal, fermentation
medium without thiamin was used. The 2 x 2 x 3 factorial arrangement of treatments
was: 1) thiamin 0 or 10 mM (thiamin effect); 2) amprolium plus (10 mM) or minus
(structural analog effect); 3) adenine (10 mM) , adenosine (10 mM) or control
(intracellular synthesis inhibition effect).

Ruminal fluid collection and preparation. Ruminal fluid was withdrawn from

two mature Holstein cows fed alfalfa hay, mixed, and strained through two layers of

48

cheese cloth. It was blended for 15 5 under 02 free-CO2 and then filtered through one
layer of cheesecloth. A cell washing procedure was then used to eliminate thiamin
present in the liquid phase of the ruminal fluid. Ruminal fluid was centrifuged at 300 x g
and 4 °C for 10 min in capped tubes under C02. The pellet was discarded and the
supernatant centrifuged at 20,000 x g and 4 °C for 20 min. The supernatant was
discarded and an equal volume of buffer (Bryant and Burkey, 1953), previously
autoclaved and anaerobically prepared, was used to resuspend the cell pellet. The last
step was repeated, and 1 mL of the cell suspension was anaerobically delivered into 25-
mL I-Iungate tubes.

Media preparation. Hungate tubes contained 15 mL of an autoclaved, ruminal
fluid-free, thiamin-free medium (Table 2-1), sealed with a rubber stopper under an O2
free-CO2 atmosphere. Forty-one milligrams of adenine [Sigma A 8626] and 21 mg of
adenosine [Sigma A 9251] were added as solids to the corresponding tubes before
delivering the medium, so as to achieve final concentrations of 10 mM. Riboflavin and
thiamin were not autoclaved with the other vitamins in order to avoid their destruction. A
0.75 ppm riboflavin solution was prepared, filter-sterilized, and used to deliver thiamin
hydrochloride [Sigma T 4625] and amprolium [Sigma A 0542] to the corresponding
tubes by anaerobically injecting 0.5 mL into each tube, so as to achieve final
concentrations of 10 mM.. Controls received 0.5 mL of the riboflavin solution without

thiamin hydrochloride or amprolium.

49

Table 2-1. Ruminal fluid-free medium used in Experiment 1

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

Amount
Ingredient (g/L or mL/L)
Cellobiose 1.0
Soluble starch 1.0
Xylose 0.5
Arabinose 0.5
Vitamin-free amino acids‘ 5.0
NaCl ' 2.0
KH2P04 3.0
KZHPO4 3.0
(NH4)2304 1-0
CaCl2 0.2
MgSO4 0.4
Resazurin solution 2.0
Cysteine sulfur solution2 80
Trace mineral solution3 0.2
Valerie acid 0.03
Isovaleric acid 0.03
Isobutyric acid 0.03
Sodium acetate 1.968
Vitamin solution‘ 0.2

 

 

 

 

'Vitamin-free Casaminoacids, Difco Laboratories.

2L-Cysteine HCl, 2.5 g and NazS o 9H20, 2.5 g were added to deionized
water to give a final volume of 200 mL, and pH was adjusted to 6.5 with 3 N NaOH.
Cysteine sulfur solution and NaHCO3 were added after boiling the medium twice as
described by Butine and Leedle (1989).

3H3BO3, 620 ppm; ZnClz, 682 ppm; MnClz 0 4H20, 930 ppm; CoClz o
6H20, 950 ppm; NazMoO4 o 2H20, 360 ppm; NazSiO3, 122 ppm; NazSeO3,
173 ppm; NiClz 0 6H20, 130 ppm; Na2W04 o 2H20, 3 ppm; A12(SO4)3, 0.03 ppm.

‘Pyridoxamine, 1500 ppm; folic acid, 500 ppm; p-aminobenzoic acid, 300

ppm; biotin, 100 ppm; cobalamin, 100 ppm; hemin, 1000 ppm.

50

Incubation. Tubes were incubated in a shaking waterbath for 24 h, and optical
density measured at 600 nm every 6 h to assess microbial growth. At the beginning of
the experiment, three samples of medium with the added cell suspension were frozen for
subsequent determination of VFA initial concentrations. At the end of the incubation,
tubes were allowed to cool to room temperature, and total gas production was measured
as described by (Callaway and Martin, 1996). Fermentation was stopped by adding 1
mL of 12 N H2804.

Analytical procedures. Methane and CO2 were analyzed as described by
Callaway and Martin (1996), using a Gow Mac series 750 flame ionization detector gas
chromatograph (Gow Mac Instruments Co., Bridgewater, NJ) equipped with a 4' x 1/4"
DC 200 column, 5.5. (150 °C, carrier gas was N2 at 820 Kpa). A RGD2 Reduction Gas
Detector (Trace Analytical, Menlo Park, CA) equipped with the same type of column was
used for H2 analysis. Gas production was expressed as umoles at 25 °C and 1 atm. A 5-
mL aliquot of the fermentation medium was centrifuged (26,000 x g, 4 °C, 30 min).
Volatile fatty acids, lactate, formate, and ethanol were quantified by differential
refractometry with a Waters HPLC (Waters Associates Inc., Milford, MA) equipped with
a BioRad I-IPX 87H column (BioRad Laboratories, Hercules, CA). Solvent was 0.005 M
H2804 at 0.6 mL/min. Column temperature was 65 °C. Sample injection volume was 15
uL. Ammonia was analyzed as described by (Chaney and Marbach, 1962).

Calculations. Apparently fermented OM (F OM) was estimated from the VFA

stoichiometry (Demeyer and Van Nevel, 1979), but using isobutyrate instead of caproate:

51

fermented OM (mg of hexose) = (Acetate/2 + Propionate/2 + Isobutyrate + Butyrate +
Valerate + Isovalerate) x 162, with all VF A expressed in mmoles produced.

Statistical analysis. Six replicates per treatment were used. Data from five tubes
(all from different treatments) were discarded because their dark color indicated lack of
reducing conditions. The experimental model was: response = overall mean + thiamin +
amprolium + N base + thiamin x amprolium + thiamin x N base + amprolium x N base +
thiamin x amprolium x N base + residual. Data were tested for homogeneity of variances
using the Modified Levene’s test (N eter et al., 1996) and analyzed as a three-way
AN OVA, when homogeneity of variances was not rejected (P > 0.05). Probabilities of
effects were calculated using Type III sums of squares for an unbalanced design. If
homogeneity of variances was rejected, a Kruskal-Wallis test (Neter et al., 1996) was
conducted. When significant (P < 0.05) effects were found by the AN OVA or the
Kruskal-Wallis tests, and in the absence of significant (P < 0.05) interactions, factor level
means were compared through the Spj ovoll/Stoline test for unequal N (Chew, 1976). If
the interactions were significant, treatment means of one factor within another were
compared through the Spjovoll/Stoline test for unequal N. When the intracellular thiamin
synthesis inhibition effect was significant (P < 0.05), preplanned contrasts tested were: 1)
control vs average of adenine and adenosine, and 2) adenine vs adenosine. The
responses of optical density to time were modeled as fourth order polynomials (N eter et

al., 1996).

52

Experiment 2

The effects of adenine and adenosine on CH4 production and fermentation were
evaluated in batch cultures in vitro, with a ruminal fluid/buffer medium. The effects of
adenosine’s ribose moiety. alone or with adenine were also tested. The experimental
treatments were: 1) Control; 2) Adenine; 3) Ribose; 4) Adenine + ribose; 5) Adenosine.
Ruminal fluid was collected, strained and blended as in Experiment 1, and one part of
ruminal fluid mixed with four parts of buffer (Goering and Van Soest, 1975). At the
begimiing of each experiment, three samples of ruminal fluid and buffer mixture were
frozen for subsequent determination of VFA initial concentrations. Fifty milliliters of
ruminal fluid and buffer mixture were anaerobically delivered into 125-mL Wheaton
bottles. Each bottle had 600 mg of ground (0.2 mm screen) alfalfa hay (11.4% CF in the
DM) as substrate, and adenine [Sigma A 8626], ribose [Sigma R 7500], adenine and
ribose, or adenosine [Sigma A 9251] added so as to achieve 10 mM final concentrations
of each of the compounds. The bottles were sealed under an O2 free-CO2 atmosphere and
incubated in a shaking water bath at 39 °C for 24 h. Fermentation was terminated by
injecting 3 mL of 12 N H2804 into each bottle. Total gas production and composition,
VFA and NH: analysis, and the calculation of apparently fermented OM, were done as in
Experiment 1.

F our replicates per treatment were used. The experimental model was: response =
overall mean + treatment + residual. Data were tested for homogeneity of variances
using the Modified Levene’s test (N eter et al., 1996) and analyzed as a one-way AN OVA,
when homogeneity of variances was not rejected (P > 0.05). If homogeneity of variances

was rejected, a Kruskal-Wallis test (N eter et al., 1996) was conducted. Planned contrasts

53

of interest were 1) control vs adenine; 2) control vs adenosine; 3) adenine vs adenosine;
4) ribose vs adenosine; 5) adenine + ribose vs adenosine. A Bonferroni adjustment
(Neter et al., 1996) for five non-orthogonal comparisons was used, and significance
declared at P < 0.01 (for an experimentwise type I error probability of 0.05). All other

comparisons were done by the Scheffé test.

Experiment 3

The effects of thiamin structural analogs amprolium and oxythiamin on CH4
production and fermentation were evaluated in batch cultures in vitro, with a ruminal
fluid and buffer medium. The experimental treatments were 1) Control; 2) Oxythiamin 5
mM; 3) Oxythiamin 10 mM; 4) Amprolium 5 mM; 5) Amprolium 10 mM. Ruminal fluid
was collected and prepared as in Experiment 2. Delivery of the ruminal fluid/buffer
mixture and incubation procedures were the same as in Experiment 2. Each bottle had
200 mg of ground (0.2 mm screen) alfalfa hay (11.4% CF in the DM) as a substrate. One
milliliter of 0.255 or 0.510 M solutions of amprolium or oxythiamin were added to the
corresponding Wheaton bottles so as to achieve final concentrations of 5 or 10 mM.
Controls received 1 mL of deionized water. Fermentation was terminated by injecting 1
mL of a 10% phenol solution, instead of H280,” as in Experiments 1 and 2, so that final
pH could be measured (Digital Benchtop pH Meter, Cole-Parmer Instrument Company,
Vernon Hills, IL). Total gas production and composition, VFA and NH,+ analysis, and

the calculation of apparently fermented OM, were done as in Experiment 1.

54

The number of replicates, the model used and the statistical analyzes were the
same as in Experiment 2. Orthogonal polynomial contrasts for linear and quadratic

responses to oxythiamin and amprolium were evaluated (N eter et al., 1996).

Experiment 4

The effects of pyruvate derivatives on CH4 production and fermentation were
evaluated in batch cultures in vitro, with a ruminal fluid and buffer medium. The
experimental treatments were: 1) Control; 2) Pyruvate (positive control); 3)
Bromopyruvate; 4) Fluoropyruvate; 5) Hydroxypyruvate. Ruminal fluid was collected
and prepared as in Experiment 2. Delivery of the ruminal fluid/buffer mixture and
incubation procedures were the same as in Experiment 2. Each bottle had 300 mg of
ground (0.2 mm screen) alfalfa hay (11.4% CF in the DM) as a substrate. One milliliter
of 0.102 M solutions of Na—pyruvate [Sigma P 2256], bromopyruvate [Sigma B 9630],
Na-fluoropyruvate [Sigma F 4004] and hydroxypyruvate [Sigma H 9270] were added to
Wheaton bottles so as to achieve final concentrations of 2 mM. Controls received 1 mL
of deionized water. Fermentation was terminated by injecting 1 mL of a 10% phenol
solution. Total gas production was measured, and gas composition analyzed as in
Experiment 1. Analysis of VFA and NH], and the calculation of fermented OM, were
done as described above. The additives final concentrations were determined by HPLC
along with the VFA.

The number of replicates, the model used, and the statistical analyzes were the
same as in Experiment 2. Planned contrasts of interest were 1) control vs Na-pyruvate;

2) control vs average of pyruvate derivatives; 3) Na—pyruvate vs average pyruvate

55

derivatives; 4) bromopyruvate vs Na—fluoropyruvate; 5) bromopyruvate vs
hydroxypyruvate; 6) Na—fluoropyruvate vs hydroxypyruvate. A Bonferroni adjustment
(N eter et al., 1996) for six non-orthogonal comparisons was used, so significance was
declared at P < 0.0083 (for an experimentwise type I error probability of 0.05). Other

comparisons of interest were done using the Scheffe test (N eter et al., 1996).

Results and Discussion
Experiment 1

. Thiamin or amprolium addition did not influence bacterial growth, as measured
through optical density (data not shown) or fermented OM (Table 2-2). However,
fermented OM (Table 2-2), and optical density at 6, 12, 18 and 24 h (Figure 2-1), were
strongly increased by adenosine, and inhibited by adenine.

Total OM fermentation (OM in medium plus additives) was very low in all
treatments, ranging from 5.4 to 21.1% (Table 2-2). There was no increase in optical
density in control. When the estimated fermented OM was related to the carbohydrates
initially present in the medium only (i.e., ignoring the amino acids and additives),
fermented OM was higher, ranging between 30.6 and 81.1% (data not shown).

Adenosine addition stimulated (P < 0.01) OM fermentation (16.6% vs 10.6% of total OM
in control and added adenosine, respectively). Adenine, in contrast, inhibited (P = 0.02)
OM fermentation (8.4% vs 10.6% of total OM with and without adenine, respectively). It
has been shown that adenosine, but not adenine, could support growth of Prevotella

(Bacteroides) ruminicola and Selenomonas ruminantium as the only energy source

56

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(Cotta, 1990). It is possible that adenosine stimulated the grth of those species in the

present experiment.

 

 

 

 

 

E
C
O
O
(0
iii
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’6
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O.)
3
g ‘0‘ control
0 MSE = 0.0502
"0., adenine
. MSE = 0.0362
01 L - A - A , "In adenosine
' o 6 12 18 24 MSE = 00522

Time (h)

Figure 2-1. Effects of the addition adenine or adenosine on optical density of ruminal

bacterial cells

Amprolium addition decreased (P < 0.01) CH, production by 42% (3.6 vs 6.2
umol; Table 2-2). Surprisingly, the addition of thiamin also decreased (P = 0.04) CH,
production by 22%. This result contradicts the hypothesis that thiamin would be used by
ruminal microorganisms for pyruvate oxidative decarboxylation, a reaction that provides
precursors for CH, formation. It agrees, though, with previous work in which thiamin
addition decreased CH, production in continuous culture between 6 and 22% (Alves de
Oliveira et al., 1996). However, in another study, there was no effect of thiamin addition

on CH, production (Alves de Oliveira et al., 1997). Thiamin may have stimulated an

58

alternative H sink in the present experiment, therefore, decreasing methanogenesis. In
agreement with Alves de Oliveira et a1. (1997), thiamin addition did not have an effect
on CO2 release in the present experiment. Neither did amprolium, which would suggest
that this structural analog of thiamin did not decrease CH, production by blocking
pyruvate oxidative decarboxylation.

Adenine and adenosine decreased (P < 0.01) CH, production by 47 (4.1 vs 8.0
umol) and 76% (2.7 vs 8.0 umol), respectively. We are unaware of previous reports
measuring the effects of nucleotides or nucleosides on CH, production in the rumen. We
had hypothesized that the addition of adenine or adenosine would hinder thiamin
intracellular synthesis. This would result in thiamin not being available to act as a
cofactor in pyruvate oxidative decarboxylation, and the reaction would be blocked.
Ultimately, this would decrease the availability of CO2 and reducing equivalents for CH,
production. Although both adenine and adenosine decrease CH, production, the addition
of thiamin did not supress their effect on methanogenesis. This suggests that the decrease
in methanogenesis caused by adenine and adenosine was unrelated to their hypothesized
inhibition of thiamin intracellular synthesis.

Thiamin addition decreased (P < 0.01) propionate molar percentage. In
agreement, Alves de Oliveira et al. (1996) found that the addition of thiamin to a ruminal
continuous culture (with normal, but not with salts-reduced, artificial saliva) decreased
propionate, and increased acetate molar percentage. Theoretically, one would expect this
result, if thiamin was used as a cofactor of pyruvate oxidative decarboxylation. However,
the addition of thiamin to the diet of sheep that served as donors of ruminal fluid

increased propionate molar percentage in ruminal fermentation in vitro at the expense of

59

butyrate (Naga et al., 1975; Candau and Kone, 1980). Naga et al. (1975) considered that
thiamin could have altered the VFA pattern directly, or indirectly by reducing ruminal
motility and outflow in the thiamin deficient animals. Candau and Kone (1980)
speculated that thiamin could act at the re-oxidation of intracellular cofactors,
withdrawing reducing equivalents from methanogenesis and diverting them to the
reduction of lactate to propionate. Alves de Oliveira et al. (1997) did not find an effect of
thiamin addition on the VFA profile. The reasons for the discrepancies between
experiments are unknown. In the present experiment, amprolium decreased (P < 0.01)
acetate molar percentage, especially when thiamin was added. Adenosine, in contrast,
prevented the decrease in acetate molar percentage caused by amprolium (P = 0.03). As
adenosine itself strongly decreased (P < 0.01) acetate molar percentage, it is possible that
both amprolium and adenosine acted on the same species of acetate producers, as their
eflects were not additive. In the absence of added thiamin, amprolium decreased (P =
0.01) propionate molar percentage. This was contrary to the experiment’s hypothesis, for
it was expected that an inhibition of pyruvate oxidative decarboxylation would divert the
C in pyruvate from acetate and butyrate to propionate formation. However, when thiamin
was present, amprolium increased (P = 0.02) propionate molar percentage. This
interaction is difficult to interpret, and, considering also that the inhibition of CH,
formation by amprolium was independent from thiamin addition, it is likely that
amprolium did not decrease CH, production by inhibiting pyruvate oxidative
decarboxylation.

Adenosine decreased (P < 0.05) butyrate molar percentage and greatly stimulated

(P < 0.05) propionate. Adenine decreased (P < 0.05) butyrate molar percentage.

60

Thiamin addition decreased (P < 0.01) NH,+ concentrations (22.3 vs 20.1 mg/dL).
This could be due to less fermentation of amino acids, or to an increase in microbial
protein synthesis, as found by Candau and Kone (1980). In contrast, Naga et al. (1975)
found a decrease in microbial growth as a result of adding thiamin. Alves de Oliveira et
al. (1996, 1997) did not find any effect of thiamin addition upon microbial protein
synthesis. Amprolium also decreased (P = 0.01) NH,+ concentration, while adenosine
increased (P < 0.05) it. This could be explained by the deamination occurring in the
catabolism of adenosine to hypoxanthine (V oet and Voet, 1995). The presence of the
ribose moiety seemed to have been a requirement for deamination to occur, as the
addition of adenine tended (P = 0.08) to decrease NH,+ concentrations.

The ratio of CO2 to CH, in the triple control (thiamin, no amprolium, and no
adenine or adenosine) was almost of 50 to 1, which is unusually large (Moss, 1993).
Microbial growth in the control, as estimated through the increase in optical density, was
almost non-existent (Figure 2-1). Some of the experimental procedures (washing the
cells with a buffer, absence of ruminal fluid in the medium, use of very rapidly
fermentable substrates) could have biased the microbial community that survived, and
created an atypical fermentation. Amprolium, adenine, and adenosine decreased
methanogenesis, although the mechanisms appeared to differ from the original
hypothesis. In Experiments 2 and 3, therefore, the effects of adenine, adenosine, and
amprolium, were studied under more classical in vitro procedures. As the mechanisms by
which adenine and adenosine decreased CH, production in Experiment 1 were unrelated
to the addition of amprolium, we studied them separately, adenine and adenosine in

Experiment 2, and amprolium in Experiment 3.

61

Experiment 2

In Experiment 1, the strong inhibition of methanogenesis caused by adenine and
adenosine was independent of thiamin addition. In Experiment 1, adenine and adenosine
had opposite effects on fermented OM and microbial growth. As this could be caused by
adenosine’s ribose moiety, a treatment with ribose alone, and a treatment with adenine
and ribose, were also included. Adenine, with or without ribose, did not affect CH,
production (Table 2-3). The addition of ribose alone increased (P = 0.03), and of
adenosine tended to increase (P = 0.07), methanogenesis. However, when CH, output
was related to F OM, there were no differences among treatments.
The addition of ribose, either pure (control vs ribose, and adenine vs adenine + ribose) or
as part of the adenosine molecule (adenine vs adenosine), always promoted (P < 0.05)
OM fermentation (Table 2-3). On the contrary, adenine was inhibitory to fermentation.
The addition of ribose, alone (control vs adenine + ribose, P = 0.69, Scheffe’ test), or as
the ribose moiety in adenosine (control vs adenosine, P = 0.58) relieved the inhibition
caused by adenine. If, as hypothesized, adenine caused an inhibition of thiamin
synthesis, the reactions of the pentose phosphate pathway catalysed by transketolase
could have been affected. However, this should not affect the supply of ribose, as ribose
phosphate is a substrate, rather than a product, of these reactions (V oet and Voet, 1995).
Therefore, the relief of adenine’s fermentation inhibition by ribose would have been due
to the use of the latter as an energy source. Although different ruminal microbial Species
differed in their ability to use ribose as an energy source, Selenomonas ruminantium
strains could attain growth rates only slightly lower than with glucose (Cotta, 1990). The

addition of adenine increased (P < 0.01) acetate molar percentage and decreased (P <

62

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0.01) pr0pionate, while adenosine decreased (P < 0.01) acetate and increased (P < 0.01)
butyrate. Ribose increased (P < 0.01) butyrate molar percentage.

It was shown that 62% of added adenine (approximately 1.2 mM) was degraded
after 4 h of incubation in ruminal fluid (McAllan and Smith, 1973). Hypoxanthine and
xanthine accounted for 33 and 7%, respectively, of the adenine initially present. All of
the added adenosine (approximately 1.2 mM) was degraded within 1 h, and 78% of its
initial concentration was recovered as inosine, which was in turn converted to
hypoxanthine. Therefore, deamination seemed to proceed faster when the ribose moiety
was present. In Experiment 1, adenosine, but not adenine, increased NH; concentrations.
In Experiment 2, the addition of adenine, with (P = 0.02) or without ribose (P < 0.01),
and of adenosine (P < 0.01), increased NH; concentrations; however, NH; was higher
(P < 0.01) with adenosine than with adenine. Ruminal protozoa have been shown to
catabolize adenine to xanthine and hypoxanthine (Coleman and Laurie, 1974, 1977);
however, we are not aware of reports on their use of adenosine. In Experiment 1, the low
speed centrifugation for isolating the cell pellet must have removed the protozoa. If
protozoa metabolize more adenine in relation to adenosine than bacteria, that could
explain why adenine increased NH; concentrations in Experiment 2 but not in
Experiment 1. Also, Cotta (1990) found that ruminal bacteria differed in their ability to
use nucleosides, N bases, and ribose. It is then possible that the procedures used in
Experiment 1 biased the microbial population so as to decrease the catabolism of adenine.

The effects of adenine and adenosine on fermentation were quite different from
Experiment 1. A washed cell suspension was used in Experiment 1, whereas, a crude

ruminal fluid and buffer mixture was used in Experiment 2. Differences in the microbial

64

species present must have existed between the two experiments. The ratio of CO2 to CH4
in Experiment 2 was more typical of a ruminal fermentation (Moss, 1993) . Therefore, it

is concluded that Experiment 1 results did not represent a typical ruminal fermentation.

Experiment 3

The depression in CH4 production caused by amprolium in Experiment 1 was
independent of the addition of either thiamin, adenine, or adenosine. Hence, the effects
of amprolium and another thiamin structural analog, oxythiamin, were studied in batch
fermentation with ruminal fluid. Methane production was decreased (P < 0.05) by 23
(244 vs 315 umol) and 8% (289 vs 315 umoles) by oxythiamin and amprolium at 5 mM,
respectively (Table 2-4). Increasing the concentrations to 10 mM did not decrease
methanogenesis further. The effects of amprolium on CH4 production were due to less
fermentation, as CH4 production per milligram of FOM did not change. However,
oxythiamin addition decreased (P < 0.01) CH4 production per milligram of FOM by
about 17% at both concentrations. As H2 concentration did not increase, a direct effect on
methanogens seems unlikely. Oxythiamin decreased (P < 0.01) the molar percentage of
acetate and butyrate and increased (P < 0.01) propionate. This would agree with an
inhibition of pyruvate oxidative decarboxylation; however, there is not evidence that this
was the mechanism by which methanogenesis was decreased. Increases in propionate
molar percentage and decreases in acetate, when methanogenesis is inhibited, have been
previously reported (Russell and Martin, 1984; Martin and Macy, 1985; Garcia-Lopez et
al., 1996). The changes in VFA profile may be a consequence of the decrease in

methanogenesis rather than the latter been caused by the increase in propionate.

65

Table 2-4. Effect of thiamin structural analogs on ruminal in vitro fermentation (Exp. 3)

 

 

 

 

 

 

 

 

 

 

 

 

 

Control Oxyl Oxy1 Amp1 AmpI SEM P = Significant
5 mM 10 mM 5 mM 10 mM contrasts2

CH4. umol 315 244 239 289 279 5.8 < 0.01 1, 2, 3
C02, umol 646 647 643 674 705 30.7 0.58“ None
H2, pmol 0.43 0.46 0.49 0.60 0.48 0.07 0.58 None
Total VFA, 48.9 47.7 47.1 46.9 45.8 0.28 0.02“ 1, 3
mM
Acetate, 814 713 697 744 724 20.3 < 0.014 1, 3
mo]
Propionate, 298 353 348 280 251 5.3 < 0.01 1, 2, 3
umol
Butyrate, 74.3 58.8 56.6 61.3 55.6 3.66 < 0.01 1, 2, 3
pmol
FOM3, °/o 56.7 53.2 51.5 51.4 48.2 1.50 0.02 1, 3
CPL/POM, 2.99 2.48 2.49 3.07 3.12 0.012 < 0.01 l
umol/mg
pH 7.3 7.2 7.1 7.2 7.2 0.07 0.18 1
NH;, 30.7 27.9 24.8 29.3 28.2 0.60 < 0.01 l, 3
mg/dL

 

 

 

 

 

 

 

 

 

fOxy = oxythiamin; Amp = amprolium.
21 = significant (P < 0.05) linear relationship for oxythiamin; 2 = significant (P < 0.05)
quadratic relationship for oxythiamin; 3 = significant (P < 0.05) linear relationship for
amprolium; 4 = significant (P < 0.05) quadratic relationship for amprolium.
3FOM = Apparently fermented OM
4The Kruskal - Wallis test was done due to heterogeneity of variances (Levene test on absolute

 

deviations P < 0.05).

Amprolium increased (P < 0.01) acetate and decreased butyrate (P = 0.02) molar
percentage. It was previously found that amprolium decreased propionate molar
percentage, without affecting the rest of the VFA profile (Horton and Stockdale, 1979).

A decrease in propionate, and an increase in butyrate, were reported when amprolium was
added to ruminal continuous cultures (Heitmann and Yehya Taka, 1970-1971). Results
from those studies the current experiment are contrary to the hypothesis that amprolium
would impair thiamin utilization for thiamin oxidative decarboxylation, resulting in

pyruvate carbon being diverted from acetate and butyrate towards propionate.

An antimethanogenic strategy based on the use of oxythiamin would require
the delivery of ruminal-protected thiamin together with oxythiamin, so as to avoid
potential toxic effects of the latter on the host animal. Given the small amounts that
methanogenesis was decreased by oxythiamin, further work in this line is not

recommended.

Experiment 4
None of the additives had any effect on CH4 or CO2 production (Table 2-5).
Disappearance of all four additives was complete (data not shown). Added pyruvate,
bromopyruvate or fluoropyruvate did not change the VFA molar proportions. This
suggests that bromopyruvate and fluoropyruvate were dehalogenated, converted into
pyruvate and metabolized. Hydroxypyruvate, however, shifted (P < 0.05) the VFA molar
proportions towards butyrate at the expense of acetate, which may indicate that part of
this additive was catabolized by an alternative pathway. Interestingly, hydroxypyruvate
increased total OM (substrate + additive) and the alfalfa substrate fermentation by 6.4 (P
< 0.01) and 5.3 (P = 0.02) percentage units, respectively. As total CH4 production was
not affected, CH4 per milligram of FOM tended (P = 0.05) to be 16% lower than in the
control.

The pyruvate derivatives did not inhibit CH, production, and, except for
hydroxypyruvate, did not alter the VFA profile. This indicates that they did not inhibit
pyruvate oxidoreductases as it was hypothesized. As they were totally metabolized, they

must have been taken up by the cells. They might have been metabolized before they

67

 

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could inhibit pyruvate oxidoreductases, or pyruvate oxidoreductases structures of ruminal

microorganisms could be different from the ones previously reported.

Implications

Pyruvate oxidative decarboxylation in ruminal fluid could not be inhibited, either
directly, or through the inhibition of thiamin utilization. It is not known if the additives
were taken up by ruminal microorganisms, and, if they were, why the intracellular effects
hypothesized did not occur. If further work in this line is to be considered, basic research
on thiamin uptake, synthesis and utilization, and on the enzymology of pyruvate

oxidoreductases of different ruminal microorganisms would be needed.

69

 

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Moss, A. R. 1993. Methane. Global Warming and Production by Animals. lst ed.
Chalcombe Publications, Kingston, Kent, UK.

Naga, M. A., J. H. Harmeyer, H. Holler, and K. Schaller. 197 5. Suspected "B"- vitamin
deficiency of sheep fed a protein-free urea containing purified diet. J. Anim. Sci.
40: 1192-1198.

Neter, J ., M. H. Kutner, C. J. Nachtsheim, and W. Wasserman. 1996. Applied Linear
Statistical Models. 4th ed. McGraw-Hill, Boston.

Russell, J. B. and S. A. Martin. 1984. Effects of various methane inhibitors on the
fermentation of amino acids by mixed rumen microorganisms in vitro. J. Anim.
Sci. 59: 1329-1338.

Russell, J. B. and R. J. Wallace. 1997. Energy-yielding and energy-consuming reactions.
In: P. N. Hobson and C. S. Stewart (eds.) The Rurnen Microbial Ecosystem. p
246-282. Blackie Academic and Professional, London.

Shigeoka, S., T. Onishi, K. Maeda, Y. Nakano, and S. Kitaoka. 1987. Thiamin uptake in
Euglena gracilis. Biochim. Biophys. Acta 929: 247-252.

Uyeda, K. and J. C. Rabinovvitz. 1971. Pyruvate-ferredoxin oxidoreductase. III.
Purification and properties of the enzyme. J. Biol. Chem. 246: 3111-3119.

Voet, D. and J. G. Voet. 1995. Biochemistry. 2nd ed. John Wiley and Sons, Inc., New
York.

Williams, K. P., P. F. Leadlay, and P. N. Lowe. 1990. Inhibition of pyruvatezferredoxin
oxidoreductase from Trichomonas vaginalis by pyruvate and its analogues.
Comparison with the pyruvate decarboxylase component of the pyruvate
dehydrogenase complex. Biochem. J. 268: 69-75.

Wolin, M. J ., T. L. Miller, and C. S. Stewart. 1997. Microbe-microbe interactions. In: P.

N. Hobson and C. S. Stewart (eds) The Rurnen Microbial Ecosystem. p 467-491.
Blackie Academic and Professional, London.

72

CHAPTER 3

Use of some novel alternative electron sinks to inhibit ruminal methanogenesis

Abstract

Several compounds were evaluated in vitro as alternative electron sinks to ruminal
methanogenesis. They were incubated with ruminal fluid, buffer mixture, and finely
ground alfalfa hay for 24 h, at O, 6, 12, and 18 mM initial concentrations. The propionate
enhancer oxaloacetic acid, the butyrate enhancer B-hydroxybutyrate, and the butyrate
unsaturated analog 3-butenoic acid were ineffective in decreasing methanogenesis.
Nevertheless, B-hydroxybutyrate increased the apparent OM fermentability of the alfalfa
hay substrate from 58.0 to 63.4%, and 3-butenoic acid seemed to increase it from 62.0 to
73.7%. Almost all of added oxaloacetic acid and most of acetoacetate disappeared during
the incubation, while only between 30.3 and 53.4% of B-hydroxybutyrate disappeared.
The butyrate enhancers acetoacetate and crotonic acid, and the butyrate unsaturated
analog 2-butynoic acid, decreased methanogenesis by a maximum of 18, 9 and 9%,
respectively. Crotonic acid at 18 mM initial concentration seemed to increase the
substrate apparent OM fermentability from 57.0 to 68.2%. Between 78.6 and 100% of
acetoacetate disappeared during the incubation. The propionate unsaturated analog
propynoic acid, and the unsaturated ester ethyl 2-butynoate, decreased methanogenesis by
a maximum of 76 and 79%, respectively. Less than 5% of propynoic acid disappeared.
The substrate apparent fermentability was decreased by propynoic acid from 62.0 to

57.4%, and seemed to have been decreased by ethyl 2-butynoate from 62.0 to 29.3%.

73

More accurate measurements of the disappearance of some of the compounds studied are

needed to better understand how they are metabolized and how they affect fermentation.

Introduction

Methane emission is an energy loss for ruminants, and also causes global
warming (Moss, 1993). It would be beneficial both for the efficiency of production and
the environment to divert reducing equivalents from ruminal methanogenesis into
alternative electron sinks with a nutritional value for the host animal (Schulrnan and
Valentino, 1976), such as propionate (Callaway and Martin, 1996).

Intermediates of the fermentation pathways that lead to propionate (“propionate
enhancers”) have been studied as alternative electron sinks to ruminal methanogenesis.
Compounds that accept one pair of electrons in their conversion into propionate include
malate (Martin and Streeter, 1995; Callaway and Martin, 1996; Lopez et al., 1999),
fumarate (Callaway and Martin, 1996), lactate, and acrylate (Lopez et al., 1999).
Oxaloacetate, however, accepts two pairs of electrons, and, theoretically, should be more
effective in competing with methanogenesis as an alternative electron sink. To our
knowledge, oxaloacetate has not been examined for this purpose.

Likewise, intermediates in the conversion of pyruvate into butyrate (“butyrate
enhancers”) also accept reducing equivalents (Miller and Jenesel, 1979). Butyrate
enhancers have not been studied as alternative electron sinks to ruminal methanogenesis.
Also, unsaturated analogs of propionate and butyrate with double and triple bonds could
be reduced to these VFA, redirecting reducing equivalents away from CH, formation.

These compounds, which are not normal intermediates of ruminal fermentation (except

74

 

for acrylate and crotonate), have not been studied as alternative electron sinks to ruminal
methanogenesis.

The objective of this study was to evaluate the effects of oxaloacetate, butyrate
enhancers, and unsaturated organic acids and esters on in vitro fermentation by mixed
ruminal microbial cultures. It was hypothesized that the addition of these compounds

would decrease CH, production in vitro by utilizing reducing equivalents.

Materials and Methods
Additives and concentrations

The intermediate of the propionate pathway, oxaloacetic acid [flee acid, Sigma 0
4126]; three intermediates of the butyrate pathway (Miller and Jenesel, 1979),
acetoacetate [Li salt, Sigma A 8509], B-hydroxybutyrate [Na salt, Sigma H 6501], and
crotonic acid [flee acid, Sigma C 4630]; the unsaturated propionate analog propynoic acid
[flee acid, Acros 13150-0100]; the unsaturated butyrate analogues 3-butenoic acid [flee
acid, Acros 15883-0250], and 2-butynoic acid [flee acid, Acros 30806-0010]; and the
unsaturated ester ethyl 2-butynoate [Aldrich 4341-76-8] were examined as alternative
electron sinks to ruminal methanogenesis in vitro. Each of the additives, except for ethyl
2-butynoate, was added to Wheaton bottles as 1 mL aqueous solutions, so as to achieve 6,
12 and 18 mM initial concentrations, respectively. The hydrophobic ester, ethyl 2-
butynoate, was added directly as a liquid (35.7, 71.3, and 107.0 uL, to achieve 6, 12 and
18 mM initial concentrations, respectively) together with 1 mL of deionized water.
Controls received 1 mL of deionized water. The initial concentrations, which could be

considered as relatively high, were chosen based on the additives hypothesized mode of

75

 

 

 

action: the effectiveness of an additive for withdrawing electrons flom methanogenesis
should be stoichiometrically related to the amount of additive reduced. As this was the
first time these compounds were studied, a wide range of initial concentrations was
chosen. Similar ranges of initial concentrations have been used in other studies that
evaluated alternative electron sinks to ruminal methanogenesis (Martin and Streeter,
1995; Callaway and Martin, 1996; Lopez et al., 1999).

Oxaloacetic acid, acetoacetate, B-hydroxybutyrate, and crotonic acid were
examined together in two experimental runs, while propynoic acid, 3-butenoic acid, 2-
butynoic acid, and ethyl 2-butynoate were examined in the third and fourth experimental

runs.

Ruminal fluid collection and incubation

Ruminal fluid was withdrawn two hours after the morning feeding from two
mature Holstein cows fed alfalfa hay. It was mixed together, and strained through two
layers of cheesecloth. It was then blended for 15 5 under C02, and again strained through
two layers of cheesecloth. One volume part of ruminal fluid was mixed with four volume
parts of a bicarbonate and phosphate buffer (Goering and Van Soest, 1975), and 50 mL of
the ruminal fluid and buffer mixture anaerobically delivered into 125-mL Wheaton
bottles. All the bottles contained 300 mg of ground (0.2 mm screen mesh) alfalfa hay
(11.4% CP in the DM) as substrate. Three samples of the ruminal fluid and buffer
mixture were flozen for subsequent determination of VFA initial concentrations. Bottles
were sealed under an Oz-flee CO2 atmosphere, and incubated in a shaking water bath at

39 °C for 24 h. At the end of the incubation, bottles were allowed to cool to room

76

 

 

 

temperature, and total gas production volume was measured (Callaway and Martin,

1996). Fermentation was then stopped by adding 1 mL of a 10% phenol solution.

Analytical procedures

Methane and CO2 were analyzed (Callaway and Martin, 1996), using a Gow Mac
series 750 flame ionization detector gas chromatograph (Gow Mac Instruments Co.,
Bridgewater, NJ) equipped with a 4' x 1/ " DC 200 stainless steel column (150 °C, carrier
gas was N2 at 820 kPa). A RGD2 Reduction Gas Detector (Trace Analytical, Menlo
Park, CA), equipped with the same type of column, was used for H2 analysis. The
volume of gas produced was expressed as umoles at 25 °C and 1 atm. A S-mL aliquot
was centrifuged (26,000 x g, 4 °C, 30 min), and the pH was measured in the supematants
(Digital Benchtop pH Meter, Cole-Partner Instrument Company, Vernon Hills, IL).
Volatile fatty acids, lactate, formate, ethanol, and the chemical additives were quantified
by differential reflactometry with a Waters HPLC (Waters Associates Inc., Milford, MA)
equipped with a BioRad HPX 87H column (BioRad Laboratories, Hercules, CA).
Separation was done by ion moderated partition. Solvent was 0.005 M HZSO, at 0.6
mL-min". Column temperature was 65 °C. Sample injection volume was 15 uL.

Ammonia was analyzed as reported before (Chaney and Marbach, 1962).

77

Calculations

Based on known biochemical pathways, some of the fermentation intermediates
added were not expected to produce gases. Consequently, calculations based on VFA
production stoichiometry (Marty and Demeyer, 197 3) would have then overestimated
apparently fermented OM (F OM). Therefore, FOM and substrate apparently fermented
were calculated by mass balance flom the net production of VFA, lactate, gases, and
ammonia. As ethanol, formate, and succinate accumulated in some of the treatments,

they were also included in the calculation:

F OM (%) = (gases + VFA + lactate + ethanol + formate + succinate + NH]) x 100
/(substrate OM + additive OM), with all fermentation products produced, substrate and

additives expressed in grams.

Substrate apparently fermented (%) =

(F OM (g) - additive disappeared during fermentation (g)) x 100 /(substrate OM (g))

Crotonic acid and 2-butynoic acid co-eluted off the HPLC column with
isovalerate and isobutyrate, respectively. As the amounts of isovalerate and isobutyrate
produced are relatively minor in comparison to acetate, propionate, and butyrate,
isovalerate was not included in the calculations for estimating FOM in the crotonic acid
treatments, and isobutyrate in the 2-butynoic acid treatments. Disappearance of crotonic
acid and 2-butynoic acid are not reported. Similarly, disappearances are not reported for

3-butenoic acid and ethyl 2-butynoate, as these additives co-eluted off the HPLC colunm

78

 

with propionate and butyrate, respectively. Organic matter and substrate fermentation are
not reported for these additives.

Hydrogen balances were calculated (Marty and Demeyer, 1973), with net
production of ammonia (one mole of ammonia produced releases one mole of reducing
equivalent pairs) also considered. Net production of ethanol, lactate, and formate were
also considered, ethanol and lactate formations releasing and accepting one pair of
reducing equivalents each (V oet and Voet, 1995), and formate incorporating one pair of

reducing equivalents (Russell and Wallace, 1997):

H produced (umoles) = 2A + P + 4B + 3V + NH,+ + E + L
H incorporated (umoles) = 2P + 2B + 4V + 4CH4 + H2 + F + E + L

H recovery (%) = H incorporated x 100 / H produced

where A = acetate, P = propionate, B = butyrate, V = valerate, E = ethanol, L =
lactate, and F = formate, all expressed as umoles. VFA and lactate were considered as
nutritionally useful H sinks, while methane, dihydrogen, formate, and ethanol were
considered as H sinks without a nutritional value. The H balance was not calculated for
3-butenoic acid and ethyl 3-butynoate, as these additives co-eluted off the HPLC column

with propionate and butyrate.

Statistical analysis
Two replicates per compound and concentration were used in each of the two

experimental runs. The experimental run was modeled as a random block (N eter et al.,

79

1996): observation = overall mean + additive concentration + run + residual. Orthogonal
contrasts were performed to determine linear, quadratic, and cubic effects of

concentration. Significance was declared at P < 0.05.

Results
Oxaloacetic acid
Production of CH, linearly increased (P < 0.01) by 7, 8, and 13%, at 6, 12, and 18
mM initial concentration of oxaloacetic acid, respectively (Table 3-1). The release of
CO2 was linearly increased (P < 0.01). H2 accumulation was similar to control.
Oxaloacetic acid was almost totally fermented. There was a linear increase in
total VFA concentration (P < 0.01), and acetate (P < 001), propionate (P = 0.01),
butyrate (P < 0.01), valerate (P = 0.01), and isovalerate (P < 0.05) production.
Production of isobutyrate, final pH, and NH,+ concentration were not affected.
Oxaloacetic acid linearly decreased (P < 0.01) the alfalfa substrate apparent
fermentability flom 58.0 to 35.8%. As the amount of FOM increased due to the additive
disappearance, CH, production per milligram of F OM was decreased (P < 0.01) by

oxaloacetic acid.

80

Table 3-1. Effects of the addition of oxaloacetic acid on in vitro ruminal fermentation

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

Linear Quadratic Cubic
Initial concentration, mM effect effect effect SEM
P = P = P =

0 6 12 18
CH4, umol 421 452 454 476 < 0.01 0.46 0.13 5.65
C02, umol 911 1091 1276 1327 < 0.01 0.37 0.98 40.1
Hz, umol 0.41 0.46 0.44 0.61 0.01 0.36 0.47 0.038
Additive - 302 606 912 < 0.01 0.87 - 5.25
disappearance,
umol
Additive - 98.7 99.0 99.3 0.64 0.98 - 0.896
disappearance,
%
Total VFA, 54.5 59.4 64.2 66.5 < 0.01 0.13 0.49 0.783
mM
Total VFA 1657 1917 2175 2296 < 0.01 0.13 0.49 41.5
production,
umol
Acetate, umol 1111 1330 1547 1688 < 0.01 0.14 0.51 23.8
Propionate, 345 365 377 372 0.01 0.09 0.81 6.67
umol
Butyrate, umol 136 151 165 172 < 0.01 0.51 0.75 5.84
Isobutyrate, 16.9 17.1 26.8 5.40 0.34 0.08 0.14 5.47
umol
Valerate, umol 20.4 23.3 26.4 26.6 0.01 0.25 0.56 1.12
Isovalerate, 27.8 30.7 32.9 33.6 < 0.05 0.89 0.82 0.071
mol
FOM, % 58.0 58.2 54.3 55.2 0.28 0.88 0.43 2.43
Substrate 58.0 52.3 41.2 35.8 < 0.01 0.96 0.44 3.09
fermentability',
%
CH4/FOM, 2.62 2.45 2.14 2.17 < 0.01 0.20 0.19 0.007
mong
Final pH 6.86 6.99 6.79 6.94 0.92 0.87 0.10 0.080
NH4+, mg/dL 26.2 24.9 23.4 25.2 0.32 0.13 0.41 0.906
H produced, 3224 3854 4401 4673 < 0.01 0.19 0.50 63.4
umol
H incorporated, 2719 2933 3040 3103 < 0.01 0.21 0.60 33.6
tunol
H recovery, % 81.8 76.2 68.5 66.5 < 0.01 0.02 0.03 0.40
Nutritionally 38.4 3 8.3 40.2 38.4 0.48 0.24 0.12 0.36
useful H, %
Nutritionally 61.6 61.7 59.8 61.6 0.48 0.24 0.12 0.36
non-useful I-I,
%

 

 

 

 

 

 

 

 

 

 

 

"Substrate apparently fermented (%) = (F OM (g) - additive disappeared (g)) x 100 /
(substrate OM (g))

81

Oxaloacetic acid linearly increased (P < 0.01) both H produced and incorporated,
but decreased (P < 0.01) H recovery flom 81.8 to 66.5%. The percentage of nutritionally

useful H incorporated was not affected by oxaloacetate (Table 3-1).

Acetoacetate

Addition of acetoacetate linearly decreased (P = 0.03) CH, production by 5, 18,
and 10% at 6, 12, and 18 mM initial concentration, respectively (Table 3-2). Release of
CO2 was not affected, while H2 accumulation was linearly decreased (P < 0.01) by 32%.

1 Acetoacetate co-eluted off our HPLC column with formate. As formate
concentration in the rumen is normally very small (Hungate et al., 1970), reasonable
disappearances could be calculated by assuming that there was no formate present. The
percentage of acetoacetate disappeared decreased linearly (P < 0.01) with the initial
concentration (Table 2-2). Total VFA concentration, and production of acetate, butyrate,
and isovalerate were linearly increased (P < 0.01) by the addition of acetoacetate.
Pr0pionate, isobutyrate, and valerate production, the substrate OM apparent
fermentability, and final pH were not affected. Methane produced per milligram of FOM
was decreased (P < 0.01) as a result of the slight decrease in CH, production and the
increase in the amount FOM due to the additive disappearance. Ammonia concentration

was lowest (P = 0.04) at 12 mM initial concentration of acetoacetate.

82

Table 3-2. Effects of the addition of acetoacetate on in vitro ruminal fermentation

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

non-useful H,
%

 

 

 

 

 

 

 

 

 

Linear Quadratic Cubic
Initial concentration, mM effect effect effect SEM
P = P = P =
0 6 12 18

CH4, umol 421 400 346 377 0.03 0.13 0.12 15.2
C02, umol 911 987 936 998 0.56 0.92 0.51 78.6
H2, umol 0.41 0.35 0.35 0.28 < 0.01 0.05 0.62 0.011
Additive - 303 571 722 < 0.01 0.02 - 15.8
disappearance,
umol
Additive - 100 93.2 78.6 < 0.01 0.09 - 1.79
disappearance, ~
%
Total VFA, 54.5 63.1 70.9 75.8 < 0.01 0.12 0.68 1.08
mM
Total VFA 1657 2117 2530 2789 < 0.01 0.12 0.68 57.3
production,
umol
Acetate, umol 1111 1542 1922 2110 < 0.01 0.03 0.51 45.7
Propionate, 345 336 330 337 0.20 0.11 0.54 4.51
pmol '
Butyrate, pmol 136 172 208 259 < 0.01 0.09 0.38 3.79
Isobutyrate, 16.9 13.3 5.75 17.4 0.77 0.13 0.28 4.51
mol
Valerate, nmol 20.4 20.2 23.7 20.0 0.70 0.20 0.08 1.24
Isovalerate, 27.8 32.3 40.5 45.1 < 0.01 0.98 0.48 2.25
mol
FOM, % 58.0 62.4 63.3 63.8 0.03 0.22 0.65 1.48
Substrate 58.0 58.3 56.6 58.8 0.93 0.55 0.41 1.53
fermentability‘,
%
CH4/FOM, 2.62 2.10 1.61 1.60 < 0.01 < 0.01 0.20 0.007
umol/mg
Final pH 6.86 6.88 6.84 . 6.91 0.69 0.56 0.43 0.046
NH4+, mg/dL 26.2 24.8 23.7 25.3 0.41 0.04 0.42 0.680
H produced, 3324 4326 5222 5817 < 0.01 0.11 0.71 80.9
umol
H incorporated, 2719 2699 2555 2782 0.88 0.09 0.13 42.6
pmol
H recovery, % 81.8 62.5 48.9 48.0 < 0.01 < 0.01 0.26 0.91
Nutritionally 38.4 40.7 46.1 45.8 < 0.01 0.31 0.13 0.45
useful H, %
Nutritionally 61.6 59.3 53.9 54.2 < 0.01 0.31 0.13 0.45

 

'Substrate apparently fermented (%) = (FOM (g) - additive disappeared (g)) x 100 /
(substrate OM (g))

83

 

Hydrogen produced was linearly (P < 0.01) increased by acetoacetate. As H was
not affected, H recovery was decreased (P < 0.01) flom 81.8 to 48.0%. The percentage
of nutritionally usefirl H incorporated was maximal (P = 0.02) at 12 mM acetoacetate

(Table 3-2).

fi-Hydroxybutyrate

Addition of B-hydroxybutyrate did not affect CH, production or H2 accumulation (Table
3-3). The release of CO2 was linearly increased (P = 0.03). Similar to acetoacetate, [3-
hydroxybutyrate co-eluted off the HPLC column with formate. As with acetoacetate, it
was assumed for calculating B-hydroxybutyrate disappearance that no formate was
present. The percentage of B-hydroxybutyrate disappeared decreased linearly (P < 0.01)
with its initial concentration. Total VFA concentration, and acetate and butyrate
production were linearly increased (P < 0.01) by the addition of fi-hydroxybutyrate. The
substrate apparent fermentability was linearly increased (P = 0.03) flom 58.0 to 63.4%.
However, as not all the additive disappeared, F OM tended (P = 0.07) to decrease flom
58.0 to 55.0%. Propionate, isobutyrate, valerate, and isovalerate production, the final
pH, and NH,+ concentration, were not affected (Table 3-3). Methane produced per
milligram of FOM was decreased (P < 0.01) as a consequence of the increase in the

amount F OM due to the additive disappearance.

84

Table 3-3. Effects of the addition of B-hydroxybutyrate on in vitro ruminal fermentation

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

non-useful H, %

 

 

 

 

 

 

 

 

Linear Quadratic Cubic
Initial concentration, mM effect effect effect SEM
P = P = P =

0 6 12 18
CH4, tunol 421 425 423 442 0.24 0.52 0.61 10.9
C02, umol 911 927 937 1041 0.03 0.22 0.52 33.0
Hz, pmol 0.41 0.39 0.50 0.43 0.28 0.36 0.05 0.029
Additive - 164 237 278 < 0.01 0.49 - 18.3
disappearance,
pmol
Additive 53.4 38.8 30.3 < 0.01 0.30 - 2.23
disappearance,
%
Total VFA, mM 54.5 59.4 61.2 65.0 < 0.01 0.59 0.31 1.01
Total VFA 1657 1917 2016 2215 < 0.01 0.65 0.39 53.3
production,
umol
Acetate, umol 1111 1296 1379 1511 < 0.01 0.47 0.35 34.1
Propionate, 345 348 337 353 0.59 0.36 0.16 5.89
punol
Butyrate, umol 136 207 242 281 < 0.01 0.17 0.44 10.6
Isobutyrate, 16.9 14.5 10.1 15.2 0.74 0.56 0.68 6.10
mol
Valerate, umol 20.4 23.1 21.1 24.1 0.29 0.94 0.24 1.76
Isovalerate, 27.8 28.2 25.9 31.1 0.51 0.35 0.37 2.44
mol
FOM, % 58.0 57.9 54.8 55.0 0.07 0.90 0.33 1.35
Substrate 58.0 58.4 58.4 63 .4 0.03 0.13 0.43 1.38
fermentability‘,
%
CH4/FOM, 2.62 2.38 2.29 2.16 < 0.01 0.23 0.41 0.003
tunol/mgf
Final pH 6.86 6.84 6.85 6.93 0.27 0.24 0.81 0.038
NH4+, mg/dL 26.2 24.9 25.4 24.8 0.51 0.79 0.58 1.17
H produced, 3324 3991 4288 4724 < 0.01 0.40 0.39 89.8
mol
H incorporated, 2719 2902 2937 3131 < 0.01 0.94 0.36 50.2
mol
H recovery, % 81.8 72.6 68.5 66.5 < 0.01 0.01 0.47 0.55
Nutritionally 38.4 41.4 42.3 43.6 < 0.01 0.10 0.31 0.35
useful H, %
Nutritionally 61.6 58.6 57.7 56.4 < 0.01 0.10 0.31 0.35

 

‘Substrate apparently fermented (%) = (FOM (g) - additive disappeared (g)) x 100 / (substrate

0M (8))

85

 

Hydrogen produced and incorporated were linearly increased (P < 0.01) by B—
hydroxybutyrate. Hydrogen recovery was decreased (P < 0.01) flom 81.8 to 66.5%. The
percentage of H incorporated into nutritionally useful products was linearly increased (P

< 0.01) by B-hydroxybutyrate flom 38.4 to 43.6% (Table 3-3).

Crotonic acid

Production of CH, was 4, 9, and 2 % lower (P < 0.05) than the control at 6,12, and
18 mM initial concentration of crotonic acid, respectively (Table 3-4). The release of
CO2 was linearly increased (P = 0.01) by 24%, and H2 accumulation was not affected.
Crotonic acid disappearance was not estimated because it co-eluted off the HPLC column
with isovalerate. Total VFA concentration (P < 0.01), and production of acetate (P <
0.01), butyrate (P < 0.01), isobutyrate (P = 0.04), and valerate (P < 0.01) were increased
by crotonic acid (Table 3-4). Propionate production and NH,+ concentration were not
affected. If control levels of isovalerate are assumed, crotonic acid increased (P < 0.05;
cubic response) the substrate apparent fermentation flom 57.0 to 68.2%. Methane
produced per milligram of FOM was decreased (P < 0.01) as a result of the decrease in
CH, production and the increase in the amount FOM due to the additive disappearance.
Final pH was linearly decreased (P < 0.01). Hydrogen produced and incorporated were
linearly increased (P < 0.01) by crotonic acid, but H recovery was linearly decreased (P <
0.01) flom 81.8 to 56.7%. The percentage of H incorporated into nutritionally useful

products was linearly increased (P < 0.01) flom 38.4 to 49.1% (data not shown).

86

Table 3-4. Effects of the addition of crotonic acid on in vitro ruminal fermentation

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

Linear Quadratic Cubic
Initial concentration, mM effect effect effect SEM
P = P = P =

0 6 12 18
CH4, umol 421 406 385 413 0.31 < 0.05 0.22 9.19
C02, mo] 91] 1013 1007 1127 0.01 0.85 0.27 44.5
Hz, umol 0.41 0.43 0.57 0.46 0.17 0.16 0.08 0.042
Additive - NA‘ NA NA NA NA NA NA
disappearance,
pmol
Additive - NA NA NA NA NA NA NA
disappearance,
%
Total VFA, 53.7 59.0 61.2 73.5 < 0.01 < 0.01 0.02 0.95
mMl
Total VFA 1629 1911 2030 2682 < 0.01 < 0.01 0.02 50.3
production,
umol'
Acetate, nmol 1111 1269 1368 1854 < 0.01 < 0.01 0.02 33.8
Propionate, 345 351 335 33 7 0.23 0.76 0.22 6.59
umol
Butyrate, umol 136 255 302 457 < 0.01 0.13 < 0.01 10.2
Isobutyrate, 16.9 1 1.2 2.22 7.39 0.04 0.14 0.27 3 .31
mol
Valerate, prnol 20.4 24.8 23.8 27.3 < 0.01 0.71 0.11 1.19
Isovalerate, 27.8 NA NA NA NA NA NA NA
umol
FOM, %2 57.0 60.0 57.6 67.2 < 0.01 0.05 0.02 1.40
Substrate 57.0 NA NA NA NA NA NA NA
fermentability’,
%
CH4/FOM, 2.62 2.24 2.03 1.74 < 0.01 0.55 0.47 0.007
mong
Final pH 6.86 6.94 6.72 6.67 < 0.01 0.19 0.04 0.045
NH4+, mg/dL 26.2 25.2 24.9 24.9 0.43 0.69 0.95 1.01

 

 

lIsovalerate not included in the calculations.

2FOM = apparently fermented OM.

3Substrate apparently fermented (%) = (FOM (g) - additive disappeared (g)) x 100 /
(substrate OM (g))

‘NA = not available. As crotonic acid co-eluted the HPLC column with isovalerate, their
final concentrations, and hence substrate fermentation, could not be determined.

87

Propynoic acid

Methane production was decreased (P < 0.01) by 65, 72, and 76%, at 6, 12, and
18 mM initial concentration, respectively (Table 3-5). The release of CO2 was linearly
decreased (P = 0.04). Propynoic acid caused (P < 0.01) a 42, 53, and 51-fold increase in
H2 accumulation, at 6, 12, and 18 mM initial concentration, respectively.

Less than 5% of propynoic acid disappeared. Disappearance was similar for all
initial concentrations of propynoic acid (Table 3-5). Total VFA concentration, acetate
production, final pH, and NH,+ concentration were linearly decreased (P < 0.01).
Propionate production was maximum at 6 mM initial concentration, and then decreased
(P < 0.01). Butyrate production increased at 6 and 12 mM concentration of propynoic
acid, and decreased at 18 mM (P = 0.04; quadratic response). Isobutyrate (P = 0.02;
cubic response) and isovalerate (P < 0.01; cubic response) production were minimum at 6
mM initial concentration. Valerate production was not affected by propynoic acid
concentration. The substrate apparent fermentability was decreased flom 62.0 to 56.6 and
57.4% at 6 and 18 mM initial concentration, respectively, but not affected at 12 mM (P <
0.01; cubic response). Although F OM (%) was decreased (P < 0.01) by pmpynoic acid,
CH, production per milligram of FOM was decreased by 63, 72, and 75%, at 6, 12, and
18 mM initial concentration, respectively.

Propynoic acid caused the accumulation of some unusual fermentation products
(data not shown). Formate was increased (P < 0.01) flom 0.48 to 5.59 mM, and ethanol
flom 0.11 to 3.17 mM, both at 12 mM propynoic acid. Also, succinate concentration was

increased (P < 0.01) flom 0.02 to 1.10 mM.

88

Table 3-5. Effects of the addition of propynoic acid on in vitro ruminal fermentation

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

Linear Quadratic Cubic
Initial concentration, mM effect effect effect SEM
P = P = P =
0 6 12 18

CH4, umol 447 155 123 108 < 0.01 < 0.01 < 0.01 5.18
C02, umol 1010 874 906 794 0.04 0.84 0.25 55.7
Hz, umol 0.93 38.6 49.5 47.5 < 0.01 < 0.01 0.01 0.81
Additive - 10.3 20.9 41.1 0.01 0.54 - 6.06
disappearance,
umol
Additive - 3.38 3.42 4.48 0.39 0.54 - 0.85
disappearance,
%
Total VFA, 56.8 52.5 51.5 49.3 < 0.01 0.13 0.14 0.62
mM
Total VFA 1784 1557 1503 1387 < 0.01 0.13 0.15 32.8
production,
urnol
Acetate, umol 1224 961 922 880 < 0.01 < 0.01 < 0.01 13.5
Propionate, 342 391 335 290 < 0.01 < 0.01 < 0.01 7.23
mol
Butyrate, umol 144 152 155 135 0.35 0.04 0.50 5.74
Isobutyrate, 18.7 8.53 25.2 22.7 0.1 1 0.31 0.02 3.53
umol
Valerate, pmol 22.5 24.2 21.5 34.1 0.30 0.42 0.52 6.49
Isovalerate, 32.4 20.8 44.2 25.6 0.85 0.33 < 0.01 3.31
mol
FOM, % 62.0 52.3 53.7 45.7 < 0.01 0.45 < 0.01 1.04
Substrate 62.0 56.6 62.5 57.4 0.16 0.95 < 0.01 1.15
fermentability‘,
%
CH4/FOM, 2.73 1.02 0.77 0.69 < 0.01 < 0.01 < 0.01 0.005
mong
Final pH 7.01 6.94 6.83 6.80 < 0.01 0.63 0.46 0.033
NH44', mg/dL 23.9 18.0 18.3 16.8 < 0.01 < 0.01 0.02 0.64
H produced, 3562 3131 3138 2912 < 0.01 0.17 0.06 48.1
mol
H incorporated, 2860 2022 2099 1914 < 0.01 < 0.01 < 0.01 37.3
mol
H recovery, % 81.2 65.0 67.6 66.0 < 0.01 < 0.01 < 0.01 0.45
Nutritionally 36.7 57.8 50.6 50.2 < 0.01 < 0.01 < 0.01 0.55
usefirl H, %
Nutritionally 62.3 42.2 49.4 49.8 < 0.01 < 0.01 < 0.01 0.55

non-useful H,
%

 

 

 

 

 

 

 

 

 

'Substrate apparently fermented (%) = (FOM (g) - additive disappeared (g)) x 100 /
(substrate OM (g))

89

 

Propynoic acid decreased (P < 0.01) H produced and incorporated. Hydrogen
recovery decreased (P < 0.01) flom 81.2 to 66.0%. The percentage of H incorporated
into nutritionally useful fermentation end products was increased (P < 0.01) flom 36.7 to

49.8% (Table 3-5).

3-Butenoic acid

Methane production tended (P = 0.07) to decrease linearly by 5%, and H2
accumulation was not affected (Table 3-6). There was a 25% linear increase (P = 0.02) in
CO2 release with the addition of 3-butenoic acid. Acetate and butyrate production were
linearly (P < 0.01) increased. Valerate and isovalerate production were maximum (P <
0.01) at 6 mM initial concentration of 3-butenoic acid. Isobutyrate tended (P = 0.07) to
increase linearly. The co-elution of 3-butenoic acid and propionate off the HPLC column
prevented us flom finding propionate production and 3-butenoic acid disappearance. If
100% disappearance of 3-butenoic acid is assumed, it would have increased (P < 0.05;
cubic response; data not shown) the substrate apparent fermentability flom 62.0 to 74.0,
68.3, and 73.7%, at 6, 12, and 18 mM initial concentration, respectively. Final pH tended

(P = 0.05) to decrease linearly. Ammonia concentration was not affected.

90

 

 

Table 3-6. Effects of the addition of 3-butenoic acid on in vitro ruminal fermentation

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

Linear Quadratic Cubic
Initial concentration, mM effect effect effect SEM
P = P = P =
0 6 12 18

CH4, umol 447 449 423 424 0.07 0.96 0.26 10.3
C02, umol 1010 1075 1046 1264 0.02 0.23 0.23 59.2
Hzfumol 0.93 0.99 0.77 3.53 0.12 0.20 0.48 0.97
Additive - NA2 NA NA NA NA NA NA
disappearance,
umol
Additive - NA NA NA NA NA NA NA
disappearance,
%
Total VFA, mM 56.8 NA NA NA NA NA NA NA
Total VFA 1784 NA NA NA NA NA NA NA
production,
umol
Acetate, umol 1224 1578 1923 2226 < 0.01 0.25 0.72 20.6
Propionate, 342 NA NA NA NA NA NA NA
pmol
Butyrate, umol 144 340 443 550 < 0.01 < 0.01 0.03 8.2
Isobutyrate, 18.7 32.9 29.1 32.3 0.07 0.21 0.20 4.01
mol
Valerate, pmol 22.5 91.0 60.7 58.3 < 0.01 0.01 < 0.01 4.63
Isovalerate, 32.4 94.4 82.9 70.2 < 0.01 < 0.01 < 0.01 4.58
mol
FOM, % 62.0 NA NA NA NA NA NA NA
Substrate 62.0 NA NA NA NA NA NA NA
fermentability‘,
%
Final pH 7.01 6.79 6.78 6.80 0.05 0.08 0.54 0.061
NH4+, mg/dL 23.9 21.4 22.6 22.0 0.20 0.22 0.12 6.97

 

 

 

 

‘Substrate apparently fermented (%) = (F OM (g) - additive disappeared (g)) x 100 /
(substrate OM (g)).

2NA = not available. As 3-butenoic acid co-eluted the HPLC with pr0pionate, their final
concentrations, and total VFA, could not be determined, and fermentation could not be
estimated.

91

 

2-Butynoic acid

Methane production was linearly decreased (P < 0.01) by 4, 6, and 9% at 6, 12,
and 18 mM initial concentration, respectively (Table 3-7). The release of CO2 and H2
accumulation, were not affected. Total VFA concentration was maximum (P < 0.01) at 6
mM propynoic acid. Acetate and propionate production were decreased (P < 0.01) at 12
and 18 mM initial concentration. Butyrate, valerate, and isovalerate production were
maximum (P < 0.01) at 6 mM initial concentration. Apparently FOM was decreased (P <
0.01) by 2-butynoic acid from 61.4 to 48.6%. Methane produced per milligram of FOM .
was unaffected. Final pH and NH,+ concentration were both linearly decreased (P < 0.01)
by 2-butynoic acid. Hydrogen produced and incorporated were highest (P < 0.01) at 6
mM 2-butynoic acid, but H recovery was not affected. The percentage of H incorporated
into nutritionally useful end products was highest (P < 0.01) at 6 mM 2-butynoic acid

(data not shown).

92

Table 3-7. Effects of the addition of 2-butynoic acid on in vitro ruminal fermentation

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

Linear Quadratic Cubic
Initial concentration, mM effect effect effect SEM
P = P = P =
0 6 12 18

CH4, umol 447 430 422 407 < 0.01 0.91 0.57 6.74
C02, umol 1010 958 1109 1074 0.17 0.88 0.13 51.4
Hz, nmol 0.93 0.75 1.08 1.32 0.15 0.34 0.52 0.21
Additive - NA‘ NA NA NA NA - NA
disappearance,
umol
Additive - NA NA NA NA NA - NA
disappearance,
%
Total VFA, 56.2 60.9 55.9 54.7 < 0.01 < 0.01 < 0.01 0.34
mM‘
Total VFA 1765 2016 1750 1686 < 0.01 < 0.01 < 0.01 18.3
production,
umol'
Acetate, umol 1224 1236 1144 1125 < 0.01 0.15 < 0.01 9.76
Propionate, 342 354 311 296 < 0.01 0.01 < 0.01 4.18
mol
Butyrate, 144 207 165 162 0.62 < 0.01 < 0.01 4.61
mol
Isobutyrate, 1 8.7 NA NA NA NA NA NA NA
umol
Valerate, 22.5 129 74.4 59.0 0.01 < 0.01 < 0.01 3.85
mol
Isovalerate, 32.4 90.0 55.6 43.9 0.99 < 0.01 < 0.01 3.70
pmol
FOM’, % 61.4 64.6 54.1 48.6 < 0.01 < 0.01 < 0.01 0.62
Substrate 61 .4 NA NA NA NA NA NA NA
fermentation’,
%
CH4/FOM, 2.73 2.65 2.85 2.55 0.58 0.41 0.20 0.019
umol/mg
Final pH 7.01 7.03 6.88 6.78 < 0.01 0.22 0.27 0.044
NH4+, mg/dL 23.9 23.6 21.3 21.6 < 0.01 0.51 0.07 4.76
‘Isobutyrate not included.

2Substrate apparently fermented (%) = (FOM (g) - additive disappeared (g)) x 100 /
(substrate OM (g))
’NA = not available. As 3-butenoic acid co-eluted the HPLC column with isobutyrate, their

final concentrations could not be determined, and hence the substrate fermentation, could not

be estimated.

93

 

Ethyl 2-butynoate

Methane production was linearly decreased (P < 0.01) by 24, 64, and 79%, at 6,
12, and 18 mM initial concentration, respectively (Table 3-8). Release of CO2 was also
linearly decreased (P < 0.01) by ethyl 2-butynoate. Ethyl 2-butynoate caused (P < 0.01) a
12, 28, and 37-fold increase in H2 accumulation, at 6, 12, and 18 mM initial
concentration, respectively. Acetate production and NH,‘ concentration were linearly
decreased (P < 0.01). Propionate, valerate, and isovalerate production were maximum at
6 mM initial concentration, and dropped at 12 and 18 mM (P < 0.01). Butyrate
production was not determined because it co-eluted off the HPLC column with ethyl 2-
butynoate. Isobutyrate production was linearly increased (P < 0.01) 7, 13, and 18-fold, at
6, 12, and 18 mM initial concentration, respectively. Apparent fermentation of OM and
substrate were not estimated as the co-elution of ethyl 2-butynoate and butyrate off the
HPLC column prevented the determination of butyrate production and ethyl 2-butynoate
disappearance. If 100% disappearance is assumed, ethyl 2-butynoate would have linearly
decreased (P < 0.01) substrate apparent fermentability flom 62.0 to 29.3% (data not
shown). Final pH was not affected.

Ethyl 2-butynoate caused the accumulation of some unusual end products of
ruminal fermentation (data not shown). Formate concentration was increased (P < 0.01)
flom 0.48 to 6.11 mM at 18 mM ethyl 2-butynoate. Also, ethanol concentration was

increased (P < 0.01) flom 0.12 to 10.4 mM at 18 mM ethyl 2-butynoate.

94

Table 3-8. Effects of the addition of ethyl 2-butynoate on in vitro ruminal fermentation

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

Linear Quadratic Cubic
Initial concentration, mM effect effect effect SEM
P = P = P =

0 6 12 18
CH4, nmol 447 340 160 93.9 < 0.01 0.63 0.33 40.3
C02, umol 1010 909 704 748 < 0.01 0.30 0.26 65.6
Hz, umol 0.93 11.1 25.8 34.4 < 0.01 0.80 0.47 3.10
Additive - NA2 NA NA NA NA NA NA
disappearance,
umol
Additive - NA NA NA NA NA NA NA
disappearance, .
%
Total VFA, 56.2 NA NA NA NA NA NA NA
mM
Total VFA 1765 NA NA NA NA NA NA NA
production,
umol
Acetate, umol 1224 1181 931 792 < 0.01 0.32 0.16 45.5
Propionate, 342 427 358 329 0.04 < 0.01 < 0.01 9.89
mol '
Butyrate, nmol 144 NA NA NA NA NA NA NA
Isobutyrate, 18.6 123 245 340 < 0.01 0.90 0.78 36.2
mol
Valerate, nmol 22.5 87.6 47.5 32.5 0.50 < 0.01 < 0.01 3.22
Isovalerate, 32.4 72.8 59.7 56.0 0.02 < 0.01 0.01 4.32
pmol
FOM, % 62.0 NA NA NA NA NA NA NA
Substrate 62.0 NA NA NA NA NA NA NA
fermentabilityl
, %
Final pH 7.01 6.97 6.91 6.87 0.17 0.93 0.92 0.071
NH4'F, mg/dL 23.9 20.0 18.3 17.0 < 0.01 0.05 0.53 0.55

 

 

'Substrate apparently fermented (%) = (F OM (g) - additive disappeared (g)) x 100 /
(substrate OM (g))

2NA = not available. As ethyl 2-butynoate co-eluted the HPLC with butyrate, their final
concentrations, and total VFA, could not be determined, and fermentation could not be
estimated.

95

Discussion
Oxaloacetate and butyrate enhancers

Acetate, followed by C02, seemed to be the major C sink of the metabolism of
added oxaloacetic acid. Therefore, most of added oxaloacetic acid was not fermented to
propionate, as was hypothesized, but perhaps decarboxylated to pyruvate, and
subsequently decarboxylated again to acetate, releasing one pair of reducing equivalents.
The increase observed in the release of CO2 suggests that oxaloacetic acid in fact
underwent decarboxylation. The slight increase in CH, production might have been a
consequence of the release of reducing equivalents in the oxidative decarboxylation of
pyruvate into acetate. Oxaloacetate, is , however, an intermediate of a ruminal
fermentation pathway leading to propionate (Russell and Wallace, 1997). It was expected
to be metabolized to propionate, rather than to acetate. It is possible that most of
externally added oxaloacetic acid was taken up by microbial species whose main
fermentation end .product is acetate, rather than propionate.

Acetate, rather than butyrate, as was hypothesized, also seem to have been the
major C sink of added acetoacetate and B—hydroxybutyrate. More of the latter, however,
seemed to be converted to butyrate. Acetoacetate could have been broken down into two
moles of acetate, which agrees with the fact that CO2 release did not increase. The
existence of a preferred pathway towards acetate could have allowed the greater
disappearance observed for acetoacetate as compared to B-hydroxybutyrate, as B-
hydroxybutyrate would need to be oxidized to acetoacetate in order to be converted to

acetate. Similar to oxaloacetic acid, it is possible that microbial species different flom the

96

butyrate producers that normally metabolize these compounds took the externally added
additives, and metabolized them to acetate.

More of the added crotonic acid was fermented to butyrate, as compared to the
other additives, but acetate was still an important C sink. Similarly, the sewage anaerobic
bacterium Syntrophomonas wolfei catabolized crotonate to acetate and smaller
proportions of butyrate and caproate (Beaty and McInemey, 1987). Thus, the added
organic acids did not seem to have been metabolized only by the pathways of which they
are intermediates in ruminal fermentation.

Oxaloacetic acid and B-hydroxybutyrate did not inhibit CH, production.
Acetoacetate caused a small decrease in CH, production, without inhibiting fermentation
or causing the accumulation of end products of fermentation without a nutritional value.

The small decrease in CH, production caused by crotonic acid could be partly due to the
decrease in pH that it caused (Van Kessel and Russell, 1996), as it was added as a flee
acid. Furthermore, the fact that the pH was not measured at the CO2 partial pressure
present in the Wheaton bottles before opening them, probably resulted in some
overestimation of the final pH, due to loss of dissolved C02, as this is in equilibrium with
H2C03 (Kohn and Dunlap, 1998). B-Hydroxybutyrate at 18 mM initial concentration

Stimflated the substrate apparent fermentability. Crotonic acid seemed to have the same
efl‘ect. Stimulation of fermentation of a high roughage substrate has been reported for
Pyruvate, lactate, fumarate, malate, 2-oxoglutarate and tartrate (Lopez et al., 1999). Due
to its low disappearance, B—hydroxybutyrate did not affect OM apparent fermentability.

B\ltyrate absorbed through the rumen and omasal walls is converted into B-

hEr'droxybutyrate, and used as an energy source (McDonald et al., 1995). Externally

97

added B-hydroxybutyrate escaped flom ruminal fermentation could be usable for the
ruminant, if it could be absorbed as such.

All of the additives decreased H recovery. The inhibition of methanogenesis may '
have stimulated some H sinks that were not measured, like sulfate and nitrate reductions
(Stewart et al., 1997), and fatty acids biohydrogenation and synthesis (Czerkawski, 1986).
Acetoacetate, B-hydroxybutyrate, and crotonic acid increased the percentage of H
incorporated into nutritionally useful sinks. B-Hydroxybutyrate did not inhibit CH,
production, so fermentation of the additive itself must not have produced additional CH,,

or produced less CH,, than the substrate fermentation.

Unsaturated organic acids and esters
A shift of the VFA profile from acetate to propionate when methanogenesis is
inhibited has been previously reported (Marty and Demeyer, 1973; Garcia-Lopez et al.,
1 996; Nagaraja et al., 1997). However, the acetate to propionate ratio decreased flom
3 - 56 to 2.44 at 6 mM propynoic acid, and then increased to 2.74 and 3.05 at 12 and 18
mM, respectively (P < 0.001; quadratic response; data not shown). Ethyl 2-butynoate
linearly decreased (P < 0.001) the acetate to propionate ratio from 3.56 at 0 mM to 2.75,
2 - 56, and 2.38, at 6, 12, and 18 mM initial concentration, respectively (data not shown).
Some unusual end products of fermentation accumulated when methanogenesis
Was inhibited by propynoic acid or ethyl 2-butynoate. Under normal conditions,
IIlethanogenesis keeps a low partial pressure of H2 in the rumen (Wolin et al., 1997).
Hydrogen accumulation has been observed with other methanogenesis inhibitors, like 2-

bromoethanesulfonate (Martin and Macy, 1985) and 9,10-anthraquinone (Garcia-Lopez et

98

 

al., 1996). Inhibition of methanogenesis using 2-bromoethanesulfonate has also caused
formate accumulation, because the increase in H2 partial pressure displaces the
equilibrium flom HCO,’ and H2 towards formate formation (Wu et al., 1993). It is also
possible that the inhibition of methanogenesis stimulated the disposal of reducing
equivalents flom pyruvate oxidative decarboxylation to acetyl-CoA into formate, a
reaction catalyzed by formate lyases (Gottschalk and Andressen, 1979) instead of
pyruvate oxidoreductases (U yeda and Rabinowitz, 1971). Reducing equivalents spared
flom methanogenesis would also have been used to reduce acetyl-CoA to ethanol, as
happens in pure cultures of Ruminococcus albus and Neocallimastixfrontalis in the
absence of methanogens (Wolin et al., 1997). The accumulation of H2, formate, and
ethanol indicates that the electrons not captured by methanogenesis were not efficiently
disposed of into other alternative pathways such as propionate formation, or fatty acids
synthesis and biohydrogenation (Czerkawski, 1986).

Succinate is a fermentation intermediate that normally does not accumulate in the
rumen or mixed ruminal cultures, as it is converted to propionate by succinate utilizers
(Wolin et al., 1997). It is interesting that the greatest accumulation of succinate occurred
at 12 and 18 mM initial concentration of propynoic acid (0.85 and 1.10 mM,
respectively), while propionate production was maximum at 6 mM. It is possible that
succinate utilizers could have been overwhelmed by the amount of succinate formed at 12
and 18 mM propynoic acid. Added succinate at 34 mM initial concentration was
metabolized to both acetate and propionate, although disappearance was not reported
(Czerkawski and Breckenridge, 1972). There might be opportunities to increase the

amounts of propionate formed by adding succinate utilizers to the fermentations, or by

99

stimulating the ones already present. Alternatively, the fact that propionate production
decreased, rather than remained constant, at 12 and 18 mM propynoic acid, suggests a
direct inhibition by propynoic acid on succinate utilizers. In support of this, it was found
that added succinate at 29.7 mM was completely consumed by a mixed ruminal culture,
and that more than 90% of it was decarboxylated to propionate (Sarnuelov et al., 1999).
The initial concentration of succinate of 29.7 mM was much greater that the accumulation
herein observed, yet succinate utilization was not overwhelmed in that study.

Despite the formation of unusual fermentation end products, propynoic acid
increased the percentage of H incorporated into products nutritionally useful for the host
animal. However, due to the decrease in H produced and to the formation of non-useful
H sinks, H spared flom methanogenesis by propynoic acid did not cause an increase in
the absolute amount of H incorporated into useful sinks. Between two thirds and four
fifths of the electrons released by fermentation were accounted by measured sinks. Little
propynoic acid was metabolized, so it could have not acted as an electron sink itself. A H
balance was not calculated for ethyl 2-butynoate because its co-elution with butyrate flom
the HPLC column prevented the determination of butyrate production.

It was hypothesized that propynoic acid and ethyl 2-butynoate would inhibit CH,
production by being alternative electron sinks. However, it is doubtful that
methanogenesis was inhibited based on electron withdrawal flom the medium since: 1)
accumulation of unusual, reduced end products like H2, formate, and ethanol, was
observed, and 2) almost all of the propynoic acid remained after 24 h incubation.

Therefore, it was not hydrogenated to propionate or acrylate.

100

Propynoic acid decreased total OM apparent fermentability partly because the
additive itself was not fermented. Apparent fermentability of the alfalfa substrate was
decreased at 6 and 18 mM, but unaffected at 12 mM initial concentration. However, a
higher proportion of the fermented products were nutritionally non-usable at 12 mM ,
compared to 6 mM initial concentration. As most of the inhibition of methanogenesis
was already achieved at 6 mM initial concentration, the utilization of lower initial
concentrations could be a way of minimizing the negative effects of propynoic acid on
fermentation. This would decrease the proportion of the OM that is not fermented.

‘ The inhibition of methanogenesis caused by 3-butenoic acid was small, but it
might have stimulated substrate fermentation. Acetate, followed by butyrate, seemed to
be the most important C sink for this additive.

2-Butynoic acid also caused small decreases in CH, production. Fermentation
was inhibited at 12 and 18 mM initial concentration, but not at 6 mM. At 6 mM initial
concentration, most 2-butynoic acid seemed to be metabolized to butyrate, valerate, and
isovalerate. Disappearance of 2-butynoic acid could not be measured as it co-eluted from
the HPLC column with isobutyrate; however, as changes in total VFA production were
relatively small at 12 and 18 mM compared to the control, it is possible that most of 2-

butynoic acid was not metabolized at those initial concentrations.

Implications
Propynoic acid and ethyl 2-butynoate decreased ruminal methanogenesis in vitro.
Propynoic acid had some adverse effects on the substrate apparent fermentability, and

ethyl 2-butynoate also seemed to be inhibitory to fermentation, although its disappearance

101

could not be measured. Both propynoic acid and ethyl 2-butynoate caused the formation
of products without nutritional value. It is possible that organic acids that seemed to
benefit fermentation, like B—hydroxybutyrate, crotonic acid, or 3-butenoic acid, could be
fed to ruminants together with propynoic acid or ethyl 2-butynoate to relieve the negative
effects on fermentation caused by the inhibitors of methanogenesis. Propynoic acid oral
LD,o to rodents is 100 mg/kg (CCOHS), although its toxicity to ruminants at the doses
inhibitory to ruminal methanogenesis would need to be evaluated. We are not aware of
toxicity trials with ethyl 2-butynoate or 3-butenoic acid. Crotonic acid LDso to rodents is
between 1 and 4.8 g/kg (CCOHS). It might be less toxic to ruminants as it is a naturally
occurring intermediate in ruminal fermentation (Miller and Jenesel, 1979). Likewise,
acetoacetate and B-hydroxybutyrate may be mildly toxic to ruminants for the same
reason. Accurate measurements of the disappearances of some of the compounds studied
are needed to fully understand what happened to, and as a consequence of, the addition of
these chemicals. Their toxicity to ruminants, as well as the potential hazards for humans

and the environment, would also need to be assessed.

102

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Callaway, T. R. and S. A. Martin. 1996. Effects of organic acid and monensin treatment

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CCOHS. 2003. Registry of Toxic Effects of Chemical Substances No. 2003. Canadian
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Chaney, A. L. and E. P. Marbach. 1962. Modified reagents for determination of urea and
ammonia. Clin. Chem. 8: 130-132.

Czerkawski, J. W. 1986. An Introduction to Rumen Studies. Pergamon Press, Oxford,
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Czerkawski, J. W. and G. Breckenridge. 1972. Fermentation of various glycolytic
intermediates and other compounds by rumen micro-organisms, with particular
reference to methane production. Br. J. Nutr. 27: 131-146.

Garcia-Lopez, P. M., L. Kung, Jr., and J. M. Odom. 1996. In vitro inhibition of microbial
methane production by 9,10-anthraquinone. J. Anim. Sci. 74: 2276-2284.

Goering, H. K. and P. N. Van Soest. 1975. Forage Fiber Analyses (Apparatus, Reagents,
Procedures and some Applications). 379, ARS-USDA, Washington DC.

Gottschalk, G. and J. R. Andressen. 1979. Energy metabolism in anaerobes. In: J. R.
Quayle (ed.) Microbial Biochemistry. p 85-115. United Park Press, Baltimore,
MD.

Hungate, R E., W. Smith, and T. Bauchop. 1970. Formate as an intermediate in the
bovine rumen fermentation. J. Bacteriol. 102: 389-397.

Kohn, R. A. and T. F. Dunlap. 1998. Calculating the buffering capacity of bicarbonate in
the rumen and in vitro. J. Anim. Sci. 76: 1702-1709.

Lopez, 5., C. J. Newbold, O. Bochi-Brum, A. R. Moss, and R. J. Wallace. 1999.
Propionate precursors and other metabolic intermediates as possible alternative

electron acceptors to methanogenesis in ruminal fermentation in vitro. S. Afl. J.
Anim. Sci. 29: 106-107.

103

Martin, S. A. and J. M. Macy. 1985. Effects of monensin, pyromellitic diimide and 2-
bromoethanesulfonic acid on rumen fermentation in vitro. J. Anim. Sci. 60: 544-
550.

Martin, S. A. and M. N. Streeter. 1995. Effect of malate on in vitro mixed ruminal
microorganism fermentation. J. Anim. Sci. 73: 2141-2145.

Marty, R. J. and D. I. Demeyer. 1973. The effect of inhibitors of methane production on
fermentation pattern and stoichiometry in vitro using rumen contents flom sheep
given molasses. Br. J. Nutr. 30: 369-376.

McDonald, P., R. A. Edwards, J. F. D. Greenhalgh, and C. A. Morgan. 1995. Animal
Nutrition. 5th ed. Longman Singapore Publishers, Singapore.

Miller, T. L. and S. E. Jenesel. 1979. Enzymology of butyrate formation by Butyrivibrio
- fibrisolvens. J. Bacteriol. 138: 99-104.

Moss, A. R. 1993. Methane. Global Warming and Production by Animals. lst ed.
Chalcombe Publications, Kingston, Kent, UK.

Nagaraja, T. G., C. J. Newbold, C. J. Van Nevel, and D. I. Demeyer. 1997. Manipulation
of ruminal fermentation. In: P. N. Hobson and C. S. Stewart (eds) The Rurnen
Microbial Ecosystem. p 523-632. Blackie Academic and Professional, London.

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Russell, J. B. and R. J. Wallace. 1997. Energy-yielding and energy-consuming reactions.
In: P. N. Hobson and C. S. Stewart (eds) The Rurnen Microbial Ecosystem. p
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Anaerobiospirillum succinoproducens for production of a succinate-based animal
feed additive. Appl. Environ. Microbiol. 65: 2260-2263.

Schulman, M. D. and D. Valentino. 1976. Factors influencing rumen fermentation: effect
of hydrogen on formation of propionate. J. Dairy Sci. 59: 1444-1451.

Stewart, C. S., H. J. Flint, and M. P. Bryant. 1997. The rumen bacteria. In: P. N. Hobson
and C. S. Stewart (eds) The Rurnen Microbial Ecosystem. p 10-72. Blackie
Academic and Professional, London.

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Purification and properties of the enzyme. J. Biol. Chem. 246: 3111-3119.

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Van Kessel, J. S. and J. B. Russell. 1996. The effect of pH on ruminal methanogenesis.
FEMS Microbiol. Ecol. 20: 205-210.

Voet, D. and J. G. Voet. 1995. Biochemistry. 2nd ed. John Wiley and Sons, Inc., New
York.

Wolin, M. J ., T. L. Miller, and C. S. Stewart. 1997. Microbe-microbe interactions. In: P.
N. Hobson and C. S. Stewart (eds) The Rumen Microbial Ecosystem. p 467-491.
Blackie Academic and Professional, London.

Wu, W.-M., R. E. Hickey, M. K. Jain, and J. G. Zeikus. 1993. Energetics and regulations

of formate and hydrogen metabolism by Methanobacteriumformicicum. Arch.
Microbiol. 159: 57-65.

105

CHAPTER 4

Some miscellaneous inhibitors of ruminal methanogenesis in vitro

Abstract

Three inhibitors of methanogenesis that had not been previously tested in mixed
ruminal fermentation were examined in vitro. The eukaryotic and archaeal DNA-
polymerase inhibitor aphidicolin did not affect methanogenesis or most of the
fermentation parameters. Interestingly, dimethylsulfoxide (DMSO), the carrier for
aphidicolin delivery, caused a strong decrease in ammonia concentrations. Antiprotozoal
properties of DMSO are suspected. The pterin, lumazine, decreased methanogenesis by
about 50%. Fermentation was also inhibited. Surprisingly, molecular hydrogen
accumulation was not observed and acetate molar proportion increased. The methyl-
CoM analog 2-bromopropanesulfonate decreased fermentation and butyrate molar
percentage and increased acetate molar percentage without affecting methane production.
It is difficult to understand how a structural analog of a coenzyme that is unique to

methanogens could have affected fermentation without altering methanogenesis.

Introduction
Methane formation in ruminants represents an energy loss and also makes some
contribution to global warming. Hence, there is an interest in decreasing methane

emissions by ruminant animals (Moss, 1993).

106

Aphidicolin is a specific inhibitor of DNA-polymerases types or, 6 and e in
eukaryotes (V oet and Voet, 1995). It inhibited DNA replication in crude extracts of
methanogens but not of Escherichia coli, and completely prevented the growth of several
methanogens studied (Zabel et al., 1985). The effects of aphidicolin on ruminal
fermentation have not been examined.

The pterin compound lumazine [2, 4-(1H, 3H)-pteridinedione] has been shown to
inhibit the growth of several methanogens, but had little effect on other Archaea, bacteria
or yeasts (Nagar-Anthal et al., 1996). Lumazine is a structural analog of some cofactors
involved in methanogenesis, like methanopterin and deazaflavin F,20 (White and Zhou,
1993). Its effects upon ruminal fermentation have not been studied.

2-Bromoethanesulfonate (BBS) is a chemical analogue of coenzyme M (CoM), a
cofactor involved in the last step of methanogenesis (McAllister et al., 1996). It inhibits
methyl-COM reductase as a competitive inhibitor with CoM (Smith, 1983), being a very
specific and potent methanogenesis inhibitor. 3-Bromopropanesulfonate (BPS), however,
was found to be a more potent inhibitor in the in vitro reaction, achieving 50% inhibition
at a concentration 80-fold lower (Ellerman et al., 1989). To our knowledge, the effects of
BPS on methanogenesis have not been studied with either pure or mixed microbial
cultures.

The objectives of this work were to study the effects of aphidicolin, lumazine and
BPS on ruminal fermentation of mixed ruminal cultures. It was hypothesized that these

compounds would decrease CH, production.

107

 

Materials and Methods
Incubations

Ruminal fluid was withdrawn flom two mature Holstein cows fed alfalfa hay,
mixed and strained through four layers of cheesecloth. It was then blended for 15 s and
strained through one layer of cheesecloth. One part of ruminal fluid was mixed with four
parts of buffer (Goering and Van Soest, 1975), and 15 mL of the mixture anaerobically
delivered into 25 mL Hungate tubes. All the tubes contained 0.2 g of finely ground
alfalfa hay (11.4% CP DM basis) as substrate. Samples of the ruminal fluid/buffer
mixture were taken and flozen to measure VFA initial concentrations. Tubes were sealed

under an Oz-flee CO2 atmosphere and incubated in a shaking water bath at 39 °C for 24 h.

Treatments

The experimental treatments were: aphidicolin 0, 30, 60 and 150 uM, lumazine 0,
300, 600 and 1,200 uM, and BPS and BES (positive control) 0, 1, 10 and 50 uM. Final
concentrations of aphidicolin were achieved by adding 46.5, 93 or 233 uL, respectively,
of a 3.4 mg/uL solution of aphidicolin in dimethyl sulfoxide (DMSO; (Zabel et al., 1985)
to Hungate tubes. The aphidicolin control received 233 11L of DMSO. Lumazine final
concentrations were achieved by adding 0.8, 1.5 and 3.1 mg of lumazine to Hungate
tubes. One hundred microliters of 360 ppm BPS and BBS solutions were added to
Hungate tubes to achieve a final concentration of 1 11M. Final BPS and BES
concentrations of 10 and 50 nM were achieved by adding 0.10 and 0.50 mL, respectively,

of 3600 ppm BPS and BES solutions to Hungate tubes. All tubes with 0.10 mL solutions

108

 

 

 

also received 0.40 mL of deionized water to equalize volumes. Control tubes for

lumazine, BPS and BES treatments had 0.5 mL of deionized water added.

Analysis

At the end of the incubation, gas exceeding 1 atrn was measured (Callaway and
Martin, 1996) and the fermentation was stopped by adding 0.25 mL of a 10% phenol
solution. A 0.1 mL gas sample was taken flom each tube and analyzed for gas
composition (CH,, C02, and H2) by gas chromatography (Gow Mac Gas Chromatograph
series 750, Gow Mac Instruments Co., Bridgewater, NJ; carrier gas was N2 at 820 KPa; 4'
x 1/ " DC 200 column, 5.5., operated at 150 °C; Gow Mac 069-50 Ruthenium Methanizer
for CO2 analysis; flame ionization detector for CH, and CO2 analyses and RGD2
Reduction Gas Detector, Trace Analytical, Menlo Park, CA; for H2 analysis; 3390 A
Integrator, Hewlett Packard, Avondale, CA). A 5 mL aliquot was taken and centrifuged
at 26,000 x g for 30 min. The pellet was discarded and the supernatant used for measuring
pH (Digital Benchtop pH Meter, Cole-Parmer Instrument Company, Vernon Hills, IL)
and VFA. VFA, lactate, formate and ethanol were analyzed by HPLC using a Waters 712
Wisp (Waters Associates Inc., Milford, MA), and separation was performed with a
BioRad HPX 87H column (BioRad Laboratories, Hercules, CA). Detection was done by
differential reflactometry (Waters 410, Waters Associates Inc., Milford, MA). Ammonia

was analyzed as described before (Chaney and Marbach, 1962).

109

 

Calculations

Apparently fermented organic matter (F OM) was estimated flom the VFA
stoichiometry (Demeyer and Van Nevel, 1979), but replacing caproate with isobutyrate:
FOM (mg hexose) = (Ac/2 + Pro/2 + Isobut + But +Val + Isoval) x 162, where Ac is
acetate, Pro is propionate, Isobut is isobutyrate, But is butyrate, Val is valerate and Isoval

is isovalerate, all expressed in mmoles.

Statistical analysis

1 Five replicates per compound and concentration were used. Dependent variables
were modeled as: observation = control + concentration + concentration2 + concentration3
+ residual. Models with non-significant (P > 0.05) highest order coefficients were
considered non-significant. A comparison between water and DMSO (aphidicolin)

controls was done by the Scheffe test (Neter et al., 1996).

Results and Discussion
Aphidicolin

Aphidicolin addition did not affect methanogenesis, CO2 release or H2
accumulation (Table 4-1). Final pH, FOM, CH,/F OM, total VFA concentration, acetate,
propionate, and isovalerate production were similar. There were linear tendencies
towards decreases in butyrate (P = 0.07) and isobutyrate (P = 0.06) production by
maximums of 26 and 46% reductions, respectively. Valerate production peaked at 60 uM
aphidicolin (P < 0.01; quadratic response). Aphidicolin at 60 [AM was shown to

completely inhibit the growth of five methanogens, and partially inhibited

110

 

 

Methanospirillum hungatei and Methanosarcina barkeri (Zabel et al., 1985). The growth
of Methanobacterium formicicum was not affected at 60 uM, but it was completely
inhibited at 210 uM. It is possible that the maximum concentration used in the present
experiment, 150 uM, was insufficient to inhibit ruminal methanogens if they are
intrinsically more resistant. The different sensitivities among methanogens could be
caused by different cell envelopes (Zabel et al., 1985). These investigators found,
however, that crude preparations of DNA-polymerases of M. formicicum and
Methanobacterium wolfei were resistant to aphidicolin at 30 uM, whereas the activities of
DNA-polymerases of Methanococcus vannielii were inhibited by 73%. M formicicum
DNA-polymerase activity was inhibited by about 20% at 150 uM of aphidicolin, which is
the maximum concentration used in this experiment. Klirnczak et al. (1986) found that
DNA-polymerase of Methanobacterium thermoautotrophicum was insensitive to
aphidicolin. Methanogens DNA-polymerases differ, therefore, in their sensitivity to
aphidicolin. It is also possible that other ruminal microorganisms or substances present

in ruminal fluid caused aphidicolin destruction and prevented inhibition of methanogens.

lll

Table 4-1. Effects of aphidicolin on ruminal fermentation in vitro

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

Aphidicolin (uM Linear Quadratic Cubic SEM3
regression regression regression

0 30 60 120 P = P =‘ P =2
CH4, mm] 52.1 35.9 54.9 58.1 0.48 0.65 0.30 28.1
Hz, umol 0.06 0.12 0.07 0.15 0.16 0.76 0.24 0.08
C02, umol 712 410 600 875 0.18 0.13 0.30 304
Total VFA, 70.1 69.2 68.9 66.1 0.20 0.83 0/ 89 4.94
mM
Acetate, 556 530 533 557 0.75 0.16 0.71 35.9
mol
Propionate, 209 199 190 189 0.34 0.61 0.91 32.4
umol
Butyrate, 97.3 106 104 71.9 0.07 0.12 0.99 24.4
umol
Isobutyrate, 37.2 44.6 33.4 20.2 0.06 0.37 0.43 16.7
mol
Valerate, 5.21 7.73 10.4 1.02 0.15 < 0.01 0.46 4.19
mol
Isovalerate, 8.31 10.4 21.2 7.3 0.98 0.19 0.41 15.4
mol
FOM, % 42.2 41.4 40.9 39.2 0.18 0.94 0.92 3.52
CH4/FOM, 0.68 0.47 0.73 0.81 0.35 0.61 0.31 0.38
urnol/mg
Final pH 7.05 7.02 7.03 6.88 0.18 0.60 0.79 0.19

 

 

 

 

 

 

 

 

 

 

‘Significance of the quadratic term.

2Significance of the cubic term.

3Standard error of the mean of the most significant model. If more than one model had a
P < 0.01, the highest order model was used.

112

Surprisingly, NH,+ concentration was elevated by the addition of 30 11M
aphidicolin, and to a lesser extent with 60 11M, but returned to control levels at 120 uM of
aphidicolin (Table 4-2). Moreover, NH,‘ concentration was substantially lower (P <
0.01; Scheffe analysis) in the DMSO control than in the water control (3.6 vs 17.5 mg/dL,
respectively). It is then quite clear that this strong decrease in NH,+ was not caused by
aphidicolin but by its vehicle, DMSO, as the DMSO controls received the same amount
of DMSO as the 120 uM treatment.

Although part of the drop in NH,+ concentrations could be explained by a trend (P
= 0.18) towards less fermentation, the decline was too pronounced to be accounted for
solely by this. It is possible that DMSO had antiprotozoal properties, as defaunation
decreases ruminal NH; concentration (Williams and Coleman, 1997). Alternatively, it
might have inhibited proteolytic bacteria. A reduction in protein degradation and
intraruminal N recylcling can be beneficial as it can result in a greater flow of amino
acids to the small intestine (Nagaraja et al., 1997). Further research is needed into the

mechanisms by which DMSO lowered ruminal NH,+ concentrations.

113

Table 4-2. Effects of aphidicolin and DMSO on ammonia levels and FOM

 

 

 

 

 

 

 

Treatment NH4+ (mg/ 100 mL) F OM (%)
Water control (no DMSO) 17.5 43.5
Aphidicolin control (233 uL DMSO) 3.36 42.2
Aphidicolin 30 uM (46.5 uL DMSO) 9.45 41.4
Aphidicolin 60 uM (93 [LL DMSO) 5.92 40.9
Aphidicolin 120 uM (233 uL DMSO) 2.97 39.2
SEM 1.051:2 3.511:3

 

 

 

 

 

‘Regressions do not include the water controls.
‘ 2Cubic regression (P < 0.01).
3Linear regression (P = 0.18).

Lumazine

Addition of lumazine at 600 and 1,200 1.1M decreased (P < 0.01) CH, formation
by 50% of the control (Figure 4-1). Dihydrogen accumulation was not affected. Carbon
dioxide release decreased in a quadratic (P < 0.05) fashion. There was little benefit of
increasing lumazine beyond 600 uM. Lumazine at 450 or 600 pM strongly inhibited
growth of the four methanogens studied, which included non-ruminal strains of
Methanobacteriumformicicum and Methanosarcina barkeri (Nagar—Anthal et al., 1996).
Apparently fermented OM was decreased (P < 0.01; quadratic response) by 12, 17 and
17% by lumazine at 300, 600 and 1,200 uM, respectively (Table 4-3). A decline in
fermentation with methanogenesis inhibitors has previously been observed (Prins, 1965;
Chalupa et al., 1980). and attributed to an interference with the interspecies H transfer,

which is necessary for the reoxidation of cofactors (Nagaraja et al., 1997). The final pH

114

which is necessary for the reoxidation of cofactors (Nagaraja et al., 1997). The final pH
and NH,+ concentration were not affected by lumazine addition. Methane produced per

milligram of FOM tended (P = 0.05) to decrease with lumazine addition.

 

 

 

 

 

 

160 + - - - - 1200
140 . 9 °
. l 1000
a? 120.
2
o A
E 100 l ' 800 8
O .—
b ' g
E 80 .
a: ‘600 .8
I 60 » E.
'O
5 40 \D ————— O ‘400 g "0., H2
‘I’ ' I! a SEM = 0.032
0 20r . . zoo \u. CH4
. 5 . SEM = 5.8
0 l o G - - fa”, 0 ‘k 002
o 300 600 900 1200 SEM = 50.4

Lumazine (microM)

Figure 4-1 Effects of lumazine on gases production in ruminal fermentation

Total VFA concentration was decreased (P < 0.01; quadratic response; Table 4-3)
by lumazine by a maximum of 12% at 600 uM. Acetate and valerate productions were
not affected. Propionate production tended (P = 0.07) to linearly decline by a maximum
of 11%. Butyrate production fell (P < 0.01; quadratic response) by 60%. Isobutyrate
production dropped (P < 0.01; quadratic response) by more than 10-fold at 600 uM
lumazine, although it recovered to control levels at 1,200 uM. Isovalerate production

was linearly decreased (P = 0.02) by a maximum of 29%.

115

Table 4-3. Effects of the addition of lumazine on in vitro ruminal fermentation

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

Linear Quadratic Cubic
Initial concentration, uM regression regression regression SEM
P= P=‘ P=2

0 300 600 1,200
TotalVFA, 71.1 65.9 62.9 65.2 0.02 <0.01 0.88 2.59
mM
Acetate, 527 549 538 540 0.67 0.45 0.35 27.7
urnol
Propionate, 197 193 177 176 0.07 0.53 0.48 9.17
mol
Butyrate, 134 66.9 53.0 53.5 <0.01 <0.01 0.07 15.4
mol
Isobutyrate, 45.0 10.3 4.21 42.2 0.75 <0.01 0.51 11.0
mol
Valerate, 13.4 12.8 12.5 12.5 0.25 0.50 >0.99 1.22
mol
Isovalerate, 11.6 9.78 8.54 8.27 0.02 0.20 0.96 2.07
umol
Acetate to 2.74 2.84 3.05 3.07 0.03 0.37 0.53 0.24
propionate
FOM,% 43.5 38.4 36.1 36.1 <0.01 <0.01 0.70 1.57
CH4/FOM, 1.36 1.36 0.81 0.84 0.05 0.95 0.28 0.063
umol/mg
Final pH 6.92 6.96 7.06 6.99 0.28 0.14 0.34 0.10
NH4+, 17.5 17.4 17.6 16.8 0.24 0.46 0.62 9.91
mg/dL

 

 

 

 

‘Significance of the quadratic term.

2Significance of the cubic term.

‘Standard error of the mean of the most significant model. If more than one model had a
P < 0.01, the highest order model was used.

It is unusual that the acetate to propionate ratio increased (P = 0.03; Table 4-3), as
the inhibition of methanogenesis consistently shifts the VFA profile flom acetate to
propionate (Nagaraja et al., 1997). The inhibition of methanogenesis was accompanied
by a decrease in CO2 release and H2 accumulation did not increase. It is possible that
some of the electrons spared flom methanogenesis were redirected towards acetate

synthesis flom CO2 and H2, thus compensating for the shift of fermentation flom acetate

116

to propionate. Acetogens synthesize acetate using the same precursors as methanogens,
but they do not compete well with methanogens under normal ruminal conditions due to
their higher thermodynamic threshold for H2 (Nagaraja et al., 1997). Nevertheless, it was
shown that when methanogenesis was inhibited, reductive acetogenesis increased when
the acetogen Peptostreptococcus productus was added to the fermentation medium
(Nollet et al., 1997). It is tempting to think that lumazine could somehow have

stimulated ruminal acetogens.

BPS I

BPS did not affect CH, production (Table 4-4), whilst BES decreased (P < 0.01) it
by 50 and 48% at 10 and 50 uM, respectively (Table 4-5). BES effects on
methanogenesis were in agreement with previous observations (Martin and Macy, 1985;
Nollet et al., 1997). BPS was, however, expected to be a more potent inhibitor according
to previous observations with a pure preparation of methyl-COM reductase flom
Methanobacterium thermoautotrophicum (Ellerman et al., 1989). BPS greater
effectiveness as an inhibitor of methyl-CoM reductase is due to a more similar structure
to methyl-COM as compared to BES (Ellerman et al., 1989). This should not change
even if methyl-COM reductase of ruminal methanogens is different flom the one of M.
thermoautotrophicum. There is, therefore, no clear explanation on the lack of

effectiveness of BPS in decreasing ruminal methanogenesis.

ll7

Table 4-4. Effects of the addition of BPS on in vitro ruminal fermentation

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

Linear Quadratic Cubic
Initial concentration, uM regression regression regression SEM3
P = =‘ P =2
0 1 10 50
CH4, umol 109 75.0 89.6 103 0.21 0.61 0.25 30.5
Hz, umol 0.29 0.24 0.33 0.33 0.59 0.56 0.66 0.19
C02, umol 746 560 694 682 0.70 0.56 0.78 294
Total VFA, 71.1 63.8 65.7 64.4 0.15 0.34 < 0.01 2.52
mM
Acetate, 527 539 538 534 0.97 0.56 0.43 21.2
mol
Propionate, 197 174 182 177 0.51 0.73 0.14 23.5
mol
Butyrate, 134 65.9 78.5 72.8 0.13 0.16 < 0.01 30.0
mol
Isobutyrate, 45.0 6.75 18.2 1 1.5 0.15 0.37 < 0.01 10.4
umol
Valerate, 13.4 13.6 13.1 13.1 0.47 0.58 0.67 0.97
mol
Isovalerate, 1 1.6 9.51 9.02 10.3 0.99 0.24 0.27 2.67
umol
FOM, % 43.5 37.2 38.6 37.7 0.11 0.24 < 0.01 1.57
CH4/FOM, 1.36 1.10 1.26 1.49 0.29 0.85 0.37
umol/mg
Final pH 6.92 7.01 6.86 6.95 0.97 0.23 0.16 0.11
NH47, 17.5 18.2 17.8 17.9 0.83 0.97 0.27 0.88
mg/dL

 

 

 

 

 

 

 

 

 

 

 

‘Significance of the quadratic term.

2Significance of the cubic term.

3Standard error of the mean of the most significant model. If more than one model had a P < 0.01,
the highest order model was used.

BPS did not affect H2 or CO2 release, acetate, propionate, valerate, or isovalerate
production, CH, production per milligram of FOM, final pH, or NH,+ concentration. BPS
decreased (P < 0.01) FOM flom 43.5 to 37.7%. Total VFA concentration, butyrate and
isobutyrate production were decreased (P < 0.01). It is difficult to understand how BPS

could have altered decreased butyrate and isobutyrate production. A major part of

118

butyrate production is carried out be Butyribivrio fibrisolvens (Stewart et al., 1997) and
protozoa (Huhtanen, 1992; Jaakkola and Huhtanen, 1993). Isobutyrate is a product of
valine fermentation (V oet and Voet, 1995). As CoM is a cofactor unique to methanogens
(Balch and Wolfe, 1979), methyl-COM analogs, like BPS, should not affect other ruminal

microorganisms.

Table 4-5. Effects of the addition of BES on in vitro ruminal fermentation

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

Linear Quadratic Cubic
Initial concentration, uM regression regression regression SEM3
P = P =‘ P =1
0 1 10 50
CH4, umol 109 1 19 54.0 56.4 0.01 < 0.01 < 0.01 21.7
Hz, umol 0.29 0.35 0.15 0.26 < 0.01 < 0.01 0.63 1.95
C02, umol 746 562 403 456 0.13 0.22 0.22 255
Total VFA, 71.1 65.9 62.9 65.2 0.27 0.16 < 0.01 2.29
mM
Acetate, 527 549 538 540 0.01 < 0.01 0.88 18.2
mol
Propionate, 197 193 177 176 < 0.01 0.02 0.81 23.9
mol
Butyrate, 134 66.9 53.0 53.5 0.14 0.16 < 0.01 15.0
mol
Isobutyrate, 45.0 10.3 4.21 42.2 0.10 0.06 < 0.01 8.85
umol
Valerate, 13.4 12.8 12.5 12.5 < 0.01 < 0.01 0.24 1.38
umol
Isovalerate, 1 1.6 9.78 8.54 8.27 0.43 0.81 0.30 2.58
umol
FOM, % 43.5 38.5 36.1 36.1 0.20 0.14 < 0.01 1.42
CH4/FOM, 1.36 1.36 0.81 0.84 0.01 < 0.01 0.02 0.29
mong
Final pH 6.92 6.96 7.06 6.99 0.67 0.27 0.94 0.14
NH4“', 17.5 17.4 17.6 16.8 0.12 0.37 0.79 12.7
mg/dL

 

‘Significance of the quadratic term.
‘Significance of the cubic term.

3Standard error of the mean of the most significant model. If more than one model had a P < 0.01,

the highest order model was used.

119

 

Conclusions

Aphidicolin and 2-bromopropanesulfonate did not decrease ruminal
methanogenesis in vitro. Aphidicolin’s vehicle DMSO, lowered NH; concentration by
more than 80%, and could act as an antiprotozoal agent. More research is needed to
establish how DMSO lowered NH; concentration. Lumazine decreased CH, production
by about 50%, although fermentation was decreased. Unexpectedly, the acetate to
propionate ratio increased. The effects of lumazine on pure cultures of ruminal

methanogens and acetogens need to be evaluated.

120

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on in vitro mixed ruminal microorganism fermentation of cracked corn. J. Anim.
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Chalupa, W., W. Corbett, and J. R. Brethour. 1980. Effects of monensin and amicloral on
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Chaney, A. L. and E. P. Marbach. 1962. Modified reagents for determination of urea and
ammonia. Clin. Chem. 8: 130—132.

Demeyer, D. I. and C. J. Van Nevel. 1979. Effect of defaunation on the metabolism of
rumen micro-organisms. Br. J. Nutr. 42: 5 15-524.

Ellerrnan, J ., S. Rospert, R. K. Thauer, M. Bokranz, A. Klein, M. Voges, and A.
Berkessel. 1989. Methyl-coenzyme-M reductase flom Methanobacterium
thermoautotrophicum (strain Marburg). Purity, activity and novel inhibitors. Eur.
J. Biochem. 184: 63-68.

Goering, H. K. and P. N. Van Soest. 1975. Forage Fiber Analyses (Apparatus, Reagents,
Procedures and some Applications). 379, ARS-USDA, Washington DC.

Huhtanen, P. 1992. The effects of barley vs. barley fiber with or without distiller's
solubles on site and extent of nutrient digestion in cattle fed grass-silage-based
diets. Anim. Feed Sci. Tech. 36: 319-337.

Jaakkola, S. and P. Huhtanen. 1993. The effects of forage preservation method and

proportion of concentrate on nitrogen digestion and ruminal fermentation in cattle.
Grass Forage Sci. 48: 146-154.

Klimczak, L. J ., F. Grummt, and K. J. Burger. 1986. Purification and characterization of
DNA polymerase flom the archaebacterium Methanobacterium
thermoautotrophicum. Biochemistry 25: 4850-4855.

Martin, S. A. and J. M. Macy. 1985. Effects of monensin, pyromellitic diimide and 2-

bromoethanesulfonic acid on rumen fermentation in vitro. J. Anim. Sci. 60: 544-
550.

121

McAllister, T. A., E. K. Okine, G. W. Mathison, and K.-J. Cheng. 1996. Dietary,
environmental and microbiological aspects of methane production in ruminants.
Can. J. Anim. Sci. 76: 231-243.

Moss, A. R. 1993. Methane. Global Warming and Production by Animals. lst ed.
Chalcombe Publications, Kingston, Kent, UK.

Nagaraja, T. G., C. J. Newbold, C. J. Van Nevel, and D. I. Demeyer. 1997. Manipulation
of ruminal fermentation. In: P. N. Hobson and C. S. Stewart (eds) The Rurnen
Microbial Ecosystem. p 523-632. Blackie Academic and Professional, London.

Nagar-Anthal, K. .R., V. E. Worrell, R. Teal, and D. P. Nagle. 1996. The pterin lumazine
inhibits growth of methanogens and methane formation. Arch. Microbiol. 166:
136-140.

Neter, J ., M. H. Kutner, C. J. Nachtsheim, and W. Wasserman. 1996. Applied Linear
Statistical Models. 4th ed. McGraw-Hill, Boston.

Nollet, L., D. Demeyer, and W. Verstraete. 1997. Effect of 2-bromoethanesulfonic acid
and Peptostreptococcus productus ATCC 35244 addition on stimulation of
reductive acetogenesis in the ruminal ecosystem by selective inhibition of
methanogenesis. Appl. Environ. Microbiol. 63: 194-200.

Prins, R. A. 1965. Action of chloral hydrate on rumen microorganisms in vitro. J. Dairy
Sci. 48: 991-993.

Smith, M. R. 1983. Reversal of 2-bromoethanesulfonate inhibition of methanogenesis in
Methanosarcina sp. J. Bacteriol. 156: 516-523.

Stewart, C. S., H. J. Flint, and M. P. Bryant. 1997. The rumen bacteria. In: P. N. Hobson
and C. S. Stewart (eds) The Rurnen Microbial Ecosystem. p 10-72. Blackie
Academic and Professional, London.

Voet, D. and J. G. Voet. 1995. Biochemistry. 2nd ed. John Wiley and Sons, Inc., New
York.

White, R. H. and D. Zhou. 1993. Biosynthesis of the coenzymes in methanogens. In: J. G.
Ferry (ed.) Methanogenesis. Ecology, Physiology, Biochemistry & Genetics. p
409-444. Chapman and Hall, New York / London.

Williams, A. G. and G. S. Coleman. 1997. The rumen protozoa. In: P. N. Hobson and C.

S. Stewart (eds) The Rurnen Microbial Ecosystem. p 73-139. Blackie Academic
and Professional, London.

122

Zabel, H., H. Fischer, E. Holler, and J. Winter. 1985. In vivo and in vitro evidence for
eukaryotic alfa-type DNA-polymerase of Methanococcus vannielii. Syst. Appl.
Microbiol. 6: 111-118.

123

CHAPTER 5

Efl'ects of two lipids on in vitro ruminal methane production

Abstract

As CH4 emissions by ruminants are a loss of energy and also contribute to global
warming, there is an interest in decreasing rumen methanogenesis. Fats and oils usually
decrease CH4 production both in vitro and in vivo, although they can inhibit
fermentation. The effects of olive oil and a hexadecatrienoic acid (cis-C15;6,9, 12) on
mixed ruminal cultures were studied in 24 h-batch fermentation. The hexadecatrienoic
acid was extracted flom the Hawaiian algae Chaetoceros. Initial concentrations of both
oils were 0.5, 1 and 2 mL/L. The hexadecatrienoic acid linearly decreased CH4
production by 97 %, while olive oil did not affect it. The hexadecatrienoic acid also
increased H2 accumulation. Release of C02 was linearly decreased by the
hexadecatrienoic acid, while olive oil increased it linearly. None of the oils had an effect
on final pH. Apparently fermented OM, as estimated through the VFA stoichiometry,
was linearly decreased by the hexadecatrienoic acid by 42%, while olive oil did not affect
it. The hexadecatrienoic acid decreased acetate and butyrate production, while
propionate production peaked at 1 mL/L. Olive oil tended to decrease acetate production
and increased propionate and butyrate. The hexadecatrienoic acid linearly decreased
NH; concentration. The hexadecatrienoic acid was a strong inhibitor of
methanogenesis, but it decreased fermentation and increased H2 accumulation. Olive oil

could be used to increase dietary energy without negatively affecting fermentation.

124

Introduction

Methane emissions by ruminants represent a loss of energy and also contribute to
global warming (Moss, 1993). Therefore, there is an economic and environmental
interest in decreasing CH4 formation in the rumen. Unsaturated fatty acids could
compete with methanogenesis for reducing equivalents during biohydrogenation in the
rumen (Czerkawski et al., 1966a, b).

Fats and oils have a variety of effects on ruminal fermentation and microbial
activities. There is a consistent decrease in ciliate protozoal numbers (Nagaraja et al.,
1997; Machmfiller and Kreuzer, 1998; Machmi'rller et al., 1998). Methanogenesis is
decreased in vitro and in vivo, and the VFA profile is generally shifted flom acetate and
butyrate towards propionate. However, the inhibition of CH4 production cannot be fully
accounted for by biohydrogenation competing for metabolic H or by the elimination of
methanogens associated with ciliate protozoa (N agaraja et al., 1997). Direct toxicity on
methanogens has been shown in mixed (Dong et al., 1997) and pure culture studies (Prins
et al., 1972). The reasons for oil toxicity on methanogens and Gram positive bacteria are
poorly understood, but they may involve alterations in cell membrane permeability that
affect nutrient uptake and regulation of intracellular pH (Nagaraja et al., 1997).

A problem associated to the use of fats and oils is an inhibition of ruminal
fermentation of non-lipid energy sources, especially fiber (N agaraja et al., 1997),
although some products like canola oil or cod liver oil have not affected NDF
disappearance (Dong et al., 1997). The objective of the present study was to examine the

effects of a hexadecatrienoic acid (C16; 6, 9, 12) isolated flom the Hawaiian marine algae

125

Chaetoceros, and olive oil predominant fatty acids, on fermentation by mixed ruminal

cultures. It was hypothesized that the oils would decrease CH4 production.

Materials and Methods
Oils and concentrations

An unsaturated hexadecatrienoic fatty acid (cis-C16;6,9, 12), isolated flom the
Hawaiian marine algae Chaetoceros (courtesy of J. -K. Wang, Department of Biosystems
and Engineering, University of Hawaii), and food grade olive oil (Pompeian Inc.,
Baltimore, MD) were examined as potential inhibitors of ruminal methanogenesis in
vitro. Initial concentrations of both oils were 0.5, 1, and 2 mL/L. Densities of the
hexadecatrienoic fatty acid and olive oil were 0.90 and 0.91 g/mL, respectively. Initial
concentrations were achieved by adding 25, 50, and 100 uL of each oil to 125 mL-

Wheaton bottles.

Ruminal fluid collection and incubation

Ruminal fluid was withdrawn prior to the morning feeding flom two mature
Holstein cows fed alfalfa hay. It was mixed together, and strained through two layers of
cheesecloth. It was then blended for 15 s, and again strained through two layers of
cheesecloth. One volume part of ruminal fluid was mixed with four volume parts of a
bicarbonate and phosphate buffer (Goering and Van Soest, 1975), and 50 mL of the
mixture anaerobically delivered into 125 mL-Wheaton bottles. All the bottles contained
250 mg of ground (0.2 mm screen mesh) alfalfa hay (1.8% N and 3.5% ash on a DM

basis) as substrate. Three samples of the ruminal fluid and buffer mixture were frozen for

126

subsequent determination of VFA initial concentrations. Bottles were sealed under an
O2-free CO2 atmosphere, and incubated in a shaking water bath at 39 °C for 24 h. At the
end of the incubation, bottles were allowed to cool to room temperature, and total gas
production volume was measured (Callaway and Martin, 1996). Fermentation was then

st0pped by adding 1 mL of a 10% phenol solution.

Analytical procedures

Methane and CO2 were analyzed (Callaway and Martin, 1996) using a Gow Mac
series 750 flame ionization detector gas chromatograph (Gow Mac Instruments Co.,
Bridgewater, NJ) equipped with a 4' x 1/4" DC 200 stainless steel column (150 °C,
carrier gas was N2 at 820 kPa). A RGD2 Reduction Gas Detector (Trace Analytical,
Menlo Park, CA), equipped with the same column, was used for H2 analysis. The
volume of gas produced was expressed as umoles at 25 °C and 1 atrn. A 5 mL-aliquot
was centrifuged (26,000 x g, 4 °C, 30 min) and the pH was measured in the supernatant
(Digital Benchtop pH Meter, Cole-Parmer Instrument Company, Vernon Hills, IL).
Volatile fatty acids, lactate, formate, ethanol, and the chemical additives were quantified
by differential reflactometry with a Waters 712 Wisp HPLC (Waters Associates Inc.,
Milford, MA) equipped with a BioRad HPX 87H column (BioRad Laboratories,
Hercules, CA). Separation was done by ion moderated partition. Solvent was 0.005 M

H2SO4 at 0.6 mL/min. Column temperature was 65 °C. Sample injection volume was 15

uL. Ammonia was analyzed as reported before (Chaney and Marbach, 1962).

127

Calculations

Apparently fermented substrate OM (F OM) was estimated flom the VFA
stoichiometry (Demeyer and Van Nevel, 1979), but using isobutyrate instead of caproate:
apparently fermented substrate (%) = (Acetate/2 + Propionate/2 + Isobutyrate + Butyrate
+ Valerate + Isovalerate) x 162 / substrate OM (mg), with all VFA expressed in

millimoles produced. It was assumed the oils were not fermented to VFA.

Statistical analysis

Four replications per oil and initial concentration were used. Responses were
modeled as: observation = control + concentration + concentration2 + concentration3 +
residual. Models with non-significant (P > 0.05) highest order coefficients were

considered non-significant.

Results and Discussion

The hexadecatrienoic acid decreased (P < 0.01) CH4 production by 97% (Table 5-
1). Possibly, the highly unsaturated hexadecatrienoic acid was toxic to methanogens
(Prins et al., 1972). In contrast, olive oil did not affect CH4 production (Table 5-1). Lack
of effect of other oils on ruminal methanogenesis has also been reported for protected fat
and crushed canola seed (Machmiiller et al., 1998). Oleic acid is the main component of
both olive oil (Kirisakis and Christie, 2000) and canola oil (Dong et al., 1997). However,
a product containing 74.6% oleic acid at a concentration of 0.5 g/L (equivalent to 0.37
g/L of olive oil) inhibited the growth of Methanobrevibacter ruminantium, regarded as

the main ruminal methanogen (Sharp et al., 1998), by 82% (Henderson, 1973). In the

128

current experiment, 2 mm of olive oil would be expected to contain at least 1.1 g/L of
oleic acid (Kirisakis and Christie, 2000). Given the high percentage of oleic acid in olive
oil, it is difficult to explain the lack of effect on CH4 production. Sensitivity of bacteria
to fatty acids is related to cell wall structure (Galbraith et al., 1971; Galbraith and Miller,
1973). The cell envelope of Methanobrevibacter differs florn other ruminal methanogens
in the presence of a pseudomurein layer instead of an S-layer (Sprott and Beveridge,
1993). It is possible that the presence of an S-layer confers other ruminal methanogens
some protection against fatty acid toxicity. Toxicity of oleic acid on pure cultures of
ruminal methano gens other than Methanobrevibacter needs to be investigated.

A 5-fold increase (P < 0.01) in H2 accumulation was observed at the maximum
hexadecatrienoic acid concentration (Table 5-1). An increase in H2 accumulation is often
a consequence of the inhibition of methanogenesis (Martin and Macy, 1985). Although
there is no stoichiometrical relationship, several experiments have shown greater H2
accumulation with stronger methanogenesis inhibition (Dong et al., 1997; Machmiiller
and Kreuzer, 1998). Similarly, in the present study, H2 accumulation increased about by
2-fold with 0.5 and 1 mL/L hexadecatrienoic acid, with a 25 and 53% decrease in CH4
production, respectively, and by 5-fold with 97% inhibition of methanogenesis at 2

mL/L.

129

Table 5-1. Effects of a hexadecatrienoic oil and olive oil on ruminal fermentation in vitro

Initial concentration, mL/L effect effect effect SEM3
P = P =* P =2

Acetate,
ionate, l
1
mol
Valerate, ol
Isovalerate,

fermented substrate, 60.8
%
CH4/FOM, l/m

Acetate,
' umol
l
Isobutyrate,
Valerate,
Isovalerate, umol

fermented substrate, 60.8
%

CH4/FOM,

 

9

term.
ZSignificance of the cubic term.

3 Standard error of the mean of the most significant model. If more than one model has a
P < 0.01, the highest order model was used.

130

There was a 16-fold increase in formate accumulation at 2 mL/L hexadecatrienoic
acid, although no formate accumulated at the lower concentrations (P = 0.04; quadratic
response; data not shown). Formate is used as a CH4 precursor in the rumen (Asanuma et
al., 1998). The hexadecatrienoic acid also increased (P < 0.05; cubic response; data not
shown) ethanol production. Interestingly, there was a 4.6- and 9-fold increase (P < 0.01;
data not shown) in formate accumulation at l and 2 leL olive oil, respectively. The
increase in this unusual H sink is difficult to understand, as olive oil did not inhibit CH4
prodirction. As there was a cubic tendency (P = 0.06; Table 5-1) for olive oil at 1 and 2
mL/L to increase OM apparent fermentability, it is possible that higher rates of
fermentation than formate utilization by methanogens (Asanuma et al., 199 8) or succinate
producers caused some increase in formate accumulation.

There was a linear decrease (P < 0.01) in CO2 release with the hexadecatrienoic
acid (Table 5-1). In previous results (Dong et al., 1997), supplementation with coconut
oil, but not with canola oil, caused a decrease in CO2 release. Cod liver oil caused a
slight decrease in CO2 release when supplementing grass hay, but not with concentrate
(Dong et al., 1997). In contrast, in the present study the release of CO2 was increased (P
= 0.02) by olive oil by 17% at 2 mL/L. This is difficult to reconcile with the changes it
caused in VFA production. A decrease (P = 0.05) in acetate of 46 umol, an increase in
propionate of 29 umol (P < 0.01), and an increase in butyrate of 42 umol (P = 0.02),
should all result in a not release of 47 umol C02 (C02 = acetate/2 + propionate/4 + 1.5 x
butyrate; (Bliimmel et al., 1997). This is clearly insufficient to explain the net increase

observed in CO2 release of approximately 300 umol.

131

The hexadecatrienoic acid decreased (P < 0.01) acetate and butyrate production
by more than 50%. Propionate production peaked at 1 mL/L hexadecatrienoic acid and
then decreased (P = 0.01; quadratic response). Olive oil increased (P < 0.01) propionate
production by 9%. This is important, as propionate is the main glucose precursor in
ruminants (Nagaraja et al., 1997). Both oils decreased (P < 0.01; data not shown) the
acetate to propionate ratio. Fats and oils generally, but not always, shift the VFA profile
flom acetate to propionate (N agaraja et al., 1997). The hexadecatrienoic acid tended (P =
0.07; data not shown) to decrease butyrate molar percentage. Butyrate molar percentage
is generally decreased by fats and oils as protozoal numbers and activities are decreased
(Nagaraja et al., 1997). On the contrary, olive oil increased (P = 0.02; data not shown)
butyrate molar percentage. In agreement, canola oil, which is also rich in oleic acid
(Dong et al., 1997), did not affect butyrate molar percentage or protozoal numbers in
continuous culture (Machmfiller et al., 1998). Other results have shown a decrease in
butyrate molar percentage when supplementing a concentrate diet, but not a roughage
diet, with canola (Dong et al., 1997).

There was a net disappearance of isobutyrate in all treatments. The reasons for
this are not clear, as isobutyrate initial concentration was not unusually high as compared
to similar experiments (data not shown). The hexadecatrienoic acid caused a 4-fold
decrease (P < 0.01) in isovalerate production. Isovalerate can be produced by the
catabolism of leucine (V oet and Voet, 1995). The hexadecatrienoic acid did inhibit
deamination, as reflected by the linear decrease (P < 0.01) in NH,+ concentration, which

is a consistent result flom oil supplementation (Nagaraja et al., 1997). Olive oil did not

132

seem to affect deamination, as reflected by the lack of change in N114+ and isoacids
concentrations.

The hexadecatrienoic acid caused a linear decrease (P < 0.01) in the alfalfa hay
substrate apparent fermentation and CH4 produced per milligram of FOM. In contrast,
supplementation with olive oil did not affect the substrate fermentation or CH4 produced
per milligram of FOM. Digestibility of roughage diets has been shown to be decreased
by coconut oil (Dong et al., 1997; Machmiiller and Kreuzer, 1998; Machmi'rller et al.,
1998) but not by protected fat, crushed canola seed, crushed linseed (Machmiiller et al.,
1998), canola or cod liver oil (Dong et al., 1997). Likely, the high degree of unsaturation
of the hexadecatrienoic acid was toxic for Ruminococcusflavefaciens and R. albus, which
are important cellulolytic bacteria with a Gram-positive-type cell wall (Nagaraja et al.,
1997)

Oleic acid, which is the most abundant fatty acid in olive oil (Kirisakis and
Christie, 2000), did not affect ruminal Gram negative bacteria (Henderson, 1973;
Maczulak et al., 1981), but was strongly inhibitory for Butyrivibrio B 835 and
Ruminococcus 4263/ 1. A product containing 74.6% oleic acid at a concentration of 0.1
g/L inhibited the growth of these species by more than 80% (Henderson, 1973).
Interestingly, oleic acid at lower concentrations (0.01 g/L or less) was actually
stimulatory for Butyrivibrio B 835 (Henderson, 1973; Maczulak et al., 1981), although it
inhibited other strains of B. fibrisolvens (Maczulak et al., 1981). One strain of
Ruminococcus albus, and two strains of R flavefaciens were strongly inhibited by oleic
acid, even at a concentration as low as 0.005 g/L (Maczulak et al., 1981). Hence, it is

difficult to explain why olive oil at 2 mL/L (which would represent at least 1 mL/L, or

133

0.891 g/L, of oleic acid) did not inhibit fermentation or CH4 production. It is possible
that Gram negative cellulolytic species, like F ibrobacter succinogenes (Maczulak et al.,
1981), could have colonized fiber surfaces left by the inhibited Ruminococci. Also, as
large strain differences can occur (Henderson, 1973), it is possible that different strains of
the same species were present during the current experiment incubations and were less
affected.

Alternatively, cholesterol and ergocalciferol at 0.45 mM have caused some
reversal of the inhibition caused by Na lauryl sulfate, lauric, and linoleic acids on non-
ruminal microorganisms (Galbraith et al., 1971). The presence of steroids in olive oil
(Kirisakis and Christie, 2000) may have relieved the inhibition caused by long-chain fatty
acids. However, the concentrations of steroids in the present study incubations should
have been much lower than levels reported to reverse the fatty acid inhibition.

In agreement with the present results, the effects of oil supplementation on
ruminal pH are generally minor (Dong et al., 1997; Nagaraja et al., 1997; Machmi'rller et

al., 1998).

Conclusions

The hexadecatrienoic acid extracted flom the Hawaiian algae Chaetoceros was a
very strong inhibitor of ruminal methanogenesis in vitro. Before it could be used as an
inhibitor of methanogenesis in vivo, it would be necessary to overcome its negative
effects on fermentation, as well as the increase it caused in H2 and formate accumulation,
and ethanol production. Perhaps the addition of alternative electron acceptors, or

acetogens (Nollet et al., 1997), could rechannel electrons away flom H2, formate and

134

ethanol, and also improve fermentation. Although olive oil did not inhibit CH4
production, it could be used to increase the energy content of ruminant diets and the
supply of propionate, the main glucose precursor, without depressing ruminal

fermentation.

135

REFERENCES

Asanuma, N., M. Iwamoto, and T. Hino. 1998. Formate metabolism by ruminal
' microorganisms in relation to methanogenesis. Anim. Sci. Technol. (Jpn.) 69:
576-584.

Bliimmel, M., H. P. S. Makkar, and K. Becker. 1997. In vitro gas production: a technique
revisited. J. Anim. Physiol. a Anim. Nutr. 77: 24-34.

Callaway, T. R. and S. A. Martin. 1996. Effects of organic acid and monensin treatment

on in vitro mixed ruminal microorganism fermentation of cracked corn. J. Anim.
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Chaney, A. L. and E. P. Marbach. 1962. Modified reagents for determination of urea and
' ammonia. Clin. Chem. 8: 130-132.

Czerkawski, J. W., K. L. Blaxter, and F. W. Wainman. 1966a. The effect of linseed oil
and of linseed oil fatty acids incorporated in the diet on the metabolism of sheep.
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Czerkawski, J. W., K. L. Blaxter, and F. W. Wainman. 1966b. The metabolism of oleic,
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production. Br. J. Nutr. 20: 349-362.

Demeyer, D. I. and C. J. Van Nevel. 1979. Effect of defaunation on the metabolism of
rumen micro-organisms. Br. J. Nutr. 42: 51 5-524.

Dong, Y., H. D. Bae, T. A. McAllister, G. W. Mathison, and K.-J. Cheng. 1997. Lipid-
induced depression of methane production and digestibility in the artificial rumen
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Galbraith, H. and T. B. Miller. 1973. Effect of long chain fatty acids on bacterial
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Galbraith, H., T. B. Miller, A. M. Paton, and J. K. Thompson. 1971 . Antibacterial activity
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Goering, H. K. and P. N. Van Soest. 1975. Forage Fiber Analyses (Apparatus, Reagents,
Procedures and some Applications). 379, ARS-USDA, Washington DC.

Henderson, C. 1973. The effects of fatty acids on pure cultures of rumen bacteria. J.
Agric. Sci. (Camb.) 81: 107-112.

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Kirisakis, A. and W. W. Christie. 2000. Analysis of edible oils. In: J. Harwood and R.
Aparicio (eds) Handbook of Olive Oil. Analysis and Properties. p 129-15 8.
Aspen Publishers, Inc., Gaithersburg, MD.

Machmiiller, A. and M. Kreuzer. 1998. Methane suppression by coconut oil and
associated effects on nutrient and energy balance in sheep. Can. J. Anim. Sci. 79:
65-72.

Machmiiller, A., D. A. Ossowski, M. Wanner, and M. Kreuzer. 1998. Potential of various
fatty feeds to reduce methane release flom rumen fermentation in vitro (Rusitec).
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Maczulak, A. E., B. A. Dehority, and D. L. Palmquist. 1981. Effects of long-chain fatty
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Prins, R. A., C. J. Van Nevel, and D. I. Demeyer. 1972. Pure culture studies of inhibitors
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137

CHAPTER 6
Effects of combinations of inhibitors of methanogenesis with crotonic acid or 3-

butenoic acid on in vitro ruminal fermentation and methane production

Abstract

It was hypothesized that the reduction of crotonate or 3-butenoate to butyrate
would utilize reducing equivalents not incorporated into CH4 formation when the latter is
inhibited by lumazine, propynoic acid, or ethyl 2-butynoate. This would avoid the
accumMation of unusual reduced products without a nutritional value (H2, formate, and
ethanol). In six experiments, three inhibitors of CH4 formation (lumazine, propynoic
acid, and ethyl 2-butynoate), each at three different initial concentrations, were combined
with either crotonic acid or 3-butenoic acid, each at two different initial concentrations.
A 4:1 buffer and ruminal fluid mixture was incubated in Wheaton bottles for 24 h, using
ground alfalfa hay as substrate. The inhibition of methanogenesis caused by lumazine
was smaller than previously observed, and there was no accumulation of reduced end
products without a nutritional value. Propynoic acid at its maximum initial concentration
decreased CH4 production by more than two thirds. Crotonic acid and 3-butenoic acid
were ineffective in avoiding the accumulation of H2 and formate or decreasing ethanol
production. Ethyl 2-butynoate suppressed methanogenesis by more than 90%. Crotonic
acid caused some decrease in H2 accumulation and ethanol production, while 3-butenoic
acid was ineffective. Crotonic acid and 3-butenoic acid were ineffective in avoiding the
formation of unusual reduced products partly because they were also metabolized to

acetate, thereby releasing, rather than incorporating, reducing equivalents. Incomplete

138

disappearance of crotonic acid could also explain the lack of effectiveness of this

additive.

Introduction

Ruminal methanogenesis represents a substantial loss of energy to the animal and
is also a major source of greenhouse gas emissions flom agriculture (Newbold et al.,
2001). However, methanogenesis is the main means of disposal of metabolic H in
ruminal fermentation (Newbold et al., 2001) and its inhibition can decrease fermentation
as the interspecies transfer of H2 is disrupted and reduced cofactors do not get reoxidized
(N agaraja et al., 1997). Propynoic acid and ethyl 2-butynoate decreased methanogenesis
in vitro by over 70%; however, total OM (substrate plus additive) apparent fermentability
was decreased by both compounds. Lumazine also decreased methanogenesis by about
50%, but inhibited fermentation (Ungerfeld et al., 2000). Ethyl 2-butynoate also
inhibited the fermentation of the alfalfa hay substrate. These compounds also caused the
formation of unusual products of fementation, like H2, ethanol, and formate, without a
nutritional value to the host animal.

B-Hydroxybutyrate, crotonic acid, and 3-butenoic acid, had little effects on
methanogenesis, but seemed to stimulate fermentation of the alfalfa hay substrate. It is
possible that these organic acids benefited fermentation by acting as alternative electron
sinks. The inhibition of methanogenesis might have caused the formation of unusual
reduced products by disrupting the interspecies H transfer. It was hypothesized that the
combination of inhibitors of methanogenesis and compounds that stimulated fermentation

would lift the constraints caused by the former. The objective of this series of six

139

experiments was to examine the effects of combinations of lumazine, propynoic acid, and
ethyl 2-butynoate, with crotonic acid or 3-butenoic acid, on fermentation of mixed

ruminal cultures.

Material and Methods
Experiment I

The experimental treatments of the 3 x 2 factorial arrangement were: 1) Lumazine
(Sigma L 0380 for all experiments) 0 mM, crotonic acid (Sigma C4630 for all
experiments) 0 mM (double control); 2) Lumazine 0 mM, crotonic acid 8 mM; 3)
Lumazine 0.3 mM, crotonic acid 0 mM; 4) Lumazine 0.3 mM, crotonic acid 8 mM; 5)
Lumazine 0.6 mM, crotonic acid 0 mM; 6) Lumazine 0.6 mM, crotonic acid 8 mM.
Lumazine was added as a solid, while crotonic acid was added as a 1 mL-solution.
Crotonic acid controls received l-mL of deionized water.

Ruminal fluid was withdrawn prior to the morning feeding flom two mature, non-
lactating Holstein cows fed alfalfa hay. It was mixed, and strained through two layers of
cheesecloth. It was then blended for 15 s and again strained through two layers of
cheesecloth. One volume part of ruminal fluid was mixed with four volume parts of
buffer (Goering and Van Soest, 1975) and 80 mL of the mixture anaerobically delivered
into 160-mL Wheaton bottles. All the bottles contained 500 mg of ground (0.2 mm mesh
screen) alfalfa hay (1 .8% N, DM base) as substrate. Lumazine was added to the
corresponding bottles as a solid, while crotonic acid was added as a solution (1
mL/bottle). Crotonic acid controls received 1 mL of deionized water. Three samples of

the ruminal fluid and buffer mixture were flozen for subsequent determination of VFA

140

initial concentrations. Bottles were sealed under an O2-flee CO2 atmosphere and
incubated in a shaking water bath at 39 °C for 24 h. At the end of the incubation, bottles
were allowed to cool to room temperature and total gas production volume was measured
(Callaway and Martin, 1996). F errnentation was then stopped by adding 1 mL of a 10%
phenol solution.

Methane and CO2 were analyzed (Callaway and Martin, 1996), using a Gow Mac
series 750 flame ionization detector gas chromatograph (Gow Mac Instruments Co.,
Bridgewater, NJ) equipped with a 4' x 1/4" DC 200 column (150 °C, carrier gas was N2 at
820 Kpa). A RGD2 Reduction Gas Detector (Trace Analytical, Menlo Park, CA),
equipped with the same type of column, was used for H2 analysis. Gas production was
expressed as micromoles at 25 °C and 1 atrn. A 5-mL aliquot was centrifuged (26,000 x
g, 4 °C, 30 min) and pH measured in the supernatant (Digital Benchtop pH Meter, Cole-
Parmer Instrument Company, Vernon Hills, IL). Volatile fatty acids, lactate, formate,
and ethanol were quantified by differential reflactometry with a Waters HPLC (Waters
Associates Inc., Milford, MA) equipped with a BioRad HPX 87H column (BioRad
Laboratories, Hercules, CA). Solvent was 0.005 M H2SO4 at 0.6 mL/min. Column
temperature was 65 °C. Sample injection volume was 15 uL. Crotonic acid co-eluted
flom the HPLC column with isovalerate. Consequently, isovalerate production is not
reported. Ammonia concentration was determined as reported before (Chaney and
Marbach, 1962).

Four replicates per compound and concentration were used. The experimental
model was: response = control + linear effect of lumazine + quadratic effect of lumazine

+ crotonic acid effect + crotonic acid by linear effect of lumazine + crotonic acid by

141

quadratic effect of lumazine + residual. Non-significant (P > 0.15) quadratic terms were
removed from the model. When significant (P < 0.05) interactions or tendencies (P <
0.15) were found, means were compared by Fisher least square differences (Rao, 1998).

Significance was declared at P < 0.05.

Experiment 2

The experimental treatments of the 3 x 2 factorial arrangement were: 1) Lumazine
0 mM, 3-butenoic acid (Acros 15883 for all experiments) 0 mM (double control); 2)
Lumazine 0 mM, 3-butenoic acid 4 mM; 3) Lumazine 0.6 mM, 3-butenoic acid 0 mM; 4)
Lumazine 0.6 mM, 3-butenoic acid 4 mM; 5) Lumazine 1.2 mM, 3-butenoic acid 0 mM;
6) Lumazine 1.2 mM, 3-butenoic acid 4 mM. Lumazine was added as a solid, while 3-
butenoic acid was added as a 1 mL-solution. 3-Butenoic acid controls received l-mL of
deionized water. Ruminal fluid collection and incubation, analytical procedures, and
statistical analysis were done as described for Experiment 1, except that H2 concentration
could not be measured because of a malfunction of the detector. 3-Butenoic acid co-
eluted flom the HPLC column with propionate. Therefore, reported propionate

production assume total disappearance of 3-butenoic acid.

Experiment 3

The 3 x 2 factorial arrangement of treatments included: 1) Propynoic acid (Acros
13150 for all experiments) 0 mM, crotonic acid 0 mM (double control); 2) Propynoic
acid 0 mM, crotonic acid 4 mM; 3) Propynoic acid 2 mM, crotonic acid 0 mM; 4)

Propynoic acid 2 mM, crotonic acid 4 mM; 5) Pr0pynoic acid 4 mM, crotonic acid 0 mM;

142

6) Propynoic acid 4 mM, crotonic acid 4 mM. Both chemicals were added as 1 mL-
solutions and controls received 1 mL of deionized water. Ruminal fluid collection and
incubation, analytical procedures, and statistical analysis, were done as described for

Experiment 1. Propynoic acid disappearance was also reported.

Experiment 4

The experimental treatments of the 3 x 2 factorial arrangement were: 1) Propynoic
acid 0 mM, 3-butenoic acid 0 mM (double control); 2) Propynoic acid 0 mM, 3-butenoic
acid 4 mM; 3) Propynoic acid 2 mM, 3-butenoic acid 0 mM; 4) Propynoic acid 2 mM, 3-
butenoic acid 4 mM; 5) Propynoic acid 4 mM, 3-butenoic acid 0 mM; 6) Propynoic acid
4 mM, 3-butenoic acid 4 mM. Both chemicals were added as 1 mL-solutions and
controls received 1 mL of deionized water. Ruminal fluid collection and incubation,
analytical procedures, and statistical analysis were done as described for Experiment 1.

Propynoic acid disappearance was also reported.

Experiment 5

The 3 x 2 factorial arrangement included: 1) Ethyl 2-butynoate (GFS Chemicals
3132 for all experiments) 0 mM, crotonic acid 0 mM (double control); 2) Ethyl 2-
butynoate 0 mM, crotonic acid 4 mM; 3) Ethyl 2-butynoate 4 mM, crotonic acid 0 mM;
4) Ethyl 2-butynoate 4 mM, crotonic acid 4 mM; 5) Ethyl 2-butynoate 8 mM, crotonic
acid 0 mM; 6) Ethyl 2-butynoate 8 mM, crotonic acid 4 mM. Ethyl 2-butynoate was
added as a liquid, while crotonic acid was added as a 1 mL-solution. Crotonic acid

controls received 1 mL of deionized water. Ruminal fluid collection and incubation,

143

analytical procedures, and statistical analysis were done as described for Experiment 1.
Ethyl 2-butynoate co-eluted flom the HPLC column with butyrate. Therefore, reported

butyrate production assume total disappearance of ethyl 2-butynoate.

Experiment 6

The experimental treatments of the 3 x 2 factorial arrangement included: 1) Ethyl
2-butynoate 0 mM, 3-butenoic acid 0 mM (double control); 2) Ethyl 2-butynoate 0 mM,
3-butenoic acid 4 mM; 3) Ethyl 2-butynoate 4 mM, 3-butenoic acid 0 mM; 4) Ethyl 2-
butynoate 4 mM, 3-butenoic acid 4 mM; 5) Ethyl 2-butynoate 8 mM, 3-butenoic acid 0
mM; 6) Ethyl 2-butynoate 8 mM, 3-butenoic acid 4 mM. Ethyl 2-butynoate was added as
a liquid, while 3-butenoic acid was added as a 1 mL-solution. 3-Butenoic acid controls
received 1 mL of deionized water. Ruminal fluid collection and incubation, analytical
procedures, and statistical analysis were done as described for Experiment 1. Separation
of crotonic acid flom isovalerate was achieved by dropping the column temperature to 45
°C. Ethyl 2-butynoate was quantified by flame ionization detector with Perkin Elmer
8500 GC equipped with an AllTech AT-l column. Solvent was ether at 4 mL/min flow

rate. Start temperature was 80 °C. Ramp rate 1 was 10 °C/min. End temperature 1 was

150 °C. Ramp rate 2 was 30 °C/min, and end temperature 2 was 250 °C.

144

Results
Experiment 1

No interactions were found for CH4 production. Lumazine decreased (P < 0.01)
CH4 production by 9 and 10% with and without crotonic acid, respectively (Table 6-1).
Crotonic acid decreased (P < 0.01) methanogenesis by 10%. There were linear by linear
and linear by quadratic interactions (P < 0.01) for CO2 release. In the absence of crotonic
acid, lumazine at 0.6 mM decreased (P < 0.05) CO2 release by 18%; however, there were
no effects in the presence of crotonic acid.

There were no interactions for H2 and formate accumulation or ethanol
production. No increase in H2 accumulation was observed (Table 6-1). Unexpectedly,
formate accumulation was strongly decreased (P < 0.01) by lumazine, while crotonic acid
had no effect. None of the additives had an effect on ethanol production.

There was a linear by quadratic interaction for total VFA concentration (P =
0.04). In the absence of crotonic acid, lumazine did not affect total VFA concentration,
while in its presence, lumazine at 0.6 mM decreased (P < 0.05) total VFA concentration.
Crotonic acid increased (P < 0.01) total VFA concentration at 0 and 0.3 mM lumazine.
There was a linear by quadratic interaction on acetate production (P < 0.01). Lumazine
at 0.6 mM decreased (P < 0.05) acetate production with and without crotonic acid.
Crotonic acid increased (P < 0.01) acetate production only at 0 and 0.3 mM lumazine.
There was a tendency (P = 0.07) for a linear by quadratic interaction for propionate
production. Lumazine decreased (P < 0.01) propionate production both in the absence

and presence of crotonic acid. Crotonic acid decreased (P < 0.05) propionate production

145

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at 0.3 and 0.6 mM lumazine. There was a linear by quadratic interaction for butyrate
production. In the absence of crotonic acid, lumazine did not affect butyrate production.
However, butyrate production was decreased (P < 0.05) at 8 mM crotonic acid, lumazine .
and 0.6 mM lumazine. Crotonic acid increased (P < 0.01) butyrate production by about
2-fold. There were linear and linear by quadratic interactions (P < 0.01) for isobutyrate
production. Lumazine at 0.3 and 0.6 mM increased (P < 0.01) isobutyrate production by
3.7- and 5.1-fold, respectively. Crotonic acid decreased (P < 0.05) isobutyrate
production only at 0.6 mM lumazine. There were no interactions for final pH. Final pH
was not affected by the additives. Ammonia concentration peaked (P < 0.01; quadratic

response) at 0.3 mM lumazine.

Experiment 2

There was a linear by linear interaction (P = 0.03) for CH4 production. Lumazine
decreased (P < 0.01; Table 6-2) CH4 production by 15 and 24 % with and without 3-
butenoic acid, respectively. 3-Butenoic acid decreased (P < 0.05) CH4 production only at
1.2 mM lumazine (linear interaction P = 0.03). There were no effects of the additives on
CO2 release.

No interactions for H2 and formate accumulation or ethanol production. No
increase in H2 accumulation was observed (Table 6-2). Surprisingly, 3-butenoic acid
increased formate accumulation, while lumazine had no effect. None of the additives
influenced ethanol production.

There were no interactions for total VFA concentration or acetate, butyrate,

valerate or isovalerate production. 3-Butenoic acid increased (P < 0.01; Table 6-2) total

147

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VFA concentration, while lumazine did not affect it. Lumazine linearly decreased (P <
0.01) acetate production, while 3-butenoic acid increased (P < 0.01) it. There was a
tendency (P = 0.09) for a linear by linear interaction for propionate production.
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decreased (P < 0.05) propionate production at 0.6 and 1.2 mM lumazine. Both lumazine
and 3-butenoic acid increased butyrate production. Lumazine at 0.6 and 1.2 mM caused
(P < 0.01) a 7- and 9-fold increase in isobutyrate production, respectively. It caused a
quadratic decrease (P = 0.03) in isovalerate production and did not affect valerate. 3-
Butenoic acid did not affect isobutyrate, valerate, or isovalerate production. There was a
linear interaction for final pH (P = 0.02; Table 6-2). In the absence of 3-butenoic acid,
lumazine did not affect pH, but it decreased it (P < 0.05) at 4 mM 3-butenoic acid.
Lumazine at 1.2 mM tended (P = 0.11; quadratic response) to decrease ammonia

concentration.

Experiment 3

There were no interactions for CH4 production or C02 release. Both at 0 and 4
mM crotonic acid, propynoic acid decreased (P < 0.01; quadratic response) CH4
production by 69% (Table 6-3). Crotonic acid decreased (P = 0.02) CH4 production by 8,
19, and 9% at O, 2, and 4 mM propynoic acid initial concentration, respectively. Release
of C02 was linearly decreased (P = 0.02) by propynoic acid, but not affected by crotonic
acid.

There was a linear by linear interaction (P = 0.04) for H2 accumulation.

Propynoic acid strongly increased (P < 0.01; quadratic response) H2 accumulation. At 4

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mM propynoic acid, crotonic acid decreased (P < 0.05) H2 accumulation (linear
interaction P = 0.04). There were also linear and linear by quadratic interactions (P <
0.01) on formate production (Table 6-3). In the absence of crotonic acid, accumulation
of formate was minimal (and negative) at 2 mM propynoic acid, but increased by more
than 3-fold of control levels at 4 mM propynoic acid (P < 0.05). At 4 mM crotonic acid,
formate accumulation was increased (P < 0.05) by 2 and 4 mM propynoic acid. Crotonic
acid decreased (P < 0.05) formate accumulation at O and 4 mM propynoic acid, but
increased (P < 0.05) it at 2 mM. There were no interactions for ethanol production.
Production of ethanol was linearly increased (P < 0.01) by propynoic acid and not
afi‘ected by crotonic acid.

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affected by its initial concentration or the presence of crotonic acid (Table 6-3).

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(P < 0.01) total VFA concentration, while propynoic acid decreased (P < 0.01; quadratic
response; Table 6-3) it. There were no interactions for acetate production. Propynoic
acid decreased (P < 0.01; quadratic response) acetate production, while crotonic acid
increased (P = 0.02) it. There was a linear by linear interaction (P = 0.04) and a tendency
(P = 0.06) for a linear by quadratic interaction for propionate production. Propynoic acid
increased (P = 0.03; quadratic response) propionate production. Crotonic acid decreased
(P < 0.05) propionate production at 0 mM propynoic acid and not affected it at 2 and 4
mM propynoic acid. There were no interactions for butyrate production. Propynoic acid
decreased (P < 0.01; quadratic response) butyrate production, and crotonic acid increased

(P < 0.01) it. There was a linear by linear interaction (P = 0.02) for isobutyrate

151

production. Isobutyrate production was decreased (P < 0.01; quadratic response) by
pr0pynoic acid. Crotonic acid decreased (P < 0.05) isobutyrate production only in the
absence of propynoic acid. Valerate production was decreased (P < 0.01) by propynoic
acid, and not affected by crotonic acid. In the absence of crotonic acid, propynoic acid
decreased (P < 0.05) final pH. There was a tendency (P = 0.07) for a linear by linear
interaction, and a linear by quadratic interaction (P = 0.04) for final pH. At 4 mM
crotonic acid, the lowest pH was observed at 2 mM propynoic acid (P < 0.05). Ammonia

concentration was decreased (P < 0.01; quadratic response) by propynoic acid.

Experiment 4

There were linear by linear and linear by quadratic interactions for CH4
production. Propynoic acid decreased (P < 0.01) CH4 production by approximately 69%
at both concentrations of 3-butenoic acid (Table 6-4). 3-Butenoic acid decreased (P <
0.05) CH4 production only at 2 mM propynoic acid (linear and linear by quadratic
interaction P < 0.01). There were no interactions for C02 release. Release of CO; was
decreased (P = 0.01) by propynoic acid, and not affected by 3-butenoic acid.

There were linear by linear (P = 0.01) and linear by quadratic (P = 0.04)
interactions for H2 accumulation. Propynoic acid increased (P < 0.01; quadratic
response) the accumulation of H2. 3-Butenoic acid increased H2 accumulation at 2 mM
propynoic acid (Table 6-4). Propynoic acid increased formate (P = 0.03; quadratic
relationship) accumulation. Unexpectedly, 3-butenoic acid also increased (P < 0.01)

formate accumulation. Ethanol production was similar among treatments.

152

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Little propynoic acid disappeared during the incubation period. Disappearance
was not affected by its initial concentration or the presence of 3-butenoic acid (Table 6-
4).

There were no interactions for total VF A concentration. Propynoic acid
decreased (P < 0.01; Table 6-4) total VFA, while crotonic acid increased (P < 0.01) it.
There were no interactions for acetate production. Acetate production was decreased (P
< 0.01) by propynoic acid, and tended (P = 0.08) to increase with 3-butenoic acid. There
were linear by linear (P = 0.04) and linear by quadratic (P = 0.02) interactions for
propionate production. In the absence of 3-butenoic acid, propionate production was
increased (P < 0.05) by propynoic acid only at 4 mM. At 4 mM 3-butenoic acid,
propionate production was increased (P < 0.05) by both concentrations of propynoic acid.
There were no interactions for butyrate or isobutyrate production. Butyrate production
was increased by both propynoic acid (P = 0.02) and 3-butenoic acid (P < 0.01). None of
the additives affected isobutyrate production. There was a linear interaction (P = 0.03)
for isovalerate production. Both at 0 (P = 0.02; quadratic response) and 4 mM 3-butenoic
acid, propynoic acid strongly decreased (P < 0.01) valerate production. 3—Butenoic acid
increased (P < 0.05) valerate production at 4 mM propynoic acid. Propynoic acid
decreased (P = 0.02) isovalerate production. There were no interactions for final pH or
NH," concentration. Fina] pH was not affected by the additives. Ammonia concentration

was decreased (P < 0.01) by both additives.

154

Experiment 5

There were no interactions for CH4 production or C02 release. Ethyl 2-butynoate
linearly decreased (P < 0.01) CH4 production by more than 90% at both concentrations of
crotonic acid. Crotonic acid did not affect methanogenesis (Table 6-5). Ethyl 2-
butynoate decreased (P < 0.01) C02 release by 39%, while crotonic acid did not afi‘ect it.

There was a linear by linear interaction (P = 0.04) and a tendency (P = 0.06) for a
linear by quadratic interaction for H2 accumulation. Ethyl 2-butynoate increased the
accumulation of H2 (P < 0.01; quadratic response; Table 6-5). Dihydrogen accumulation
was decreased (P < 0.05) by crotonic acid only at 4 mM ethyl 2-butynoate. There was a
tendency (P < 0.15) for a linear by quadratic interaction for formate accumulation. Ethyl
2-butynoate increased (P < 0.01; quadratic response) formate accumulation. Crotonic
acid increased (P < 0.05) formate accumulation at 8 mM ethyl 2-butynoate. There were
no interactions for ethanol production. Ethyl 2-butynoate increased (P < 0.01; quadratic
response) ethanol production, while crotonic acid did not affect it.

There were no interactions for total VFA concentration or acetate production.
Ethyl 2-butynoate decreased (P < 0.01; quadratic response) total VFA concentration,
while crotonic acid did not affect it (Table 6-5). Ethyl 2-butynoate decreased (P < 0.01;
quadratic response) acetate production, while crotonic acid did not affect it. There were
tendencies for linear by linear (P = 0.13) and linear by quadratic (P = 0.11) interactions
for propionate production. At 0 mM crotonic acid, propionate production was greatest (P
< 0.05) at 4 mM ethyl 2-butynoate. At 4 mM crotonic acid, propionate production was
not affected by 4 mM ethyl 2-butynoate, and decreased (P < 0.05) by 8 mM ethyl 2-

butynoate. Crotonic acid decreased (P < 0.05) propionate production only at 4 mM ethyl

155

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156

2-butynoate. There was a tendency (P = 0.07) for a linear by quadratic interaction for
butyrate production. Butyrate production was increased (P < 0.01) by crotonic acid. In
the absence of crotonic acid, butyrate production was increased by 8 mM ethyl 2-
butynoate. There were no interactions for isobutyrate and valerate production, final pH
and NH;+ concentration. Isobutyrate production was increased (P < 0.01) by ethyl 2-
butynoate, and not affected by crotonic acid. Valerate production was decreased (P <
0.01; quadratic response) by ethyl 2-butynoate, and not affected by crotonic acid. Effects
of ethyl 2-butynoate on final pH (P < 0.01; quadratic response) were of little importance.
Ammonia concentration was decreased (P = 0.01; quadratic response) by ethyl 2-

butynoate and tended (P = 0.08) to be decreased by crotonic acid.

Experiment 6

There was a linear by linear interaction (P = 0.02) and a tendency (P = 0.05) for a
linear by quadratic interaction for CH4 production. Ethyl 2-butynoate decreased (P <
0.01) CH4 below the limits of detection (Table 6-6). Methane production was increased
(P < 0.05) by 3-butenoic acid only in the absence of ethyl 2-butynoate. There were no
interactions for C02 release. Ethyl 2-butynoate decreased (P = 0.03; quadratic response)
C02 release, while 3-butenoic acid increased (P = 0.04) it.

There were no interactions for H2 and formate accumulation and ethanol
production. Ethyl 2-butynoate increased (P < 0.01; quadratic response; Table 6-6) the
accumulation of H2, which was not relieved by 3-butenoic acid. Ethyl 2-butynoate also
increased the accumulation of formate (P = 0.01; quadratic response) and ethanol

production (P < 0.01), none of which was relieved by 3-butenoic acid.

157

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158

Over 97% of ethyl 2-butynoate and 100% of 3-butenoic acid disappeared during
fermentation (data not shown).

There were no interactions for total VFA, and a tendency (P < 0.15) for a linear
by linear interaction for acetate production. Total VFA concentration and acetate
production were decreased (P < 0.01) by ethyl 2-butynoate, and increased (P < 0.01) by
3-butenoic acid (Table 6-6). There were no interactions for propionate production.
Propionate production was decreased (P < 0.01; quadratic response) by ethyl 2-butynoate
at 8 mM, and tended (P = 0.06) to be decreased by 3-butenoic acid. Butyrate production
was increased by both ethyl 2-butynoate (P < 0.01; quadratic response) and 3-butenoic
acid (P < 0.01). There were no interactions for isobutyrate, valerate and isovalerate
production, final pH and NH4+ concentration. Isobutyrate production was increased by
ethyl 2-butynoate (P < 0.01; quadratic response), and not affected by 3-butenoic acid.
Valerate and isovalerate production were decreased (P < 0.01; quadratic response) by
ethyl 2-butynoate, and not affected by 3-butenoic acid. None of the additives affected the

final pH or NH: concentration.

Discussion
Lumazine

In previous work, lumazine at 0.6 mM lowered CH4 formation by 50%, and there
was no additional reduction with 1.2 mM (Ungerfeld et al., 2002). Thus, a maximum
initial concentration of 0.6 mM was chosen for Experiment 1. However, the inhibition
achieved was substantially smaller than observed before (Ungerfeld et al., 2002). It was

therefore decided to double the initial concentrations for Experiment 2. Nevertheless, the

159

inhibition of CH4 formation was still lower than previously reported. Other responses
have also been inconsistent or difficult to explain. In the previous study, acetate
production was not affected even though CH4 production was inhibited by 50%
(Ungerfeld et al., 2002), while in Experiments 1 and 2 acetate production was decreased
despite a much lower inhibition of methanogenesis. In Experiment 1 and in previous
work (Ungerfeld et al., 2002), the acetate to propionate ratio increased (data not shown),
which is contrary to the shift from acetate to propionate generally observed when CH4
production is inhibited (N agaraja et al., 1997). In Experiment 2, the acetate to propionate
ratio was not affected by lumazine (data not shown). The observed increase in the acetate
to propionate ratio, together with the decreased C02 production and lack of H2
accumulation when methanogenesis was inhibited, led to the speculation that perhaps
lumazine could somehow stimulate reductive acetogenesis (Ungerfeld et al., 2002).
Tetrahydrofolate, like lumazine, a pterin derivative, is involved in reductive acetogenesis
(Drake, 1994). Perhaps some acetogens could use lumazine to synthesize
tetrahydrofolate.

Most puzzling, the addition of lumazine has resulted in large changes in
isobutyrate production. A quadratic response, with a lO-fold decrease in isobutyrate
production at 0.6 mM lumazine, and an almost complete recovery at 1.2 mM was first
noticed (Ungerfeld et al., 2002). In Experiment 1, 4- to S-fold increases were observed
at 0.6 mM lumazine. In Experiment 2, 8- to 9-fold increases in isobutyrate production
were observed at 1.2 mM lumazine. Isobutyrate is a product of valine fermentation (V oet
and Voet, 1995). It is possible that lumazine somehow alters valine metabolism.

Alternatively, isobutyrate may be a product of the catabolism of lumazine itself. In

160

Experiment 1, the C content of the extra isobutyrate represented 91 and 69% of the C
present in lumazine, at 0.3 and 0.6 mM, respectively (data not shown). In Experiment 2,
C in extra isobutyrate accounted for 129 and 83% of the C from added lumazine, at 0.6
and 1.2 mM, respectively (data not shown). However, as there are six C atoms per
molecule of lumazine, and four per molecule of isobutyrate, one third of the C contained
in lumazine should not appear in isobutyrate (assuming a molecule of isobutyrate is not
formed fiom C provided from two lumazine molecules). This elevates the amount of
lumazine necessary to form the extra isobutyrate to 137 and 104% of added lumazine at
0.3 and 0.6 mM lumazine in Experiment 1, and 194 and 125% of added lumazine at 0.6
and 1.2 mM lumazine in Experiment 2 (data not shown). Therefore, it seems unlikely
that the increase in isobutyrate production was a result of lumazine catabolism.

In Experiment 1, formate production at 0 mM lumazine was unusually high
(Table 6—1). Formate concentration in the double control (0 mM both additives) was 2.27
mM (data not shown), which is between about 100- and ZOO-fold higher than previously
reported values (Hungate, 1967; Hungate et al., 1970; Asanuma et al., 1998). Formate
concentration dropped by about 50 and 70% at 0.3 and 0.6 mM lumazine, respectively,
but the absolute values appeared high compared to previously reported values. Highly
fermentable substrates can lead to earlier and higher peaks of formate concentration in
mixed ruminal batch cultures due to higher rates of formate production than utilization by
methanogens (Asanuma et al., 1998). However, a slowly fermentable substrate (alfalfa
hay) was used in the current experiments. Comparison of CH4 production in Experiment
1 control treatment with controls in Experiments 2-6, would not support the premise that

lower growth rates of methanogens in Experiment 1 resulted in formate accumulation.

161

Perhaps the utilization of formate as an electron donor for succinate (Asanuma et al.,
1998) or butyrate formation was low in Experiment 1. Alternatively, more pyruvate may
have been metabolized into formate by pyruvate formate lyase reaction (Russell and
Wallace, 1997), or more formate may have been formed via glyoxylate (Asanuma et al.,

1999)

Propynoic acid

Propynoic acid was again shown to be a fairly potent inhibitor of methanogenesis.
As pointed out in Chapter 3, the inhibition of CH; production must not have been caused
by competition for reducing equivalents because H2 accumulation was consistently
observed, and disappearance of propynoic acid was very low so that few reducing
equivalents would be used to completely reduce its triple bond (Table 6-7). The effects
on methanogenesis do not seem to be related to the amount of propynoic acid
metabolized.

Propynoic acid addition caused an increase in the accumulation of H2, formate,
and ethanol in Experiment 3. However, and in contrast with previous observations (see
Chapter 2), no accumulation of succinate was observed.

As it has been consistently reported, the inhibition of methanogenesis shifted the
VFA profile fi'om acetate to propionate (N agaraja et al., 1997). However, the responses
in butyrate production were different in Experiment 3 and 4. Propynoic acid decreased
butyrate production in Experiment 3 but increased it in Experiment 4. The increase in
butyrate production in Experiment 4 led to an increase (P < 0.01) in butyrate molar
percentage from 7.5 to 10.8% (data not shown). As the procedures used in both

experiments were the same, it is difficult to explain the opposite responses of butyrate

162

production to propynoic acid. A similar response was noted for isobutyrate production.
In Experiment 3, propynoic acid strongly decreased isobutyrate by more than 4-fold, but
it did not affected in Experiment 4. Both valerate and isovalerate production were
strongly decreased by propynoic acid in both experiments. Isobutyrate, valerate, and
isovalerate are fermentation products from valine, proline, and leucine, respectively
(Nagaraja et al., 1997), and their decrease could reflect an inhibition of the fermentation
of amino acids by propynoic acid. This agrees with the decline observed in N114+
concentrations in both experiments. Decreased fermentation of amino acids has been
reported when CH4 formation was inhibited with CO (Russell and Martin, 1984).
Deamination releases one pair of reducing equivalents per mole of NH4+ formed (V oet
and Voet, 1995). The increase in H2 partial pressure that occurs when methanogenesis is
inhibited, results in an inhibition of deamination, especially for highly reduced, branched-

chain amino acids (Hino and Russell, 1985).

163

Table 6-7. Proportion of the decrease in CI-L formation accountable by the complete

reduction of propynoic acid triple bond (Experiments 3 and 4)

 

Initial Second Pairs of 2H Decrease in CH4
Experiment concentration additive (mM) potentially taken accountable by

(mM) (umoles) hydrogenation (%)

 

Crotonic acid

3 q 2 OmM 33.4 3.14

 

Crotonic acid

3 2 4 mM 25.1 2.30

 

Crotonic acid

3 4 0 mM 54.4 3.73

 

Crotonic acid

 

 

 

 

3 4 4 mM 47.1 3.48
3-Butenoic

4 2 acid 0 mM 13.4 1.96
3-Butenoic

4 2 acid 4 mM 8.10 0.71
3-Butenoic

4 4 acid 0 mM 17.7 1.33
3-Butenoic

4 4 acid 4 mM 10.5 0.75

 

 

 

 

 

 

 

164

Ethyl 2-butynoate

Ethyl 2-butynoate was again shown to be a potent inhibitor of methanogenesis.
Carbon dioxide release was also decreased, as shown before (Chapter 3).

In both Experiment 5 and 6, and as observed before (Chapter 3), ethyl 2-
butynoate increased H2, formate, and ethanol accumulation; indicating that the electrons
spared from CH4 formation were not efficiently relocated into normal end products of
fermentation like propionate or butyrate.

In both Experiment 5 and 6, the acetate to propionate ratio was decreased (P <
0.01; quadratic response; data not shown), which is a normal consequence of the
inhibition of ruminal methanogenesis (N agaraja et al., 1997). Butyrate production was
increased by ethyl 2-butynoate in both experiments, acting a H sink for part of the
reducing equivalents spared from CH4 formation. In agreement with previous results
(Chapter 3), isobutyrate production was greatly increased in both experiments. This
could be a consequence of a greater deamination of valine (N agaraja et al., 1997).
However, NH4+ concentration was decreased in Experiment 5, and this may suggest a
general decrease in amino acid fermentation. Similarly, valerate and isovalerate, which
are products of proline and leucine catabolism, respectively (N agaraja et al., 1997), were
decreased. Alternatively, isobutyrate could be a product of ethyl 2-butynoate
metabolism. The C in extra isobutyrate represented 6.5 and 20.7% of ethyl 2-butynoate
C in Experiment 6, where all the additive disappeared. In Experiment 5, it represented
16.3 and 29.7% of C in ethyl 2-butynoate. Then, isobutyrate increase could have been

the result of ethyl 2-butynoate catabolism.

165

The accumulation of formate and ethanol when methanogenesis was inhibited by
ethyl 2-butynoate was considerably greater than with propynoic acid. On the contrary,
more H2 accumulated with propynoic acid than with ethyl 2-butynoate. As almost all of
ethyl 2-butynoate, but little propynoic acid, was metabolized by the mixed ruminal
rnicrobiota, it is possible that formate and/or ethanol were products of ethyl 2-butynoate

catabolism.

Crotonic acid

As observed in Chapter 3, crotonic acid caused a mild decrease in CH4 formation
in Experiments 1 and 3, although no effect was observed in Experiment 5. It was
hypothesized that crotonic acid would serve as an electron acceptor. This would
rechannel electrons spared from methanogenesis into butyrate formation, and decrease
the accumulation of reduced products without a nutritional value, such as H2, formate,
and ethanol. In Experiment 1, the addition of 656 umoles of crotonate resulted in only
142 umoles of extra butyrate (22%). It was thought that decreasing the initial
concentration of crotonic acid to 4 mM could increase its efficiency of conversion into
butyrate. However, in Experiment 3, crotonic acid did not relieve the increase in H2
accumulation or ethanol production caused by propynoic acid and caused a small
decrease in formate accumulation at 4 mM propynoic acid. Part of crotonic acid was
converted to butyrate as hypothesized, and smaller amounts were converted to acetate.
Averaged across propynoic acid concentrations, the addition of 328 umoles of crotonic

acid resulted in an increase in butyrate production of 95 moles (29%). The reduction of

crotonate to butyrate would imply an uptake of 95 umoles reducing equivalent pairs

166

(Miller and Jenesel, 1979). The increase in acetate production, 36 umoles, would imply a
release of 18 umoles of reducing equivalent pairs (Miller and Jenesel, 1979; Russell,
2002), therefore not compensating for the metabolic H uptake needed for butyrate 1
formation. As the effects on other VFA were minor, it is not clear why in Experiment 3
the uptake of metabolic H required for the conversion of crotonic acid into butyrate did
not result in greater decreases of fermentation end products without a nutritional value.

In Experiment 5, crotonic acid caused some decrease in H2 accumulation and
ethanol production. Forty percent of added crotonic acid seemed to be converted to
butyrate, while smaller amounts seemed to be converted to acetate. There was an 82
pmoles increase in butyrate production, a numerical increase in acetate of 36 umoles, and
a numerical decrease in propionate of 40 umoles. Effects on other VFA were minor.

The increase in acetate would imply a release of 18 umoles reducing equivalents pairs,
and the decrease in propionate formation would mean that 80 moles reducing equivalent
pairs were not taken up by this pathway (Russell, 2002). Changes in acetate and
propionate, when considered together, could compensate for the 82 umoles reducing
equivalents pairs incorporated into butyrate production. This could explain why the

decrease in unusual reduced end products of fermentation was not more pronounced.

3-Butenoic acid
There was no accumulation of unusual fermentation products in Experiment 2. In
Experiment 4, 3-butenoic acid did not decrease H2 accumulation caused by propynoic

acid, and, unexpectedly, increased formate accumulation. 3-Butenoic acid increased

167

butyrate production by 133 umoles, but also increased acetate production by 131 umoles.
Effects on other VFA were minor. Increased acetate production would imply a release of
66 umoles of reducing equivalent pairs, while the increase in butyrate would need 133
umoles reducing equivalent pairs (Russell, 2002). In total, these changes would imply an
uptake of approximately 67 moles of reducing equivalents pairs. However, there was
no effect on unusual reduced end products. In Experiment 6, 3-butenoic acid was again
ineffective in decreasing H2, formate, or ethanol. The increase in acetate production (145
umoles) was slightly higher than the corresponding increase in butyrate (116 umoles).
These changes, together with a tendency (P = 0.06) to decrease propionate production by
19 umoles, would result in almost no change caused by 3-butenoic acid on the H balance
(145 umoles/Z - (-19 umoles x 2 — 116 umoles = -5.5 umoles). This would explain the
lack of effect of 3-butenoic acid in capturing reducing equivalents, and withdrawing them
from CH4 formation, or H2, formate or ethanol accumulation.

The lack of effect of crotonic acid and 3-butenoic acid on decreasing the
accumulation of H2, formate and ethanol can be explained by: a) incomplete
disappearance of the additives, and b) alternative metabolic pathways. With the
exception of Experiment 6, additive disappearance was not measured due to additives co-
eluting from the HPLC column with VFA. Significant amounts of both crotonic acid and
3-butenoic acid were metabolized to acetate. Likely, this follows the reversal of butyrate
formation in the rumen, leading tothe formation of two moles of acetyl CoA (Miller and
Jenesel, 1979), and, eventually, acetate. Metabolism of intermediates of fermentation by
the reverse of their normal pathways has been observed before (Chapter 3). The reason

for a compound being metabolized in an opposite direction than normal can be

168

understood on thermodynamic grounds. A large increase in the concentration of a
metabolic intermediate normally found at very low concentrations would make
alternative metabolic pathways thermodynamically feasible, including the reverse of the
normal pathway. It follows that to maximize the efficiency of conversion of crotonic
acid or 3-butenoic acid into butyrate, these compounds should be added to the

fermentation continuously, avoiding large increases in their concentrations.

Conclusions

' Propynoic acid and ethyl 2-butynoate were again shown to be potent inhibitors of
CH4 production. Inhibition of methanogenesis resulted in inefficient relocation of
electrons into H2, formate, and ethanol. Neither crotonic acid nor 3-butenoic acid were
effective in re-channeling the electrons spared from methanogenesis into butyrate
formation. This was partly due to the fact that significant amounts of both additives were
converted to acetate, a process that releases reducing equivalents. The relatively high
concentrations of the additives may have made this alternative pathway
thermodynamically feasible. Possibly, a more continuous supply of these additives
would increase the proportion converted to butyrate and improve the uptake of reducing
equivalents.

The decreases in methanogenesis caused by lumazine were lower than previously

reported. The changes in the VFA profile were also erratic. The inconsistent effects of
lumazine across experiments on in vitro ruminal fermentation are difficult to explain.

The effects of lumazine on pure cultures of methanogens and other ruminal

169

microorganisms would need to be evaluated in order to understand its variable effects on

mixed ruminal cultures.

170

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Callaway, T. R. and S. A. Martin. 1996. Effects of organic acid and monensin treatment

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Chaney, A. L. and E. P. Marbach. 1962. Modified reagents for determination of urea and
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Hungate, R. E., W. Smith, and T. Bauchop. 1970. Formate as an intermediate in the
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Miller, T. L. and S. E. Jenesel. 1979. Enzymology of butyrate formation by Butyrivibrio
fibrisolvens. J. Bacteriol. 138: 99-104.

Nagaraja, T. G., C. J. Newbold, C. J. Van Nevel, and D. I. Demeyer. 1997. Manipulation
of ruminal fermentation. In: P. N. Hobson and C. S. Stewart (eds.) The Rurnen
Microbial Ecosystem. p 523-632. Blackie Academic and Professional, London.

Newbold, C. J ., J. O. Ouda, S. Lopez, N. Nelson, H. Omed, R. J. Wallace, and A. R.
Moss. 2001. Propionate precursors as possible electron acceptors to methane in
ruminal fermentation. In: Greenhouse Gases and Animal Agriculture, Obihiro,
Hokkaido, Japan. p 272-275.

171

Rao, P. V. 1998. Statistical Research Methods in the Life Sciences. lst ed. Duxbury
Press, Pacific Grove, CA.

Russell, J. B. 2002. Rurnen Microbiology and Its Role in Ruminant Nutrition. lst ed,
Ithaca, NY.

Russell, J. B. and S. A. Martin. 1984. Effects of various methane inhibitors on the
fermentation of amino acids by mixed rumen microorganisms in vitro. J. Anim.
Sci. 59: 1329-1338.

Russell, J. B. and R. J. Wallace. 1997. Energy-yielding and energy-consuming reactions.
In: P. N. Hobson and C. S. Stewart (eds.) The Rurnen Microbial Ecosystem. p
246-282. Blackie Academic and Professional, London.

Ungerfeld, E. M., S. R. Rust, M. K. Jain, and R. Burnett. 2000. Novel approaches for

‘ inhibiting rumen methanogenesis. In: Conference on Rumen Function, Chicago,
IL. p 19.

Ungerfeld, E. M., S. R. Rust, M. K. Jain, and R. Burnett. 2002. Some miscellaneous
inhibitors of rumen methanogenesis in vitro. Beef Cattle, Sheep and Forage
Systems. Res. Dem. Rep. 113-122.

Voet, D. and J. G. Voet. 1995. Biochemistry. 2nd ed. John Wiley and Sons, Inc., New
York.

172

CHAPTER 7
Effects of combinations of inhibitors of methanogenesis with crotonic acid or 3-

butenoic acid on in vitro degradation and microbial biomass and N synthesis

Abstract

It was hypothesized that the combinations of the methanogenesis inhibitors
lumazine, propynoic acid, and ethyl 2-butynoate, with crotonic acid or 3-butenoic acid
would be able to relieve the fermentation constraints observed when the inhibitors of
methanogenesis are used. In six experiments, one of the three methanogenesis inhibitors,
at three different concentrations, were combined with either crotonic acid or 3-butenoic
acid, at two different concentrations. A 4:1 buffer to ruminal fluid mixture was incubated
with grass hay in 1 L Erlenmeyer flasks for 72 h. 15N incorporation was used as a
microbial marker. All three methanogenesis inhibitors decreased N degradation.
Propynoic acid and ethyl 2-butynoate at the highest concentration, also decreased OM
degradation. Crotonic acid and 3-butenoic acid were ineffective to increase degradation
of OM or N. All three inhibitors increased the production of microbial OM and N.
Propynoic acid increased the amount of microbial OM and N per kilogram of total OM
disappeared. Propynoic acid disappearance was very small. Ethyl 2-butynoate, and
perhaps lumazine, could have been used as C and energy sources. Propynoic acid
increased the proportion of total disappeared OM (substrate plus additive) that was
incorporated into microbial biomass. It might have increased the efficiencies of

microbial OM and N synthesis by decreasing bacterial cell lysis. It was concluded that

173

lumazine and ethyl 2-butynoate at low concentrations could benefit animal production by

lowering CH4 formation, decreasing proteolysis, and increasing microbial N flow.

Introduction

Methane is formed in the nrmen as an end product of fermentation and is removed
by eructation. Its release represents a loss of digestible energy for ruminants. Methane is
also a potent greenhouse gas that contributes to global warming (Moss, 1993). There
would be environmental and economic benefits in decreasing CH4 production in the
rumen. In previous work, lumazine, propynoic acid, and ethyl 2-butynoate were shown
to inhibit ruminal methanogenesis (Ungerfeld et al., 2000, 2002). Unfortunately,
apparent OM fermented, as estimated through a mass balance, was decreased.

In contrast, crotonic acid and 3-butenoic acid had little effects on methanogenesis,
but seemed to stimulate OM fermentation (Ungerfeld et al., 2000). The objectives of this
series of six experiments were to study the effects of combinations of lumazine,
prOpynoic acid, and ethyl 2-butynoate with crotonic acid or 3-butenoic acid on mixed
ruminal cultures. It was hypothesized that the combinations of inhibitors of CH4
formation with crotonic acid or 3-butenoic acid would be able to lift the constraints on

fermentation while maintaining the inhibition of methanogenesis.

174

Materials and Methods
Experiment I

The experimental treatments of the 3 x 2 factorial arrangement were: 1) Lumazine .
(Sigma L 0380 for all experiments) 0 mM, crotonic acid (Sigma C4630 for all
experiments) 0 mM (double control); 2) Lumazine 0 mM, crotonic acid 8 mM; 3)
Lumazine 0.3 mM, crotonic acid 0 mM; 4) Lumazine 0.3 mM, crotonic acid 8 mM; 5)
Lumazine 0.6 mM, crotonic acid 0 mM; 6) Lumazine 0.6 mM, crotonic acid 8 mM.
Ruminal fluid was collected prior to the morning feeding from two non-lactating Holstein
cows fed grass hay. A mixture of 150 mL of ruminal fluid and 600 mL of buffer
(Goering and Van Soest, 1975) was anaerobically delivered into 1 L-Erlenmeyer flasks.
Each flask had 6 g of ground (1 mm screen mesh) grass hay (1.0% N, 67.1% NDF, 6.3%
ash, DM basis). Lumazine was added as a solid (4 and 8 mg for 0.3 and 0.6 mM,
respectively), while crotonic acid was added as 10 mL of a 0.308 M solution. Crotonic
acid controls received 10 mL of deionized water. Three samples of the ruminal fluid and
buffer mixture were frozen for subsequent determination of VFA initial concentrations.
Flasks were sealed under an O2-free CO2 atmosphere with stoppers that allowed gas
release but not entrance, and incubated in a non-shaking water bath at 39 °C for 72 h. An
aliquot of 5 mL was taken every 24 h for subsequent determination of crotonic acid
concentrations. Crotonic acid was quantified by differential refiactometry with a Waters
712 Wisp HPLC (Waters Associates Inc., Milford, MA) equipped with a BioRad HPX
87H column (BioRad Laboratories, Hercules, CA). Solvent was 0.005 M H2SO4 at 0.6

mL/min. Column temperature was 45 °C. Sample injection volume was 15 uL.

175

At the end of the incubation period, flasks were opened, and pH measured under
CO2 (Digital Benchtop pH Meter, Cole-Partner Instrument Company, Vernon Hills, IL).
Ten milliliters of 10% (v/v) H2SO4 were added to stop the fermentation. Flasks were
immediately refrigerated until further analysis.

Each flask content was removed and centrifuged at 500 x g at 4 °C for 15 min
(Zinn and Owens, 1986). The supematants were removed using a Pasteur pipette, and the
pellets of undigested substrate with attached microbial biomass (“pellets A”) transferred
into pre-weighted aluminum pans using an 0.85% saline solution. Supematants were
centrifiiged at 4,640 x g at 4 °C for 30 min (Hsu and Fahey, 1990). The supernatant from
the second centrifugation was removed using a Pasteur pipette and discarded. The
resulting pure microbial pellets (“pellets B”) were re-suspended in an 0.85% saline
solution and re-centrifuged at 4,640 x g at 4 °C for 30 min. Pellets suspended in saline
were transferred into pre-weighted aluminum pans. Pans containing both low- and high-
speed centrifugation pellets (A & B) were fieeze-dried for 5 d (Tri-Philizer MP, F TS
Kynetics, Stone Ridge, NY). Pellet masses were determined from difference in pans
masses. Pellets were analyzed for ash (Van Soest and Robertson, 1985), total N by
generic combustion (AOAC, 1990) in a FP-2000 analyzer (Leco®, St. Joseph, MI), and
'5N enrichment (Europa Integra mass spectrometer, PDZ Europa, Northwich Cheshire,
UK; Stable Isotope Facility, University of California-Davis). Pellets A were also
analyzed for NDF content (Van Soest et al., 1991). The three samples taken for initial
VFA content were combined into one, and centrifuged at 4,640 x g at 4 °C for 30 min

(Hsu and Fahey, 1990). The 15N enrichment of the resulting pellet was defined as 15N

176

natural abundance. The proportion of microbial N in the low-speed centrifugation pellet

A (PMNA) was calculated as:

PMNA = (”N% in pellet A — natural abundance) / (”N in pellet B - natural abundance)

Total microbial N was calculated as the addition of the microbial N in pellets A
and B. Microbial OM in pellet A was calculated using the N/OM ratio of pellet B, and
total microbial OM produced was calculated as the sum of microbial OM in pellets A and
B. Microbial C was calculated from the empirical formula C6H9.3502,99N12o (Stanier and
Davies, 1981). True degradation of N (Nm) and OM (OMTD) , and apparent NDF

degradation (NDFAD), were calculated from their disappearances during fermentation:

Nm (%) = 100 x (substrate N - N in pellet A + microbial N in pellet A) / substrate N
OMTD (%) =
100 x (substrate OM - OM in pellet A + microbial OM in pellet A) / substrate OM

NDFAD (%) = 100 x (substrate NDF - NDF in pellet A) / substrate NDF

Efficiencies of microbial N and OM synthesis were expressed on a truly degraded
substrate OM basis.

Degraded OM can either form fermentation products (VF A, gases, NH], etc) or
be incorporated into microbial OM (Leng and Nolan, 1984). The proportion of total
disappeared OM (substrate plus additives) incorporated into microbial OM (PMOM), was

calculated as:

177

PMOM = microbial OM / (substrate degraded OM + OM disappeared from additives)

Three replicates per compound and concentration were used. The experimental
model was: observation = overall mean + linear effect of lumazine + quadratic effect of
lumazine + crotonic acid effect + crotonic acid by linear effect of lumazine + crotonic
acid by quadratic effect of lumazine + residual. Non-significant (P > 0.15) quadratic
terms were removed from the model. When significant (P < 0.05) interactions or
tendencies (P < 0.15) were found, means were compared by Fisher least square
differences (Rao, 1998). Significance was declared at P < 0.05.

Crotonic acid disappearance as a function of time was modeled as a logistic
response (Neter et al., 1996) for each lumazine concentration:

%disappeared = 100 X exp(|30 + 510 / [1 + expwo + [311)]

where t is time in hours.

Experiment 2

The experimental treatments of the 3 x 2 factorial arrangement were: 1) Lumazine
0 mM, 3-butenoic acid (Acros 15883 for all experiments) 0 mM (double control); 2)
Lumazine 0 mM, 3-butenoic acid 4 mM; 3) Lumazine 0.6 mM, 3-butenoic acid 0 mM; 4)
Lumazine 0.6 mM, 3-butenoic acid 4 mM; 5) Lumazine 1.2 mM, 3-butenoic acid 0 mM;
6) Lumazine 1.2 mM, 3-butenoic acid 4 mM. Ruminal fluid was collected, processed,
and incubated, as described for Experiment 1. Lumazine was added as a solid (8 and 16

mg for 0.6 and 1.2 mM, respectively). 3-Butenoic acid was added as 10 mL of a 0.308 M

178

solution. 3-Butenoic acid controls received 10 mL of deionized water. Analytical and

statistical procedures were similar to Experiment 1.

Experiment 3

The 3 x 2 factorial arrangement of treatments included: 1) Propynoic acid (Acros
13150 for all experiments) 0 mM, crotonic acid 0 mM (double control); 2) Propynoic
acid 0 mM, crotonic acid 4 mM; 3) Propynoic acid 2 mM, crotonic acid 0 mM; 4)
Propynoic acid 2 mM, crotonic acid 4 mM; 5) Propynoic acid 4 mM, crotonic acid 0 mM;
6) Propynoic acid 4 mM, crotonic acid 4 mM. Ruminal fluid was collected, processed,
and incubated, as described for Experiment 1. Propynoic acid and crotonic acid were
added as 0.156 (10 or 20 mL) and 0.312 M (10 mL) solutions, respectively. Volumes
were equalized with deionized water. Analytical and statistical procedures were the same
as in Experiment 1, except that sampling was only done at the end of the 72 h-period.
Efficiencies of microbial N and OM synthesis were expressed both on a truly degraded
substrate OM basis (substrate only) and on a truly degraded total OM (substrate plus
additive) basis.

Propynoic acid disappearance after 72 h of incubation was modeled as a response
to propynoic acid (2 or 4 mM) and crotonic acid (0 or 4 mM) concentration, and their
interaction. Crotonic acid disappearance was modeled as a quadratic response to

propynoic acid concentration.

179

Experiment 4

The experimental treatments of the 3 x 2 factorial arrangement were: 1) Propynoic
acid 0 mM, 3-butenoic acid 0 mM (double control); 2) Propynoic acid 0 mM, 3-butenoic
acid 4 mM; 3) Propynoic acid 2 mM, 3-butenoic acid 0 mM; 4) Propynoic acid 2 mM, 3-
butenoic acid 4 mM; 5) Propynoic acid 4 mM, 3-butenoic acid 0 mM; 6) Propynoic acid
4 mM, 3-butenoic acid 4 mM. Ruminal fluid was collected, processed, and incubated, as
described for Experiment 1. Propynoic acid and 3-butenoic acid were added as 0.156 (10
or 20 mL) and 0.312 M (10 mL) solutions, respectively. Volumes were equalized with
deionized water. Analytical and statistical procedures were the same as in Experiment 3.
Efficiencies of microbial N and OM synthesis were expressed both on a truly degraded
substrate OM basis (substrate only) and on a truly degraded total OM (substrate plus
additive) basis.Propynoic acid disappearance at 72 h was modeled as a response to
propynoic acid (2 or 4 mM) and 3-butenoic acid (0 or 4 mM) concentrations, and the

interaction.

Experiment 5

The 3 x 2 factorial arrangement included: 1) Ethyl 2-butynoate (GFS Chemicals
3132 for all experiments) 0 mM, crotonic acid 0 mM (double control); 2) Ethyl 2-
butynoate 0 mM, crotonic acid 4 mM; 3) Ethyl 2-butynoate 4 mM, crotonic acid 0 mM;
4) Ethyl 2-butynoate 4 mM, crotonic acid 4 mM; 5) Ethyl 2-butynoate 8 mM, crotonic
acid 0 mM; 6) Ethyl 2-butynoate 8 mM, crotonic acid 4 mM. Ruminal fluid was
collected, processed, and incubated, as described for Experiment 1. Ethyl 2-butynoate

was delivered as a liquid (0.35 and 0.70 mL for 4 and 8 mM, respectively), while crotonic

180

acid was added as 10 mL of a 0.304 M solution. Crotonic acid controls received equal
volumes of deionized water. Analytical and statistical procedures were the same as in
Experiment 1. Efficiencies of microbial N and OM synthesis were expressed both on a
truly degraded substrate OM basis (substrate only) and on a truly degraded total OM
(substrate plus additive) basis. As almost all ethyl 2-butynoate disappeared by 24 h, only
data for that time period are presented for ethyl 2-butynoate disappearance. Ethyl 2-
butynoate disappearance was modeled as a response to ethyl 2-butynoate (4 or 8 mM)
and crotonic acid (0 or 4 mM) concentrations and the resulting interaction. Crotonic acid
disappearance as a function of time was modeled as a logistic response (N eter et al.,
1996) for each ethyl 2-butynoate concentration:

%disappeared = 100 x exp(Bo + Blt) / [1 + exp(Bo + 1311)]
where t is time in hours. Comparisons of crotonic acid disappearance among ethyl 2-

butynoate concentrations at 24 h were done using the Tukey test (N eter et al., 1996).

Experiment 6

The experimental treatments of the 3 x 2 factorial arrangement included: 1) Ethyl
2-butynoate 0 mM, 3-butenoic acid 0 mM (double control); 2) Ethyl 2-butynoate 0 mM,
3-butenoic acid 4 mM; 3) Ethyl 2-butynoate 4 mM, 3-butenoic acid 0 mM; 4) Ethyl 2-
butynoate 4 mM, 3-butenoic acid 4 mM; 5) Ethyl 2—butynoate 8 mM, 3-butenoic acid 0
mM; 6) Ethyl 2-butynoate 8 mM, 3-butenoic acid 4 mM. Ruminal fluid was collected,
processed, and incubated, as described for Experiment 1. Ethyl 2-butynoate was
delivered as a liquid (0.35 and 0.70 mL for 4 and 8 mM, respectively), while 3-butenoic

acid was added as 10 mL of a 0.304 M solution. 3-Butenoic acid controls received equal

181

volumes of deionized water. Analytical and statistical procedures were the same as in
Experiment 1. Efficiencies of microbial N and OM synthesis were expressed 'on a truly
degraded substrate OM basis (substrate only) and on a truly degraded total OM (substrate -
plus additive) basis. As almost all ethyl 2-butynoate disappeared by 24 h, only data for
that time period are presented for ethyl 2-butynoate disappearance. Ethyl 2-butynoate
disappearance was modeled as a response to ethyl 2-butynoate (4 or 8 mM) and crotonic
acid (0 or 4 mM) concentrations and the resulting interaction. 3-Butenoic acid
disappearance as a fimction of time was modeled as a logistic response (Neter et al.,
1996) for each ethyl 2-butynoate concentration:

%disappeared = 100 x exp(Bo + 1510/ [1 + exp(Bo + [310]

where t is time in hours.

Results
Experiment 1

Over 90% of crotonic acid disappeared after 48 h of incubation. However,
substantially less crotonic acid had disappeared after 24 h of incubation, especially at 0.3

mM lumazine (Figure 7-1).

182

 

 

Disappearance (%)

‘oc Oliumazine
"s. 0.3 mM lumazine
o 10 20 30 4o 50 60 7o 80 "*-. 0-6 liumazine

 

 

 

 

Figure 7-1 Crotonic acid disappearance (Experiment 1)

There were no interactions for OM degradation, and none of the additives affected
it. There was a tendency for a linear by quadratic interaction (P = 0.13) for N
degradation. Lumazine tended (P = 0.06; quadratic response; Table 7-1) to decrease N
degradation. At 0 mM crotonic acid, N degradation was decreased (P < 0.05) by 0.3 mM
lumazine, while at 4 mM crotonic acid, N degradation was decreased (P < 0.05) by 0.6
mM lumazine. Crotonic acid decreased (P < 0.05) N degradation at 0.6 mM lumazine.
There were no interactions for apparent NDF degradation, and none of the additives
affected it. There were linear by linear and linear by quadratic interactions (P = 0.04) for
the proportion of OM incorporated into microbial biomass. In the absence of crotonic
acid, lumazine at 0.3 mM increased (P < 0.05) the proportion of OM incorporated into
microbial biomass, while there was no effect of lumazine at 8 mM crotonic acid. There
were tendencies for linear by linear (P = 0.06) and linear by quadratic (P = 0.09)

interactions for microbial OM production. At 0 mM crotonic acid, lumazine at 0.3 and

183

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was higher (P < 0.05) at 0.6 than at 0.3 mM lumazine. Crotonic acid tended (P = 0.09) to
increase production of microbial OM. There were no interactions for the production of
microbial N, which was increased both by lumazine (P < 0.01) and crotonic acid (P <
0.05). There were linear by linear and linear by quadratic (P = 0.03) interactions for the
efficiency of microbial OM synthesis. At 0 mM crotonic acid, lumazine at 0.3 mM
increased (P < 0.05) the efficiency of microbial OM synthesis, but there was no effect of
lumazine at 8 mM of crotonic acid. There were no interactions for the efficiency of
microbial N synthesis, which was increased (P = 0.02) by lumazine, and not affected by
crotonic acid. No interactions were present for final pH, which was decreased (P < 0.01)

by crotonic acid, and not affected by lumazine (Table,7-1).

Experiment 2
Approximately one third of 3-butenoic acid disappeared by 24 h of incubation
across all lumazine concentrations. 3-Butenoic acid completely disappeared by 48 h of

incubation at all lumazine concentrations (Figure 7-2).

185

 

 

 

 

 

 

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There were no interactions for OM, N and NDF degradation. Lumazine did not
affect OM degradation (Table 7-2). At 1.2 mM, lumazine decreased N degradation (P =
0.02; quadratic response). In contrast, 3-butenoic acid increased (P = 0.03) N
degradation, although the increase was not enough to compensate for the decrease caused
by lumazine at 1.2 mM. Apparent NDF degradation was not influenced by either
additive. There were no interactions for the proportion of total OM incorporated into
microbial biomass, which tended (P < 0.10) to increase with added lumazine. There was
a tendency (P = 0.14) for a linear by quadratic interaction for microbial OM production.
Lmnazine increased the production of microbial OM (P = 0.01). 3-Butenoic acid
increased (P < 0.05) microbial OM production only at 1.2 mM lumazine. There were no
interactions for microbial N production. Lumazine increased (P < 0.01; quadratic

response) microbial N production, and 3-butenoic acid did not affect it. There were no

186

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interactions for the efficiencies of synthesis of microbial OM and N, both of which were
increased by lumazine and not affected by 3-butenoic acid.
There were no interactions for final pH. 3-Butenoic acid decreased (P < 0.01) the

final pH, while lumazine had no effect (Table 7-2).

Experiment 3

At 0 mM propynoic acid, all of crotonic acid disappeared by the end of the
incubation period, but only 63 and 69% disappeared at 2 and 4 mM propynoic acid,
respectively (P < 0.01; quadratic response). Disappearance of propynoic acid at the end
of the incubation period was lower than 10% for all treatments. There was a tendency (P
= 0.12) for an interaction for propynoic disappearance. At 4 mM crotonic acid, it was
higher (P < 0.05) with 2 mM propynoic acid than with 4 mM, while there was no
difference at 0 mM crotonic acid.

There were no interactions for OM degradation. Propynoic acid decreased (P <

0.01; quadratic response) OM degradation from 37.6 to 25.1%, while crotonic acid had

188

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189

no effect (Table 7-3). There were linear by linear and linear by quadratic interactions (P
< 0.01) on N degradation. Propynoic acid decreased (P < 0.01) N degradation to almost
zero. Crotonic acid decreased (P < 0.05) N degradation only at 0 mM propynoic acid.
There was a tendency (P = 0.07) for a linear by linear interaction for NDF degradation.
Propynoic acid decreased (P < 0.01; quadratic response) NDF degradation. Crotonic acid
decreased (P < 0.05) NDF degradation only at 0 mM propynoic acid. There were no
interactions for the proportion of OM incorporated into microbial biomass. Propynoic
acid increased (P < 0.01; quadratic response) the proportion of OM incorporated into
microbial biomass, and crotonic acid had no effect. There were no interactions for
microbial OM production, which was increased by both additives. There was a tendency
(P = 0.10) for a linear by linear interaction for microbial N production. Propynoic acid
increased (P < 0.01; quadratic response) microbial N production. Crotonic acid increased
(P < 0.05) microbial N production at 0 mM propynoic acid. There were no interactions
for microbial OM and N synthetic efficiencies. Propynoic acid improved (P < 0.01)
microbial OM and N synthetic efficiencies, while crotonic acid had no effect.

There were linear and linear by quadratic interactions (P < 0.01; Table 7-3) on
final pH. Propynoic acid did not affect final pH at 0 mM crotonic acid, but at 4 mM

crotonic acid pH was minimmn (P < 0.05) at 2 mM propynoic acid.

Experiment 4

Less than 20% propynoic acid disappeared by the end of the incubation period

(Table 7-4). Significant effects and interactions were of little biological importance.

190

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191

There were tendencies for linear (P = 0.06) and linear by quadratic (P = 0.08)
interactions for OM degradation. Propynoic acid decreased (P < 0.05) OM degradation
at 0, but not at 4 mM 3-butenoic acid (Table 7-4). 3—Butenoic acid increased (P < 0.05)
OM degradation only at 2 mM propynoic acid. Propynoic acid decreased (P = 0.04) N
degradation, while 3-butenoic acid did not affect it. There were tendencies for linear by
linear (P = 0.10) and linear by quadratic (P = 0.11) interactions for NDF degradation.
Propynoic acid decreased (P < 0.01) NDF degradation. 3-Butenoic acid increased (P <
0.05) NDF degradation only at 2 mM propynoic acid. There were linear by linear and
linear by quadratic interactions (P = 0.03) for the proportion of OM incorporated into
microbial biomass. Propynoic acid increased (P < 0.01; quadratic response) the
proportion of total OM incorporated into microbial biomass. At 2 mM propynoic acid, 3-
butenoic acid lowered (P < 0.05) , the proportion of OM incorporated into microbial
biomass. There was a tendency (P = 0.11) for an interaction for microbial OM
production. Propynoic acid increased (P < 0.01; quadratic response) microbial OM
production. Butenoic acid increased (P < 0.05) microbial OM production at 2 and 4 mM
propynoic acid. There were no interactions for microbial N production, which was
increased both by propynoic acid (P < 0.01; quadratic response) and 3-butenoic acid (P =
0.02). Propynoic acid increased (P < 0.01; quadratic response) the efficiency of
microbial OM synthesis calculated on a degraded substrate basis, while 3-butenoic acid
did not affect it. There were linear by linear and linear by quadratic interactions (P =
0.03) for the efficiency of microbial OM synthesis calculated on a total degraded OM
basis. Propynoic acid increased (P < 0.01; quadratic response) the efficiency of

microbial OM synthesis calculated on a total degraded OM basis, while 3-butenoic acid

192

decreased (P < 0.05) it only at 2 mM propynoic acid. Propynoic acid increased (P <
0.01; quadratic response) the microbial efliciency of N synthesis calculated on a
degraded substrate basis, while 3-butenoic acid did not affect it. There were linear by
linear (P = 0.01) and linear by quadratic (P = 0.02) interactions for the microbial
efficiency of N synthesis calculated on a total degraded OM basis. Propynoic acid
increased (P < 0.01; quadratic response) the microbial efficiency of N synthesis
calculated on a total degraded OM basis, while 3-butenoic acid decreased (P < 0.05) it
only at 2 mM propynoic acid.

There were no interactions for final pH. 3-Butenoic acid tended (P = 0.09) to

decrease final pH, while propynoic acid did not affect it (Table 7-4).

Experiment 5

More than 97% of ethyl 2-butynoate disappeared after 24 h of incubation. There
were no interactions for ethyl 2-butynoate disappearance. Disappearance was not
affected by its initial concentration or by the presence of crotonic acid (Table 7-5).
Virtually all crotonic acid disappeared after 48 h of incubation (Figure 7-3). However,
disappearance during the first 24 h was greater (P < 0.01) at 0 mM ethyl 2-butynoate.

There were no interactions for OM, N and NDF degradation. Ethyl 2-butynoate
at 8 mM decreased OM degradation from 36.0 to 30.6%. However, ethyl 2-butynoate at
4 mM did not impair OM degradation (P = 0.04; quadratic response; Table 7-5).
Crotonic acid did not affect OM degradation. Ethyl 2-butynoate strongly decreased (P <
0.05; quadratic response) N degradation from 47.9 to 19.5%, while there was no effect of

crotonic acid. The effect of ethyl 2-butynoate on apparent NDF degradation was

193

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194

quadratic, with a maximum at 4 mM. Crotonic acid tended (P = 0.11) to increase
apparent NDF degradation. There were no interactions for the proportion of OM
incorporated into microbial biomass, which was not affected by either additive. There
were no interactions for microbial OM and N production and the synthetic efficiencies.
Despite the decrease in OM degradation, ethyl 2-butynoate increased the production of
microbial OM (P = 0.02) and N (P < 0.01) through increased efficiencies of synthesis.
However, there was no improvement in the efficiencies when total OM disappearance

was accounted. Crotonic acid had no effect on the production of microbial OM and N, or

their efficiencies of synthesis.

 

 

Disappearance (%)

l \OmMEB
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Time (h)

Figure 7-3 Crotonic acid disappearance (Experiment 5)

There were no interactions for final pH. Ethyl 2-butynoate decreased (P < 0.01)

final pH, while crotonic acid did not affect it (Table 7-5).

195

Experiment 6

More than 95% of ethyl 2-butynoate disappeared after 24 h of incubation.
There were no interactions for ethyl 2-butynoate disappearance. Disappearance was not
affected by its initial concentration or by the presence of 3-butenoic acid (Table 7-6). All
of 3-butenoic acid had disappeared by 48 h of incubation. However, the disappearance of
3-butenoic acid’s was considerably smaller at 24 h incubation, especially at 8 mM ethyl

2-butynoate (Figure 7-4).

 

 

 

 

 

 

 

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There were no interactions for OM or N degradation. Ethyl 2-butynoate did not affect
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There was no effect of 3-butenoic acid upon OM or N degradation. NDF degradation and the
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treatments. Tendencies to increase the efficiencies of microbial OM (P = 0.08) and N (P = 0.06)

196

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197

synthesis calculated on a degraded substrate OM basis allowed ethyl 2-butynoate to increase (P <
0.01; quadratic response) the production of microbial OM and N production. However, there
was no improvement in synthetic efficiencies when total OM was accounted. 3-Butenoic acid
did not affect microbial OM and N production or their efficiencies of synthesis.

There were no interactions for final pH. Ethyl 2-butynoate tended (P = O. 12) to

decrease the final pH, while 3-butenoic acid had no effect (Table 7-6).

Discussion

Lumazine, propynoic acid, and ethyl 2-butynoate decreased N degradation.
However, their negative effects on OM and NDF degradation were milder or non-
existent. As NDF was the major non-nitrogenous component of the substrate OM, the
effects of the inhibitors on OM and NDF degradation were similar. Lumazine did not
affect OM or NDF degradation. In Experiment 5, ethyl 2-butynoate at 8 mM, but not at 4
mM, decreased OM and NDF degradation. In Experiment 6, ethyl 2-butynoate did not
decrease OM or NDF degradation at any concentration. Propynoic acid had the most
negative effects on OM and NDF degradation. However, in Experiment 3, the inhibition
exhorted by propynoic on OM and NDF degradation was lower than the inhibition of N
degradation.

It is unlikely that the decreased OM, N, or NDF degradation, could have been
caused by low pH. Although ethyl 2-butynoate and some propynoic acid combinations
decreased final pH, pH was not sufficiently reduced to impair degradation (Orskov and

Ryle, 1990).

198

As the additives affected more proteolysis than fibrolysis, it is possible that the
additives were more toxic to proteolytic than to fibrolytic microorganisms. Proteolytic
activity is quite widespread among ruminal bacteria, although the main cellulolytic
bacteria are possible exceptions (Wallace et al., 1997). Among cell-free extracts from 14
different entodiniomorphid protozoa, the cellulolytic species had the lowest proteolytic
activity (Coleman, 1983). Reports on the proteolytic activity of ruminal fibrolytic fimgi
are somewhat conflicting, although their role in proteolyisis in vivo is probably minor
(Wallace et al., 1997). Thus, there seems to be little overlapping between ruminal
proteolytic and cellulolytic microbial species, and it is possible that the inhibitors of
methanogenesis herein studied were more toxic to the former than the latter.

However, the fact that the three different compounds affected proteolysis more
than fibrolysis suggests that a more general explanation related to the inhibition of CH;
production could be possible. Propynoic acid and ethyl 2-butynoate increased the
accumulation of H2. It has been observed that H2 accumulation resulting from the
inhibition of CH4 formation decreased the redox potential of ruminal cultures (Sauer and
Teather, 1987). An increase in H2 pressure interferes with the interspecies H transfer and
the re-oxidation of cofactors (N agaraja et al., 1997). Deamination releases one mole of
reducing equivalent pairs per mole of NH: released (V oet and Voet, 1995). When
hydrogenase activity and subsequent CH4 formation were inhibited by CO, the
catabolism of amino acids was decreased. Deamination of branched-chain amino acids,
which are highly reduced, was particularly inhibited (Russell and Martin, 1984).

Furthermore, deamination of branched-chain amino acids was inhibited by NADH (Hino

199

and Russell, 1985). However, these results were not entirely reproducible with another
inhibitor of methanogenesis, chloroform (Russell and Martin, 1984).

Possibly, the inhibition of CH4 production had adverse consequences on
deamination, which could result in decreased proteolytic activity if amino acids
accumulated (Blackburn, 1968; Cotta and Hespell, 1986), and/or NH4+ concentration
decreased (Blackburn, 1968). Other work, however, did not find marked alterations in
proteolytic activity of ruminal bacteria caused by changes in the concentrations of NH],
amino acids or peptides (Blackburn, 1968), and it was concluded that proteases are
constitutive (Wallace and Brammall, 1985).

An inhibition of deamination is consistent with the decreased NI-L;+ concentration
observed with propynoic acid, ethyl 2-butynoate, and lumazine. Yet, responses in VFA
products of amino acid catabolism have not always been consistent with a decrease in
deamination (Chapters 3, 4, and 6). Lumazine decreased isovalerate, but caused large
increases in isobutyrate production. Isovalerate and isobutyrate are products of leucine
and valine catabolism, respectively (Nagaraja et al., 1997). Propynoic acid has been
shown to decrease the production of isobutyrate, valerate, and isovalerate, although,
strangely, the responses were sometimes quadratic. Valerate is a product of proline
catabolism (Nagaraja et al., 1997). Ethyl 2-butynoate decreased valerate and isovalerate,
but greatly increased isobutyrate production.

Another problem with this explanation for greater reduction in proteolysis than
fibrolysis, is that hexose fermentation, the same as deamination, releases metabolic H.
Glycolysis releases two moles of reducing equivalent pairs per mole of glucose fermented

(V oet and Voet, 1995), and more metabolic H is released when acetate is the end product

200

of fermentation. Then, a decrease in redox potential should inhibit both proteolysis and
cellulolysis.

Proteolytic activity can also be decreased by reducing agents as a result of
hydrogenation of cysteine residues involved in the catalytic mechanism (Wallace and
Brammall, 1985). Sodium sulfide, L—cysteine and dithiothreitol decreased proteolytic
activity by Butyrivibriofibrisolvens (Cotta and Hespell, 1986). Dithiothreitol also
decreased the proteolytic activity of both mixed ruminal fluid and monocultures of
ruminal bacterial species (Wallace and Brammall, 1985). However, other results in
which proteolytic activity of Prevotella (Bacteroides) ruminicola was increased by
dithiothreitol and cysteine have also been reported (Hazlewood et al., 1981).

Under most situations, an inhibition of ruminal proteolysis is not undesirable.
Protein fermentation in the rumen is generally considered wasteful (Russell, 1983). The
conversion of feed protein into microbial protein requires energy, part of dietary amino
acids are fermented, and NH: released is only partially captured into microbial protein
(Nagaraja et al., 1997), increasing N voided in the environment.

Crotonic acid was ineffective in improving OM, N, or NDF degradation. There
was some improvement in N degradation by supplementing lumazine at 0.6 mM with 3-
butenoic acid, but no consistent improvements in degradation of OM, N or NDF were
observed with 3-butenoic acid. Previous results had shown that these additives could
improve apparent OM fermentation (Chapter 3). A shift in degraded OM towards more
end more end products of fermentation and less microbial biomass may explain the
increase in apparent OM degradation with no change in true degradation. However, only

the combination of crotonic acid with lumazine at 0.3 mM, and 3-butenoic acid with

20]

propynoic acid at 2 mM, decreased the proportion of disappeared OM incorporated into
microbial biomass. Overall, crotonic acid only tended (P = 0.11) to decrease the
proportion of total OM (substrate plus additives) incorporated into microbial biomass in
Experiment 1, and did not affect it in Experiments 3 and 5. 3-Butenoic acid did not affect
the pr0portion of OM incorporated into microbial biomass in any of the experiments. It
is not clear therefore why crotonic acid and 3-butenoic acid stimulated apparent OM
degradation in previous experiments, but failed to improve true OM degradation in the
present experiments.

Nitrogen degradation in the absence of added additives ranged between 43.7 and
72.3% across experiments. The reasons for this ample variation are not apparent, as the
same source of ground hay was used in all the experiments. Differences in the
proteolytic activity of the ruminal fluid used as inoculum are unlikely as the incubations
were conducted for 72 h.

Unexpectedly, all of the inhibitors of CH4 formation improved the efficiencies of
production of microbial OM and N, expressed on a truly degraded substrate OM basis.
Previous reports of the effects of the inhibition of ruminal CH4 production on microbial
efficiency of N synthesis are scarce. No changes in the efficiency of microbial N
synthesis were found when CH4 production was inhibited with 2-trichloromethyl-4-
dichloromethylene benzo[1,3] dioxin 6-carboxylic acid or monensin (Stanier and Davies,
1981). Unfortunately, the efficiency of microbial N synthesis was expressed on an
apparent OM degradation basis in that study.

It is tempting to think that the improvements in microbial efficiency were a

general consequence of changes in H dynamics. The inhibition of CH4 formation

202

decreases the redox potential (Sauer and Teather, 1987), which could favor anabolic
processes that incorporate reducing equivalents, like amination and fatty acid synthesis
(V oet and Voet, 1995).

However, ethyl 2-butynoate did not improve the efficiencies of microbial OM and
N synthesis when the disappearance of OM from the additives was considered. It is
apparent therefore that ethyl 2-butynoate was used as a source of C and/or energy. Figure
7-5 shows that disappearance of C from ethyl 2-butynoate would be sufficient to explain
the increase in microbial C observed. In contrast, the potential supply of C from
disappeared propynoic acid would be largely insufficient to explain the increases

observed in microbial C.

0.16 T

 

0.12 0* ....... ....... .. ....... 2-- .......

0,03 .1. ....... ....... ....... .......
. ° Propynoic acid
004 9 ...... ....... ....... 2 mM

' ' ' 3 .i 0 Propynoic acid

5 5 = 5 ‘3 4 mM

0 t... ....... .....A.-. ....... .-... ....... A Ethyl 2 butynoate
9 4 mM

L. ‘ Ethyl 2-butynoate
0 0.05 0.1 0.15 0.2 0.25 0.3 0.35 0.4 0.45 0.5 0.55 8 mM

Increase in microbial C (9)
01° .

 

 

-0.04

 

C disappeared in additive (9)

Figure 7 -5 Relationship between C disappeared from additives and the increase in

microbial C synthesis

203

Propynoic acid increased microbial efficiency of OM and N synthesis without
being used as a C or energy source. It also increased the proportion of OM that was
incorporated into microbial biomass. Protozoa release a significant proportion of
digested bacterial material into the medium and increase VFA concentration (Williams
and Coleman, 1997). Propynoic acid has been shown to decrease VFA concentration
(Chapters 3 and 6). It is then possible that the shift in the partition of disappeared OM
towards microbial biomass and less fermentation products was caused by antiprotozoal
activity of propynoic acid. This would decrease bacterial predation and N turnover, and
increase the production of microbial N (Williams and Coleman, 1997).

An improvement in the efficiency of microbial N synthesis may also occur if
propynoic acid inhibited deamination more than proteolysis, as this would increase the
availability of amino acids. There is evidence that amino acids and peptides can
stimulate the efficiency of microbial N synthesis (Leng and Nolan, 1984), although this is
generally observed with rapidly degradable energy sources. It may not be true for energy
substrates that are degraded more slowly such as hay used in the present study (Wallace
et al., 1997).

A lumazine derivative, 6, 7-dimethyl-8-ribityllumazine, is a riboflavine precursor
(Weimar and Neims, 1975), and therefore an FAD precursor (V oet and Voet, 1995). If
lumazine is used to synthesize FAD, this may relieve some of the decrease in redox
potential that occurs when CH4 production is inhibited (Sauer and Teather, 1987). It
could be of interest to evaluate whether there are responses in degradation and microbial

biomass synthesis to riboflavine when methanogenesis is inhibited.

204

Conclusions

All the inhibitors of methanogenesis decreased N degradation. Organic matter
and NDF degradation were decreased by propynoic acid and ethyl 2-butynoate at 8 mM.
Possible reasons for a greater inhibition of N than OM and NDF degradation are: l) a
greater toxicity to proteolytic than cellulolytic microorganisms (as NDF comprised most
of non-nitrogenous OM); 2) inhibition of proteolysis due to less deamination resulting
from a lower redox potential; and/or 3) inhibition of proteolysis as a result of
hydrogenation of cysteine residues in catalytic sites. Neither crotonic acid nor 3-butenoic
acid were shown to consistently improve OM, N, or NDF degradation, and it does not
appear to be beneficial to use these acids to relieve the depression in degradation caused
by inhibitors of CH4 formation.

Decreases in CH4 production caused by lumazine have been erratic, and lower
than with other additives. However, lumazine did not impair OM or NDF degradation,
and it increased microbial N and OM production. It decreased substrate proteolysis,
which can decrease N flow into NH; in the rumen and reduce N release into the
environment.

Ethyl 2-butynoate at a concentration of 4 mM did not impair OM or NDF
degradation. Methane production has been shown to decrease by approximately 50% at 4
mM ethyl 2-butynoate (Chapter 6), although 6 mM caused only a 24% decrease (Chapter
3). The additive was totally fermented and increased microbial N production. It could
decrease N flow to NH4+ in the rumen, improve N retention by the animal, and decrease

N voided to the environment. Ethyl 2-butynoate was catabolized within 24 h of

205

fermentation, which would decrease the risks of toxicity for the animal or the
environment.

All three additives improved microbial synthetic efficiencies. Relocation of
electrons spared from CH4 formation may have favored anabolic processes such as
amino acids or fatty acid biosynthesis. Alternatively, ethyl 2-butynoate could have
improved synthetic efficiencies by being a C source. However, the increases in microbial
C caused by propynoic acid were greater than the C disappeared from this additive. The
shift in the partition of OM towards more microbial biomass and less fermentation
products could indicate a reduction in bacterial lysis. Changes in protozoal numbers
could be useful to understand the effects of propynoic acid on degradation and synthetic

efficiencies.

206

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Blackburn, T. H. 1968. Protease production by Bacteroides amylophilus strain H 18. J.
Gen. Microbiol. 53: 27-36.

Coleman, G. S. 1983. Hydrolysis of fraction 1 leaf protein and casein by rumen
entodiniomorphid protozoa. J. Appl. Bacteriol. 55 : 111-118.

Cotta, M. A. and R. B. Hespell. 1986. Proteolytic activity of the ruminal bacterium
Butyrivibriofibrisolvens. Appl. Environ. Microbiol. 52: 51-5 8.

Goering, H. K. and P. N. Van Soest. 1975. Forage Fiber Analyses (Apparatus, Reagents,
Procedures and some Applications). 379, ARS-USDA, Washington DC.

Hazlewood, G. P., G. A. Jones, and J. L. Mangan. 1981. Hydrolysis of leaf fiaction 1
protein by the proteolytic bacterium Bacteroides ruminicola R8/4. J. Gen.
Microbiol. 123: 223-232.

Hino, T. and J. B. Russell. 1985. Effect of reducing-equivalent disposal and NADH/NAD
on deamination of amino acids by intact rumen microorganisms and their cell
extracts. Appl. Environ. Microbiol. 50: 1368-1374.

Hsu, J. T. and G. C. Fahey, Jr. 1990. Effects of centrifugation speed and freezing on

composition of ruminal bacterial samples collected from defaunated sheep. J.
Dairy Sci. 73: 149-152.

Leng, R. A. and J. V. Nolan. 1984. Symposium: protein nutrition of the lactating dairy
cow. J. Dairy Sci. 67: 1072-1089.

Moss, A. R. 1993. Methane. Global Warming and Production by Animals. lst ed.
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Nagaraja, T. G., C. J. Newbold, C. J. Van Nevel, and D. I. Demeyer. 1997. Manipulation
of ruminal fermentation. In: P. N. Hobson and C. S. Stewart (eds.) The Rurnen
Microbial Ecosystem. p 523-632. Blackie Academic and Professional, London.

Neter, J ., M. H. Kutner, C. J. Nachtsheim, and W. Wasserman. 1996. Applied Linear
Statistical Models. 4th ed. McGraw-Hill, Boston.

Orskov, E. R. and M. Ryle. 1990. Energy Nutrition in Ruminants. lst ed. Elsevier
Science Publishers Ltd., Essex, UK.

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Russell, J. B. 1983. Fermentation of peptides by Bacteroides ruminicola Bl4. Appl.
Environ. Microbiol. 45: 1 566-1 574.

Russell, J. B. and S. A. Martin. 1984. Effects of various methane inhibitors on the
fermentation of amino acids by mixed rumen microorganisms in vitro. J. Anim.
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Sauer, F. D. and R. M. Teather. 1987. Changes in oxidation reduction potentials and
volatile fatty acid production by rumen bacteria when methane synthesis in
inhibited. J. Dairy Sci. 70: 1835-1840.

Stanier, G. and A. Davies. 1981. Effects of the antibiotic monensin and an inhibitor of

methanogenesis on in vitro continuous rumen fermentations. Br. J. Nutr. 45: 567-
578.

Ungerfeld, E. M., S. R. Rust, M. K. Jain, and R. Burnett. 2000. Novel approaches for
‘ inhibiting rumen methanogenesis. In: Conference on Rurnen Function, Chicago,
IL. p 19.

Ungerfeld, E. M., S. R. Rust, M. K. Jain, and R. Burnett. 2002. Some miscellaneous
inhibitors of rumen methanogenesis in vitro. Beef Cattle, Sheep and Forage
Systems. Res. Dem. Rep. 113-122.

Van Soest, P. N. and J. B. Robertson. 1985. Analysis of Forages and Fibrous Feeds,
Cornell University, Ithaca, NY.

Van Soest, P. N., J. B. Robertson, and B. A. Lewis. 1991. Methods for dietary fiber,
neutral detergent fiber and nonstarch polysaccharides in relation to animal
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Voet, D. and J. G. Voet. 1995. Biochemistry. 2nd ed. John Wiley and Sons, Inc., New
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Wallace, R. J. and M. L. Brammall. 1985. The role of different species of bacteria in the
hydrolysis of protein in the rumen. J. Gen. Microbiol. 131: 821-832.

Wallace, R. J ., R. Onodera, and M. A. Cotta. 1997. Metabolism of nitrogen-containing
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208

Williams, A. G. and G. S. Coleman. 1997. The rumen protozoa. In: P. N. Hobson and C.
S. Stewart (eds.) The Rumen Microbial Ecosystem. p 73-139. Blackie Academic
and Professional, London.

Zinn, R. A. and F. N. Owens. 1986. A rapid procedure for purine measurement and its
use for estimating net ruminal protein synthesis. Can. J. Anim. Sci. 66: 157-166.

209

CHAPTER 8

Effects of several inhibitors on pure cultures of ruminal methanogens

Abstract

The effects of five inhibitors of methanogenesis, 2-bromoethanesulfonate (BES),
3-bromopropanesulfonate (BPS), lumazine, propynoic acid, and ethyl 2-butynoate, on
CH4 production of ruminal methanogens Methanobrevibacter ruminantium,
Methanosarcina mazei, and Methanomicrobium mobile were examined. M. ruminantium
was the most sensitive species to BES, propynoic acid, and ethyl 2-butynoate. M. mazei
was the least sensitive species to those chemical additives, and M. mobile was
intermediate. Inhibition caused by propynoic acid and ethyl 2-butynoate appeared to be
non-competitive. BPS failed to inhibit any of the methanogens. All three species were
almost completely inhibited by 50- and 100%-lumazine saturated media, but the
inhibition was somewhat lower with a 25%-lumazine saturated media. There were
important differences among species of methanogens regarding their sensitivity to the
different inhibitors. The presence of resistant species of methanogens should be
considered when designing strategies of inhibition of ruminal methanogenesis. Long
term changes in the populations of ruminal methanogens caused by inhibitors of

methanogenesis need to be investigated.

Introduction

Methane formation in the rumen implies an energy loss for ruminants, and also

contributes to global warming. There is, therefore, an interest in decreasing CH4

210

production in the rumen (Moss, 1993). Chemical additives with different modes of
action have been used to decrease ruminal methanogenesis. Some compounds have been
shown to considerably decrease CH4 production of mixed nmiinal cultures (Van Nevel
and Demeyer, 1996; Nagaraja et al., 1997).

Although the main interest when evaluating potential inhibitors of
methanogenesis is their effects on the mixed ruminal microbial community, it is
important to know how individual species of methanogens are affected. If some species
are resistant to an inhibitor, its long term use in vivo could result in their selection, and
CH4 formation could return to the levels observed before the inhibitor was fed. For
example, 2-bromoethanesulfonate (BES) is a very potent inhibitor of methanogenesis in
batch cultures, but ruminal rnicrobiota of sheep adapted to the compound after BES was
administered to the animal for three days (Van Nevel and Demeyer, 1996). Also, the
relative proportion of methanogenic species can be influenced by ruminant species (Lin
et al., 1997), diet composition and management (Rowe et al., 1979; Jarvis et al., 2000),
and geographical area (V icini et al., 1987). This may result in different responses to
chemical additives used to inhibit CH4 production in the rumen. BES (Nagaraja et al.,
1997), 3-bromopropanesulfonate (BPS), lumazine, propynoic acid, and ethyl 2-butynoate
have been previously studied regarding their effects on CH4 production by mixed ruminal
cultures (see Chapters 3 and 4). The objectives of the present study were to investigate
the effects of these five chemicals on CH4 production of pure cultures of three ruminal
methanogens. It was hypothesized that there would be differences among methanogens

in their sensitivity to the inhibitors.

211

Materials and Methods
Cultures

The experiment was conducted at the Department of Biology, Portland State
University, Portland, OR, USA. Methanobrevibacter ruminantium Ml (OCM 146) and
Methanosarcina mazeii CW3A (OCM 20) were obtained from the Oregon Collection of
Methanogens (Department of Biology, Portland State University, Portland, OR 97207,
USA). Methanomicrobium mobile 1539 (DSM 1539) was obtained from DSMZ (DSMZ
Versand, 38124 Braunschweig, Germany). All three strains have been isolated from

ruminal fluid.

Medium preparation

All three cultures were grown in MS medium (Boone et al., 1989), which
contained per liter: 4.0 g NaOH, 2.0 g yeast extract (Difco Laboratories, Detroit, MI), 2.0
g Trypticase peptones (BBL Microbiology Systems, Cockeysville, MD), 1.0 g NH4Cl,
1.0 g MgC12-6H20, 0.2 g 2-mercaptoethanesulfonic acid (coenzyme M), 0.4 g
K2HP04-3HZO, 0.4 g CaC12-2H20, 5.0 mg disodium EDTA-ZHZO, 1.5 mg CoClz-6HzO,
1.0 mg resazurin, 1.0 mg of MnClz'4H20, 1.0 mg F eSO4-7HzO, 1.0 mg ZnClz, 0.4 mg
AlCl3-6HzO, 0.3 mg Na2WO4°2HzO, 0.2 mg CuClz-ZHZO, 0.2 mg NiSO4-6HzO, 0.1 mg
H28603, 0.1 mg of H3B03, and 0.1 mg NaMoO4-2HzO, 250 mg Nags-9H20, and 250 mg
L-cysteine . Twenty five percent (v/v) of deionized water was replaced with clarified
ruminal fluid (Leadbetter and Breznak, 1996). Methanol (30 mM final concentration)
was also included in the medium for M mazei (P01 and Demeyer, 1988). Medium was

anaerobically mixed and delivered under a 50:50 02-free Nz/COz gas mixture. pH of the

212

medium before delivery into tubes was 6.90. Five milliliters of medium were delivered
into 25 mL-Hungate tubes and hermetically sealed with butyl rubber septum and
aluminum seals (Bellco Glass Company, Vineland, NJ). Tubes were then autoclaved at
121 °C for 20 min. Approximately 24 h before inoculation, 0.05 mL of a previously
autoclaved, 02-free, 2.5% (w/v) NaZS-Hzo stock solution, were aseptically added to the

tubes through the butyl rubber septum.

Chemicals

Chemical inhibitors studied were BBS and BPS at 0, 10, 50, and 250 uM,
propynoic acid at O, 2 and 4 mM, ethyl 2-butynoate at O, 4 and 8 mM, and lumazine as a
O, 25, 50 or 100% lumazine-saturated medium. BBS [Na salt, Sigma B 9008], BPS [Na
salt, B2912], and propynoic acid [free acid, Acros 13150-0100] were dissolved in
deionized water, filter-sterilized, and 0.1 mL of each solution delivered into the
corresponding tubes. Ethyl 2-butynoate [Aldrich 4341-76-8] was directly injected into
the tubes through the butyl rubber septum using a 10 uL Hamilton syringe (2.35 and 4.7
uL of ethyl 2-butynoate to achieve 4 and 8 mM, respectively). Two milligrams of
lumazine [~12.2 umoles; Sigma L 03 80] were added to 50 mL of medium in a Wheaton
bottle, shaken, and left overnight. The presence of visible yellowish particles of lumazine
at the bottom of the bottle indicated that the medium was saturated with lumazine. The
lumazine-saturated medium was filter-sterilized, and 5, 2.5, and 1.25 mL were mixed
with O, 2.5, and 3.75 mL of previously filter sterilized medium, respectively, and injected

into sealed, previously autoclaved Hungate tubes, to achieve 100, 50, and 25% lumazine-

213

saturated medium. Lumazine solubility could not be measured because it was difficult to

establish the minimum saturating concentration.

Inoculation and incubation

Stock cultures were grown on MS medium (Boone et al., 1989) and 0.2 mL
delivered into autoclaved Hungate tubes with MS medium. Tubes were then pressurized
with O2-free H2 at 138 kPa. Tubes were then horizontally placed in a shaking incubator

at 37 °C at 120 rpm.

Measurements and calculations

Methanogen growth can be followed by CH4 production (Balch and Wolfe, 1976;
Balch et al., 1979). At 4 and 6 d of incubation, tubes were analyzed for CH4 production
using a flame ionization-detection GC (Chong-Song et al., 2002). A 10% CH; gas
mixture at 69 kPa was used as a standard. Micromoles of CH4 produced per tube were

calculated from its partial pressure and headspace volume.

Statistical analysis

The experimental model for each of the chemical additives was: CH4 production =
overall mean + species + concentration + day (4 or 6) + random effect of tube nested
within species and concentration + species x concentration + species x day +
concentration x day + species x concentration x day + residual. Concentration of BES,
BPS, and lumazine (% of saturation) was log transformed (euler base). Type III mean

squares for tube nested within species and concentration were used as the error term for

214

species and concentration effects and their interaction (N eter et al., 1996). Depending on
the significance of triple and double interactions, linear and quadratic contrasts for
concentration were evaluated separately for each species or each combination of species
by day. If both linear and quadratic contrasts were significant (P < 0.05), a quadratic
response was declared. Comparisons of interest between means of the same species and
chemical concentration on different days were done by the Scheffe test (N eter et al.,
1996). Due to heterogeneity of variances, the repeated! group statement of SAS was used
to estimate separate variances per concentration for BES, per species for lumazine, and

per combination of species and concentration for propynoic acid (SAS®, 1999).

Results and Discussion
BES

At 4 (1, CH4 production of M. ruminantium was decreased by BES at 10 uM and
was minimal at 50 and 250 uM (quadratic contrast P = 0.03; Table 8-1). By day six, M.
ruminantium had some recovery at 50 mM BES (P < 0.01; means, but not probability, are
shown in Table 8-1). In contrast, M mazei and M. mobile were minimally affected by
BES at 10 or 50 uM (species x concentration; P < 0.01). M. mobile showed minimal CH4
production with 250 uM BES (linear and quadratic contrast P < 0.01), while M mazei
was inhibited by 250 uM BES during the first 4 d, but showed a strong recovery by day
six P < 0.01; means, but not probability, are shown in Table 8-1). In agreement with the
current results, methano gens resistant to BES have been found in sheep that were never

fed the chemical (Ungerfeld, 1998).

215

Methanogens of the family Methanobacteriaceae, of which M. ruminantium is the
main ruminal species (Sharp et al., 1998), were dominant in a protozoa-enriched fraction
of ruminal fluid from a Holstein cow. On the contrary, the order Methanomicrobiales, of
which M mobile is the only ruminal species isolated, was negatively associated with the
presence of protozoa (Sharp et al., 1998). M. mobile was also the dominant methanogen
in sheep ruminal fluid in which few protozoa had ecto or endosymbiotic methanogens
(Yanagita et al., 2000). Higher resistance to BES was sometimes found in defaunated
compared to faunated ruminal fluid (Ungerfeld, 1998). This may be explained by a high
proportion of M mobile in total archaea in the absence of protozoa, as M mobile had a

greater resistance to BES compared to M ruminantium in the present experiment

Table 8- 1. Effects of 2-bromoethanesulfonate (BES) on CI-L production (umoles) of
three ruminal '

     
 

concentratron
res l
118 7 ll
ruminantium
< . 1
mazei .
. < 0.
mobile . . < . 1
< . < . 1 < . pecres x
Concentration; P < 0.01); Species by concentration (P < 0.01); Species by days
(P < 0.01); Concentration by days (P < 0.01); Species by concentration by days
(P < 0.01); SEM 0 uM = 0.32; SEM 10 uM = 1.24; SEM 50 uM = 1.74 uM;
SEM 250 uM = 1.31; 2Significance of linear contrast; 3Significance of quadratic
contrast; ‘ND = not detected.

   

BES is a structural analog of coenzyme M (CoM) and inhibits the reductive
demethylation of methyl-S-CoM in the last step of methanogenesis (Miiller et al., 1993).
Resistance to BES in the non-ruminal Methanococcus voltae (Santoro and Konisky,

1987) and a non-ruminal species of Methanosarcina (Smith, 1983) was shown to be

216

based on an inability to transport BES into the cell. Inhibition of M ruminatium (Balch
and Wolfe, 1979a) and a non-ruminal Methanosarcina (Smith and Mah, 1981) by BES
was lower when CoM concentration in the medium was elevated. Coenzyme M
prevented the uptake of BES and protected Methanococcus voltae cells from BES
inhibition (Santoro and Konisky, 1987). Methanogens that have the ability to synthesize
CoM are less dependent on CoM taken from the medium. In a medium containing 5 uM
CoM, M mobile and Methanobrevibacter smithii (M ruminantium PS) had 90% lower
CoM uptake than M ruminantium M1 (Balch and Wolfe, 1979b). Both M mobile and
M smithii can synthesize CoM intracellularly (Balch and Wolfe, 1979b; Stewart et al.,
1997). M ruminantium M1, in contrast, requires CoM in the medium, and was shown in
the current experiment to be highly sensitive to BES. M mazei, which was more resistant
to BES, has the ability to synthesize the CoM (Stewart et al., 1997). Methanogens that
can synthesize CoM exhibit lower rates of transport of external CoM into the cell and are
likely more resistant to BES. As the medium used in the present experiment contained
around 1,200 uM CoM, the inhibition of M mobile and M mazei might have been
greater with less or no added CoM.

M ruminantium is the only mminal methanogen isolated that requires CoM
(Stewart et al., 1997). The implications would be that feeding BES could result in a
selection of other methanogens that would occupy the empty niche left by M
ruminantium, causing BES-resistance of the mixed ruminal rnicrobiota as observed in
vivo (Van Nevel and Demeyer, 1996). The accumulation of H2 observed when ruminal
methanogenesis is inhibited (Van Nevel and Demeyer, 1996) could allow for faster

growth rates of the non-inhibited, BES-resistant methanogens.

217

In the present experiment, M mazei seemed to adapt to 250 uM BES between 4
and 6 d. In a previous report, BES at 150 uM inhibited methanol utilization in sheep
ruminal fluid 1 h after its addition, indicating that the chemical was toxic for
Methanosarcina at that concentration (Pol and Demeyer, 1988); however, it was not
reported whether the inhibition remained over a longer period. There are, nevertheless,
previous reports on spontaneous acquisition of resistance to BES in Methanosarcina spp.
(Smith and Mah, 1981). The non-ruminal methanogen Methanosarcina strain 227 was
inhibited by 24 uM BES, but spontaneously resumed growth after 6 d. It was shown that
the acquisition of resistance was based on heritable changes and was not due to biological
inactivation of BES, or inactivation through reaction with medium components.
Interestingly, BES resistance was retained even after 30 transfers in the absence of BES
(Smith and Mah, 1981). In the present experiment, M ruminantium also showed some
adaptation between 4 and 6 d.

It is apparent , therefore, that BES adaptation of mixed ruminal rnicrobiota (Van
Nevel and Demeyer, 1996) is based on at least two mechanisms: selection of species of
methanogens with greater resistance to BES, and selection of BES-resistant strains or
mutants within species. It is also possible that BES degradation by the mixed ruminal
rnicrobiota occurs. Reduction of BES sulfonic moiety to sulfide, presumably by sulfate-
reducing bacteria, has been reported for the microbial community of a river sediment (Ye

et al., 1999).

218

BPS

BPS did not affect CH4 production in any of the species studied (P = 0.17; Table
8-2). Previously, methanogenesis of a mixed ruminal culture was shown to be unaffected
by BPS (Ungerfeld et al., 2002). BPS, however, has been found to be a very potent
inhibitor of methyl-COM reductase in a pure enzyme-substrate system (Ellerman et al.,
1989). The opposite results observed with the purified enzyme-substrate system and with
pure methanogen and mixed cell cultures, suggest that BPS is not transported into cells.
BPS has a closer structure to methyl-COM than BES (Ellerman et al., 1989). Coenzyme
M and BES have been shown to share a common transport system (Smith, 1983; Santoro

and Konisky, 1987). Coenzyme M transport system may not be suitable for transporting
methyl-COM or BPS into the cell.

Table 8- 2. Effects of 3- bromopropanesulfonate (BPS) on CH4 production
(umoles of three ruminal

   
   

COI'ICCDII‘aUOD
ICS

ruminantium

res ( < .

Tube (Species x Concentration; P < O. 01); Species x Concentration (P= 0.19);
Species x Days (P < 0. 01); Days x Concentration (P= 0. 69);

Species x Concentration x Days (P = 0.41); SEM = 1.19.

219

Lumazine

For the three species studied, CH4 production in a medium that was 50 or 100%
saturated with lumazine was minimal (P < 0.01; Table 8-3). Inhibition was less at 25%
lumazine saturation. M ruminantium increase in CH4 production at 25% lumazine
saturation by d 6 was non-significant (P = 0.91; means, but not probability, are shown in
Table 8-3).

Table 8-3. Effects of lumazine on CH4 ) of three ruminal methanogensl

8312111911011

    
   

ruminantium

mazei

mobile

       

  

( < - ( < - ( = - );
Tube (Species x Concentration; P = 0.01); Species x Concentration (P < 0.01);
Species x Days (P = 0.07); Days x Concentration (P < 0.01);

Species x Days x Concentration (P < 0.01); SEM M ruminantium = 2.89;
SEM M mazei = 0.60; SEM M mobile = 0.08; 2Significance of linear contrast;
3Significance of quadratic contrast; ‘ND = not detected.

Lumazine concentration in the 50 and 100% lumazine-saturated media was lower
than 0.12 and 0.24 mM, respectively. However, the inhibition achieved was much
greater than what has been observed in mixed cultures with 1.2 mM lumazine (Chapters 4
and 6). It is possible that microbial species present in mixed cultures took up lumazine
and prevented it from inhibiting methanogens. Adsorption of lumazine to solid particles
could perhaps prevent its uptake by methanogens. Also, solid particles harbor a dense
adhered microbial population (Craig et al., 1987; Olubobokun and Craig, 1990), that may

accelerate lumazine catabolism, if it occurred.

220

Propynoic acid

Propynoic acid at 2 mM was strongly inhibitory for M ruminantium and M
mobile, and less inhibitory for M mazei (P < 0.01; Table 8-4). At 4 mM, propynoic acid
caused further inhibition of M ruminantium, but had minimal further effects on M mazei
or M mobile (P < 0.01). At both concentrations, inhibition of CH4 production of M
ruminantium and M mobile was greater than previously observed with ruminal mixed
cultures (Chapter 6). As M ruminantium and M mobile are generally the dominant
ruminal methanogens (Lin et al., 1997; Sharp et al., 1998), it would be expected that
propynoic acid inhibition of methanogenesis in mixed ruminal cultures would follow the
pattern observed for these two species in pure culture. As little propynoic acid
disappeared (Chapters 3 and 6), it is unlikely that the lower inhibition of methanogenesis
observed with mixed cultures was due to propynoic acid metabolism by otherruminal
microorganisms. It is also improbable that the lower inhibition of methanogenesis
observed in mixed cultures was due to propynoic acid adsorption to solid particles, as
propynoic acid is highly hydrophilic. M mazei was more resistant to propynoic acid than
the two other species examined. Other species not studied herein, like Methanosarcina
barkeri, could be more resistant to propynoic acid and perhaps explain the lower
inhibition of CH; production observed with mixed cultures. Importantly, no adaptation to

propynoic acid by any of the three species occured.

221

Table 8-4. Effects of propynoic acid on CH4 production (umoles) of three
ruminal ‘

    
  

res
1

ruminantium
Me 1 5
mazei
1
mobile 1 l 11 .
pecres < . < . 1 = . x
Concentration; P = 0.31); Species x Concentration (P < 0.01); Species x Days
(P = 0.93); Concentration x Days (P = 0.53); Species x Concentration x Days
(P = 0.49); SEM M ruminantium 0 mM = 0.43; SEM M ruminantium 2 mM = 0.34;
SEM M ruminantium 4 mM = 0.55; SEM M mazei 0 mM = 0.77; SEM M. mazei
2 mM = 0.86; SEM M mazei 4 mM = 0.65; SEM M mobile 0 mM = 2.15; SEM
M mobile 2 mM = 0.050; SEM M mobile 4 mM = 0.32 2Significance of linear
contrast; 3Significance of quadratic contrast.

 

Previous experiments with mixed ruminal cultures had found an increase in H2
accumulation when methanogenesis was inhibited by propynoic acid (Chapters 3 and 6).
This suggested that CH4 production was not being inhibited through competition for
electrons to reduce propynoic acid triple bond, as was originally hypothesized. The
present results support that conclusion, as CH4 production was inhibited at H2 partial
pressures 500 to 5,000 greater than those found in the rumen (Hungate, 1967; Kohn and
Boston, 2000). Furthermore, the complete reduction of propynoic acid triple bond at 4
mM concentration would have demanded 4 x 10'5 moles of H2, equivalent to
approximately 3.5% of the H2 originally present in the headspace. Thus, even if
propynoic acid’s triple bond had been completely reduced, methanogenesis would have
still been amply thermodynamically favorable. The inhibition exhorted by propynoic

acid, must be, therefore, non-competitive.

222

Ethyl 2-butynoate
Methane production of M ruminantium was almost completely inhibited at both

concentrations of ethyl 2-butynoate. M mazei was resistant to this chemical at the
concentrations studied. M. mobile was not affected by ethyl 2-butynoate at 4 mM, but
almost completely inhibited at 8 mM (M mobile quadratic contrast for days 4 and 6 P <
0.01; Table 8-5).

Table 8-5. Effects of ethyl 2-butynoate on CH4 production (umoles) of three

ruminal '

(mM)
.4

ruminantium
ina mazei

1 < .
l l 1 1 1 < .
pecres < . P < . < .
(P < 0.01); Tube (Species x Concentration; P < 0.01); Species x Days (P = 0.05);
Concentration x Days (P < 0.01); Species x Concentration x Days (P < 0.01); SEM = 1.02;
2Significance of linear contrast; 3Significance of quadratic contrast; ‘ND = not detected.

ium 4

 

Cell envelope differences could be related to the differences observed in toxicity
to ethyl 2-butynoate. The presence of an S-layer in M mazei and M mobile, which is
absent in Methanobrevibacter spp. (Sprott and Beveridge, 1993), may confer the former
some resistance to ethyl 2-butynoate. The methanochondroitin layer of M mazei (Sprott
and Beveridge, 1993) might confer this organism some additional protection to resist a
higher concentration of 8 mM ethyl 2-butynoate. Also, Methanosarcina aggregates in
large clumps (P01 and Demeyer, 1988; Moss, 1993) that may restrain the access of ethyl
2-butynoate to cells. Microscopic observations, however, did not find Methanosarcina

clump formation in the present experiment (not shown).

223

Resistance of M mazei to ethyl 2-butynoate can be a problem for its practical use
in vivo. If M mazei or other Methanosarcina are completely resistant to this chemical,
its long term use may preferentially select for them, and the inhibition of CH4 production
dissipate as the proportions of methanogens change.

The same rationale for the nature of inhibition of CH4 formation with propynoic
acid applies to ethyl 2-butynoate. At 8 mM initial concentration, the complete reduction
of ethyl 2-butynoate triple bond would have implied the utilization of approximately 7%
of H2 initially present in the headspace. Even if ethyl 2-butynoate triple bond had been
completely reduced, methanogenesis would still be largely thermodynamically favorable.
Other hypothetical reductions that this molecule could undergo (e. g., keto group in the
ester bond) would have little quantitative significance as a H sink in this experiment.
One must conclude that, as with propynoic acid, the inhibition exhorted by ethyl 2-
butynoate is non-competitive. Triesters such as trilinolein or trilinolenin at 10 mM have
been shown to be non-toxic to Methanobrevibacter (Methanobacterium) ruminantium
(Prins et al., 1972). Perhaps the toxicity of ethyl 2-butynoate to methanogens involves its
a smaller molecule size compared to those triglycerides, and/or to the presence of the

triple bond.

Conclusions

There were important differences among the three ruminal methanogens
examined with regard to their sensitivity to some of the compounds investigated. M
ruminantium was the most sensitive species to BES, propynoic acid, and ethyl 2-

butynoate, M mazei was the least sensitive species to those three chemicals, and M

224

mobile was intermediate. Conversely, M ruminantium was the least sensitive species to
lumazine. Differences among species in resistance to chemicals can have important '
practical implications as they may result in the selection of the more resistant species.
Adaptation of the ruminal rnicrobiota to BES was found after 3 d of feeding this chemical
to sheep (Van Nevel and Demeyer, 1996). This was likely related to differences among
methanogens regarding their sensitivity to the inhibitor. There is a need to monitor long
term changes in the populations of different methanogens when inhibitors of
methanogenesis are administrated. 16S rRNA probes could be useful for this purpose
(Lin et al., 1997; Sharp et al., 1998), especially to study unculturable organisms

(Y anagita et al., 2000; Tajima et al., 2001).

There seemed to be adaptation by the pure cultures to some of the chemicals. M
mazei seemed to adapt to 250 mM BES after 6 d. M ruminantium showed some
recovery at 50 mM BES between 4 and 6 d. Mechanisms of action of the inhibitors, and
of subsequent adaptation of individual species of methanogens, need to be studied before
in vivo strategies to inhibit ruminal methanogenesis can be developed.

Perhaps future in vivo strategies could be based on combinations or rotations of
different inhibitors of methanogenesis. Possibly, lumazine or propynoic acid could be
used to specifically target Methanosarcina and combined with other inhibitors to which

Methanosarcina is more resistant.

225

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Balch, W. E. and R. S. Wolfe. 1976. New approach to the cultivation of methanogenic
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Balch, W. E. and R. S. Wolfe. 1979a. Specificity and biological distribution of coenzyme
M (2-mercaptoethanesulfonic acid). J. Bacteriol. 137: 256-263.

Balch, W. E. and R. S. Wolfe. 1979b. Transport of coenzyme M (2-
mercaptoethanesulfonic acid) in Methanobacterium ruminantium. J. Bacteriol.
137: 264-273.

Boone, D. R., R. L. Johnson, and Y. Liu. 1989. Diffusion of the interspecies electron
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measurement of K”. for H2 or formate uptake. Appl. Environ. Microbiol. 55: 1735-
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Chang-Song, C., Y. Liu, M. Cummins, D. L. Valentine, and D. R. Boone. 2002.
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Jarvis, G. N., C. Str'o‘mpl, D. M. Burgess, L. C. Skillman, E. R. B. Moore, and K. N.

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Leadbetter, J. A. and J. A. Breznak. 1996. Physiological ecology of Methanobrevibacter
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hindgut of the termite Reticulitermesflavipes. Appl. Environ. Microbiol. 62:
3620-363 1.

Lin, C., L. Raskin, and D. A. Stahl. 1997. Microbial community structure in
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Muller, V., M. Blaut, and G. Gottschalk. 1993. Bioenergetics of methanogenesis. In: J. G.
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Rowe, J. B., M. L. Loughman, J. V. Nolan, and R. A. Leng. 1979. Secondary
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Sharp, R., C. J. Ziemer, M. D. Stern, and D. A. Stahl. 1998. Taxon-specific associations
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Smith, M. R. 1983. Reversal of 2-bromoethanesulfonate inhibition of methanogenesis in
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Smith, M. R. and R. A. Mah. 1981. 2-Bromoethanesulfonate: a selective agent for
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Stewart, C. S., H. J. Flint, and M. P. Bryant. 1997. The rumen bacteria. In: P. N. Hobson
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Tajima, K., T. Nagamine, H. Matsui, M. Nakamura, and R. I. Aminov. 2001.
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existence of a novel group of archaea not associated with known methanogens.
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Ungerfeld, E. M. 1998. Characterisation of resistance to 2-bromoethanesulphonate (BES)
in sheep rumen fluid. MSc thesis, Aberdeen University, Aberdeen.

Ungerfeld, E. M., S. R. Rust, M. K. Jain, and R. Burnett. 2002. Some miscellaneous
inhibitors of rumen methanogenesis in vitro. Beef Cattle, Sheep and Forage
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Van Nevel, C. J. and D. I. Demeyer. 1996. Control of rumen methanogenesis. Environ.
Monit. Asses. 42: 73-97.

Vicini, J. L., W. J. Brulla, C. L. Davis, and M. P. Bryant. 1987. Quin's oval and other
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Yanagita, K., Y. Kamagata, M. Kawaharasaki, T. Suzuki, Y. Nakamura, and H. Minato.
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detection of Methanomicrobium mobile by fluorescence in situ hybridization.
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I

Ye, D., J. F. Quensen, 111, J. M. Tiedje, and S. A. Boyd. 1999. 2-Bromoethanesu1fonate,
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65: 327-329.

229

CONCLUSIONS

None of the compounds or combinations of compounds studied substantially
decreased CH4 production without having some adverse effects on fermentation.
However, those effects were milder at low concentrations of the inhibitors, and there
were some unexpected beneficial effects such as decreased proteolysis and increased
microbial N production.

It was hypothesized that the inhibition of pyruvate oxidative decarboxylation
would result in less CO2 and H2 available for methanogenesis (Chapter 2). Inhibition of
pyruvate oxidoreductases was attempted directly and through the inhibition of thiamine
utilization, a cofactor of this reaction. Inhibition achieved in CH4 production was small
or non-existent. If further work in this line is to be considered, basic research on
thiamine uptake and metabolism in ruminal microorganisms and the enzymology of their
pyruvate oxidoreductases would be needed.

Alternative electron sinks to methanogenesis were examined (Chapter 3). Four
compounds, oxaloacetate, acetoacetate, B-hydroxybutyrate, and crotonate, are naturally
occuring intermediates in ruminal fermentation. Another four compounds, propynoic
acid, 3-butenoic acid, 2-butynoic acid, and ethyl 2-butynoate, are unsaturated analogs of
VFA or other compounds. Propynoic acid and ethyl 2-butynoate were strong inhibitors
of methanogenesis. However, fermentation also was reduced and fermentation products
without a nutritional value accumulated. It was apparent that methanogenesis was

inhibited directly, rather than through competition for reducing equivalents. B-

230

1

 

 

*4 ~_r-- unru-

Hydroxybutyrate, crotonate, and 3-butenoic acid, in contrast, had minimal effects on CH4
production, but seemed to stimulate fermentation.

Three novel compounds with unrelated mechanisms of inhibition of CH;
formation were studied in Chapter 4. The archaeal DNA-polymerase inhibitor aphidicolin
did not affect CH4 production. ~ A pterin, lumazine, decreased CH4 formation by about
50%, but also decreased apparent OM fermentation from 43.5 to 36.1%. Surprisingly, H2
accumulation was not observed and acetate molar percentage increased. Lumazine
inhibited the reduction of methyl-COM to CH4 in the last step of methanogenesis (Nagar-
Anthal and Nagle, 1997). The methyl-COM analog 2-bromopropanesu1fonate (BPS) did
not affect CH4 formation.

The effects of a novel hexadecatrienoic acid extracted from a marine algae and
olive oil on CH4 production and fermentation were investigated in Chapter 5. The
hexadecatrienoic acid decreased CH4 production by 97%, but decreased apparent OM
fermentation and increased H2 accumulation. Olive oil did not affect CH4 production or
fermentation, but increased propionate and butyrate production, and tended to decrease
acetate production. Perhaps olive oil could be used to increase dietary energy and
glucose supply without negatively affecting fermentation.

Some of the compounds studied in Chapters 2 through 5 decreased CH4
production considerably, but had other undesirable effects. Previous reports have shown
that other inhibitors of CH4 production result in an increased accumulation of
nutritionally non-usable products like H2, formate, and ethanol and decrease fermentation
(Van Nevel and Demeyer, 1996). These problems are a general consequence of an

inefficient relocation of the electrons spared from CH4 formation (Van Nevel and

231

Demeyer, 1996). The organic acids that seemed to stimulate fermentation in Chapter 3
could have acted as H sinks. Consequently, it was hypothesized that they could relieve
the negative effects on fermentation of the inhibitors of CH4 formation. Three inhibitors
of CH4 formation, lumazine, propynoic acid, and ethyl 2-butynoate, and two organic
acids that stimulated fermentation, crotonic acid, and 3-butenoic acid, were selected for
further investigation. The effects on fermentation of combinations between lumazine,
propynoic acid or ethyl 2-butynoate, and crotonic acid or 3-butenoic acid were examined
in Chapter 6. The effects of those combinations on OM, N, and NDF degradation and
microbial OM and N production were examined in Chapter 7.

Resistance of individual species of methanogens to chemical inhibitors could lead
to adaptation of the ruminal rnicrobiota, and difficulties to maintain long term inhibition
of CH4 production (Ungerfeld, 1998). The effects of lumazine, propynoic acid, ethyl 2-
butynoate, 2-bromoethanesulfonate (BES) and BPS, on pure cultures of three ruminal

methanogens were examined in Chapter 8.

Lumazine

Inhibition of CH; formation obtained with lumazine has been inconsistent,
ranging from 9 (Chapter 6) to 50% (Chapter 4). Ruminal fluid donors and substrate used
were similar in the experiments reported. However, the numbers of different species of
methanogens could have changed throughout time as experiments in Chapters 4 and 6
were more than one year apart. Previous observations with sheep suggested that the
numbers of methanogens resistant and sensitive to BES changed throughout time in

animals whose diet and management remained constant (Ungerfeld, 1998). Differences

232

in the proportions of methanogens in ruminal fluid may be responsible for the variable
results, as resistance to lumazine (measure through CH4 production by monocultures) was
shown to vary among species. Methanobrevibacter ruminantium was shown to be more
resistant to lumazine than Methanosarcina mazei or Methanomicrobium mobile (Chapter
8).

Other effects of lumazine as an inhibitor of ruminal CH4 formation have been
peculiar. No increase in unusual reduced end products of fermentation (Hz, formate, or
ethanol) were observed, which indicates that metabolic H was efficiently relocated into
other pathways. This cannot be totally explained by the relatively mild inhibition of CH;
formation observed with lumazine. A 50% decrease in CH; formation caused by
lumazine (Chapter 4) was not accompanied by any increase in H2, formate or ethanol
accumulation, while similar degrees of inhibition caused by intermediate concentrations
of propynoic acid or ethyl 2-butynoate resulted in increases in these products (Chapters 3
and 6). Although apparent OM fermentation (estimated through a mass balance)
decreased with lumazine addition (Chapter 4), it was then shown that true OM
degradation was unaffected (Chapter 7). A decrease in OM apparent fermentation
without changes in true degradation suggests a shift of OM sinks fiom fermentation
products to microbial biomass; however, the proportion of degraded OM incorporated
into microbial biomass was largely unaffected by lumazine (Chapter 7). Like the
accumulation of unusual reduced end products of fermentation such as H2, formate and
ethanol, a decrease in fiber degradation is another consequence of changes in electron
disposal and cofactor re-oxidation when methano genesis is inhibited (Van Nevel and

Demeyer, 1996). The lack of decrease of truly degraded OM (Chapter 7) again suggests

233

T

that the inhibition of CH4 formation by lumazine somehow did not create an electron
disposal problem.

It is also of interest that lumazine was a much stronger inhibitor of CH4 formation
in pure cultures of methanogens (Chapter 8) than in mixed ruminal cultures (Chapters 4
and 6). It seems likely that in mixed ruminal cultures lumazine was taken up and
metabolized. Decreases in N degradation and large changes in isobutyrate production
suggest that lumazine also affected non-methanogens. This is contrary to a previous
report, where there were no effects of lumazine on growth of non-ruminal bacteria
(N agar-Anthal et al., 1996). The effects of lumazine would need to be evaluated in pure
cultures of ruminal non-methanogens, and changes in the numbers of different microbial
groups in mixed ruminal cultures would need to be assessed.

It could be worthwhile to do further research on the effects of lumazine on
ruminal fermentation and animal production. From the applied nutrition point of view,
lumazine could cause some decrease in CH4 production in the rumen, and thus increase
the retention of fermented energy into usable products. It also may decrease ruminal
proteolysis and NH,“ concentration and increase microbial N production (Chapter 7).
Lumazine may have potential to increase the retention of dietary energy and N into
usable products. This may have beneficial environmental consequences, such as less CH4
and N released into the environment. The toxicological properties of lumazine have not
been investigated (MSDS, 2000), so its toxicity to ruminants and the environment would
need to be studied.

It is of great interest to understand why lumazine does not cause the problems in

the relocation of electrons that have been observed with other inhibitors of ruminal

234

methanogenesis. Lumazine increased (Chapter 4 and 6) or did not affect (Chapter 6) the
acetate to propionate ratio, which is contrary to what has been observed with other
inhibitors of ruminal methanogenesis (N agaraja et al., 1997). Stimulation of reductive
acetogenesis by lumazine could perhaps explain the shift towards acetate, the lack of
increase in H2 accumulation, and the decrease in C02 release sometimes observed
(Ungerfeld et al., 2002). It would be of interest to study the effects of lumazine on
ruminal acetogens.

A lumazine derivative, 6, 7-dimethyl-8-ribityllumazine, is a precursor of
riboflavin (Weimar and Neims, 1975). Riboflavin is in turn a precursor of the electron
acceptor FAD (V oet and Voet, 1995). The inhibition of CH4 formation with its
consequential disruption of the interspecies H transfer might elevate the requirements of
the mixed ruminal rnicrobiota for riboflavin, and perhaps also nicotinamide, an NAD

precursor. This would be another potential area for further research.

Propynoic acid

Propynoic acid was a strong inhibitor of ruminal methanogenesis in vitro.
Contrary to what was hypothesized originally, propynoic acid did not decrease CH4
formation by capturing reducing equivalents, but inhibited methanogens directly (Chapter
8). Propynoic acid had undesirable effects on degradation and increased the
accumulation of nutritionally non-usable fermentation products. Accumulation of these
products was not relieved by crotonic acid or 3-butenoic acid. 3-Butenoic acid could

mitigate the negative effects of propynoic acid on OM degradation (Chapters 6 and 7).

235

 

Since propynoic acid is moderately toxic to rodents and is not readily
metabolized, its practical utilization in ruminant diets is difficult. However, propynoic
acid caused substantial increases in microbial OM and N production, due to considerable
improvements in microbial synthetic efficiency (Chapter 7). It is of great interest to
understand how propynoic acid addition increased the efficiencies of microbial OM and
N synthesis. As little additive disappeared, these improvements were not caused by the
utilization of the additive as a C or energy source. If propynoic acid was deleterious to
protozoa, the improvement in microbial synthetic efficiency could be a consequence of
less intrarruminal N recycling due to less protozoal predation on bacteria (Williams and
Coleman, 1997). The decrease in VF A and NH4+ concentrations caused by propynoic
acid would be consistent with an inhibition of protozoa (Williams and Coleman, 1997).
However, lower VFA concentrations also could be a consequence of the lower OM
degradation, and lower NH4+ concentrations could be a consequence of the higher
production of microbial N observed with propynoic acid. Also, pr0pynoic acid at 2 mM
greatly improved microbial efficiencies of OM and N synthesis, but was not particularly
inhibitory of butyrate production (Chapter 6). As protozoa are strong butyrate producers
(Huhtanen, 1992; Jaakkola and Huhtanen, 1993), this may suggest that protozoa were
unaffected by propynoic acid. Protozoal numbers should be monitored to assess the
effects of propynoic acid.

Alternatively, propynoic acid may have inhibited deamination (Chapter 6). An
increase in the supply of preformed amino acids can improve microbial efficiency (Leng
and Nolan, 1984). Nevertheless, beneficial effects of amino acids occur with rapidly

degradable energy sources (Wallace et al., 1997), and there might be no benefit with

236

slowly degradable substrates as used in this study. Regardless, if protozoal activity or
lower deamination are not involved in the increased microbial synthetic efficiencies, it
would be very important to understand how propynoic acid caused these improvements,

and how this could be applied to manipulate ruminal fermentation.

Ethyl 2-butynoate

Similarly to propynoic acid, ethyl 2-butynoate was a strong inhibitor of ruminal
methanogenesis in vitro. It did not decrease CH4 formation by withdrawing reducing
equivalents, but inhibited methanogens directly (Chapter 8). Ethyl 2-butynoate caused
the accumulation of nutritionally non-usable fermentation products, which was not
relieved by crotonic acid or 3-butenoic acid. However, ethyl 2-butynoate at an initial
concentration of 4 mM did not decrease OM and NDF degradation. Furthermore, it
strongly increased microbial OM and N production through increased efficiencies of
synthesis. However, when the additive’s disappearance was considered, synthetic
efficiencies were unchanged.

The inclusion of ethyl 2-butynoate in ruminant diets to provide average ruminal
concentrations of approximately 4 mM (or slightly lower, as average concentrations in
vitro were lower than initial due to disappearance) could have an applied nutrition
interest. In vitro degradation of OM was not affected by 4 mM initial concentration of
ethyl 2-butynoate. Furthermore, in vivo small decreases in ruminal degradation could be
compensated post ruminally. Ethyl 2-butynoate at 4 mM could decrease CH4 production
by about 50%, decrease ruminal proteolysis and increase microbial N flow. This could

result in improvements in N retention and reduce N voided to the environment in feces

237

 

and urine. Ethyl 2-butynoate toxicity has not been investigated (GFS Chemicals, 1998),
but it is advantageous that it completely disappeared by 24 h of incubation. This would
decrease risks for the animal or the environment. The greatest problem to resolve,
however, would be to relieve the accumulation of H2, formate, and ethanol.

Effectiveness of the relocation of electrons spared from methanogenesis into nutritionally
useful electron sinks needs to be improved, either by using new electron acceptors, or by
developing new strategies to better utilize the electron acceptors already studied.

The negative effects of H2 accumulation on fermentation could be milder in vivo,
if the animal could release the increased gas pressure more frequently. Hydrogen
released to the atmosphere could perhaps escape the earth without causing the deleterious
effects that CH4 does.

Resistance of M mazei to 8 mM ethyl 2-butynoate and of M mobile to 4 mM
(Chapter 8) can be a problem for their potential use in vivo. If some methanogens are
resistant to this chemical, it is conceivable that long term use of ethyl 2-butynoate could
select the resistant organisms and lead to adaptation of the microbial community to the
chemical, as it has been found with BES (Van Nevel and Demeyer, 1996). Ethyl 2-
butynoate could be combined with lumazine, which exhorts a better control on M mazei
and M mobile.

Toxicity of ethyl 2-butynoate to ruminants and the environment, and potential

problems with its practical application also would need to be addressed.

238

 

I

Crotonic acid, 3-butenoic acid, and other electron acceptors

The inhibition of CH4 formation disrupts the interspecies H transfer and causes
the formation of end products without a nutritional value. It was hypothesized that the
addition of an external electron acceptor could solve this problem. However, both
crotonic acid and 3-butenoic acid were largely ineffective in decreasing the accumulation
of H2, formate, or ethanol (Chapter 6).

It was found that crotonic acid and 3-butenoic acid stimulated the substrate
apparent OM fermentation, as estimated through a mass balance (Chapter 3). Thus, it
was hypothesized that these organic acids could relieve the constraints on fermentation
caused by the inhibitors of CH4 formation. Crotonic acid was ineffective in relieving the
decreased OM and NDF degradation caused by propynoic acid or ethyl 2-butynoate
(Chapter 7). 3-Butenoic acid relieved some of the negative effects on OM and NDF
degradation caused by propynoic acid. Overall, the effects of crotonic acid and 3-
butenoic acid on true OM degradation were much smaller than initially observed on
apparent OM fermentation in Chapter 3. The combination of crotonic acid with
lumazine, and 3-butenoic acid with propynoic acid at 2 mM, decreased the proportion of
disappeared OM incorporated into microbial biomass. As true OM degradation was
unaffected, apparent OM degradation would increase. Although this mechanism can
explain why the increases in OM fermentation were not repeated for these two
combinations of inhibitors and organic acids, it does not apply to other combinations or
to crotonic acid and 3-butenoic acid overall. In addition, increased substrate apparent
OM fermentation was not consistently observed in Chapter 6 (data not shown): crotonic

acid increased (P < 0.01) the substrate apparent OM fermentability when combined with

239

f

lumazine, but not with propynoic acid or ethyl 2-butynoate. 3-Butenoic acid increased (P
= 0.04) the substrate apparent OM fermentability only when combined with ethyl 2-
butynoate.

It was a constant pattern throughout this work that externally added fermentation
intermediates were not completely converted to the fermentation pathways end products.
Most of added oxaloacetate did not seem to be converted to propionate, but to acetate.
Important amounts of the butyrate enhancers acetoacetate, B—hydroxybutyrate, and
crotonate also seemed to be converted to acetate (Chapter 3). Similar results have been
reported before for other organic acids (Lopez et al., 1999). These unexpected results can
be understood on thermodynamic grounds. Fermentation intermediates are normally
present at very low concentrations. Yet, their conversion into the next compound in the
pathway is still thermodynamically favorable, because the AGO of the reaction and the
concentration of the next intermediate in the pathway are sufficiently low. Reverse
reactions are thermodynamically unfeasible under normal conditions. However,
artificially elevated concentrations of a fermentation intermediate normally present at
very low concentrations could reverse the direction of the reaction.

One approach to resolve this problem would be to add the compounds
continuously, so that their concentration at any time would be only slightly elevated.
Thus, it may be more appropriate to do research on alternative electron acceptors in
continuous culture than in batch culture. Experiments done in the chemostat would more
closely reproduce events in the rumen when animals consume several meals per day.

However, the thermodynamic threshold for the reverse reaction of a pathway may

still be too low in terms of the added compound concentration. A second approach to

240

 

address this problem would be to combine intermediates of different pathways to achieve
the same molarity as with a single compound. This would allow lower concentrations of
each externally added compound. While keeping reverse reactions thermodynamically
unfavorable, it would still provide potential electron acceptors. Electron acceptors that
are not natural fermentation intermediates could also be added if conversion into other
compounds of the pathway does not occur. For example, if no isomerization occurs, a
50:50 molar ratio solution of 3-butenoic acid and crotonic acid would in theory be more
efficiently converted to butyrate than each compound added separately at a similar
molarity. Experiments with tracers would be needed to study how the concentration of
externally added fermentation intermediates affects the proportion that is converted to the
pathway end product.

Reductive acetogenesis could be an interesting alternative to rechannel electrons
into acetate formation, because it does not involve the addition of a metabolite at high
concentrations. Under normal conditions, H2 pressure in the rumen is too low for
reductive acetogenesis to compete with methanogenesis (Kohn and Boston, 2000).
However, with the increase in H2 partial pressure caused by the inhibition of CH4
production, reductive acetogenesis could become thermodynamically favorable, and H2

used to form acetate (Nollet et al., 1997).

241

 

 

 

 

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243

I

 

 

 

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297

Table A-42 Unprocessed data for Chapter 8

reatment
ruminantium
ruminantium contro
ruminantium
ruminantium
ruminantium
ruminantium
ruminantium
. ruminantium
ruminantium
ruminantium
ruminantium
. rum inantium
ruminantium
ruminantium
ruminantium
ruminantium
ruminantium
ruminantium
. ruminantium
ruminantium
ruminantium
ruminantium propynonc
ruminantium
ruminantium propynmc
ruminantium propynmc
. ruminantium propynmc
ruminantium propynmc
ruminantium y
ruminantium
ruminantium
. ruminantium
ruminantium
ruminantium
ruminantium
ruminantium
ruminantium
ruminantium
ruminantium
ruminantium
ruminantium
ruminantium
ruminantium
ruminantium
ruminantium
mazei
mazei
mazei
mazei

 

298

Table A-42 (cont’d)

. mazei
mazei
mazei
. mazei
mazei
mazei
mazei
mazei
mazei
mazei
mazei
M. mazei
mazei
mazei
mazei propynonc
mazei
mazei propynmc
. mazei propynmc
mazei propynmc
M. mazei propynmc
M. mazei
mazei
mazei
mazei
mczei
mazei
mazei
mazei
mazei
. mazei
mazei
mazei
mazei
mazei
mazei
mazei
mazei
mazei

 

299

Table A-42 (cont’d)

Treatment

ruminantium
ruminantium
ruminantium
rum inantium
ruminantium
ruminantium
ruminantium
rum inanti um
rum inantium
rum inantium
ruminantium
ruminantium

 

300

Table A-42 (cont’d)

reatment
ruminantium
ruminantium
ruminantium
ruminantium
ruminantium
ruminantium
ruminantium
rum inantium propynonc
ruminantium
ruminantium
rum inantium
rum inantium
ruminantium
ruminantium
. rum inantium
rum inamium
ruminantium
rum inantium
ruminantium
rum inantium
. ruminantium
ruminantium
ruminantium
ruminantium
ruminantium
ruminantium
mazei
mazei
mazei
mazei
mazei
mazei
mazei
mazei
mazei
mazei
mazei
mazei
mazei
mazei
mazei
mazei

 

301

Table A-42 (cont’d)

Days reatment

mazei
mazei
mazei
mazei
mazei
mazei
mazei
mazei
mazei
mazei
mazei
mazei
mazei
mazei
mazei
mazei
mazei

 

302

Table A-42 (cont’d)
Treatment
1

propynorc
PTOPYTIO

 

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REFERENCES

Acros. 1998-1999. Commercial catalog. Fisher Scientific.

Beilstein Database. 2002. MDL Information Systems GmbH. Beilstein Institut zur
Forderung der Chemischen Wissenschafien, Germany.

CRC Handbook of Chemistry and Physics. 1973. 53rd ed. CRC Press, Cleveland, OH.

Sigma. 2002-2003. Commercial catalog. Sigma-Aldrich Biotechnology LP.

The Merck Index. 1989. 1 1th ed. Merck and Co., Inc., Rahway, NJ.

308