improving the potency and reliability of...
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IMPROVING THE POTENCY AND RELIABILITY OF EXOGENOUS FIBROLYTIC
ENZYMES FOR ENHANCING FORAGE UTILIZATION BY DAIRY CATTLE
By
JUAN JOSE ROMERO GOMEZ
A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL
OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT
OF THE REQUIREMENTS FOR THE DEGREE OF
DOCTOR OF PHILOSOPHY
UNIVERSITY OF FLORIDA
2013
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© 2013Juan Jose Romero Gomez
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To my dear parents, Luz and Marino and my beloved wife Ana
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ACKNOWLEDGMENTS
I would like to thank my committee supervisor, Dr. Adesogan for his invaluable support
and dedication during my Ph.D. program. He has not only been my professor, but he has been a
mentor and guide for my professional development. I would also like to thank Dr. C.R. Staples,
Dr. C.F. Gonzalez and Dr. W. Vermerris for their priceless advice and guidance as members of
my committee. Special thanks to Dr. J.E.P. Santos for his guidance during my dairy trial even
though he was not in my committee.
I am indebted to Miguel Zarate, Zhengxin Ma, and Edis Macias for their hard work and
dedication during my experiments. Miguel is a dear and good friend who played a pivotal role
during my first and second experiments. Miguel also allowed me to participate in his thesis
experiment, which taught me a great deal about parasite control in goats. Zhengxin recently
became a very good friend and I appreciate her support during my third and fourth experiments. I
am grateful that she allowed me to assist with her experiments, which gave me ‘hands on’
experience working with mycotoxins. I am indebted to Edis for his dedication during my dairy
trial. His attention to detail and high quality work allowed me to conduct my dairy trial to a very
high standard.
I am grateful to Dr. Oscar Queiroz for his great advice and friendship. Oscar allowed me
to collaborate with him on two of his experiments. I obtained invaluable experience about
conducting and managing dairy cow experiments. This experience was very useful for my fourth
experiment. I also appreciate the support of Dr. Kathy Arriola during my first experiment. I
learnt a lot from her about silage analysis and conservation during her experiments.
I would like to thank Eric Diepersloot, Grady Byers, Brad Dicks, and the other Dairy
Unit staff for their help during my dairy cow experiment. Their many contributions allowed the
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experiment to proceed in a timely and efficient manner. Special thanks to Eric for his useful
advice and help during the planning and implementation stages of the dairy experiment.
I am very grateful to Jan Kivipelto for her support in the laboratory, even while going
through a tough time in her life. Thanks to Dr. Miriam Garcia for her friendship, advice and
assistance on various occasions during my experiments. I am thankful to Julio Schlaefli for
helping during the dairy trial, Shirley Levi for being the best secretary ever, and Mihai Giarnacu
and Carlos Martinez for their advice on the statistical analyses. Also special thanks are due to
Joseph Chakana Hamie, from whom I learnt about the role and importance of legume
supplementation in goat diets. Also, I am grateful to YeonJae Jang, Andres Pech, Fabio Kamada,
Uly Carneiro, Diego Garbuio, Rafael Marcondes, Bibiana Coy, Fabiola Martinez, Yun Jiang,
Kelly Mills, Illeana Brody and Chelsea Curry for their dedication and hard work while working
on my experiment. Each of them has a brilliant future ahead.
Thanks to Maggie and Daisy, our cannulated cows, for being such great animals and for
providing ruminal fluid for my experiments. Juanita who is at the Dairy Unit was also a great
cow at the Dairy Unit and I will miss her a lot.
There are no words to describe the great appreciation I have for my wife Ana. She has
been a great partner who understood the demands of doing experiments with animals. I thank her
a lot for her patience and support during the last three years. I promise her that we will spend
more quality time together after I graduate!
Finally, thanks to my parents, for all their support and guidance during my life. I could
not have accomplished my life goals without them. Thanks mom for raising me when dad passed
away and for all the sacrifices you did to help me achieve my dreams.
I am grateful to God that these great people were involved in my life.
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TABLE OF CONTENTS
page
ACKNOWLEDGMENTS ...............................................................................................................4
LIST OF TABLES .........................................................................................................................10
LIST OF FIGURES .......................................................................................................................13
LIST OF ABBREVIATIONS ........................................................................................................14
ABSTRACT ...................................................................................................................................16
CHAPTER
1 INTRODUCTION ..................................................................................................................18
2 LITERATURE REVIEW .......................................................................................................21
Overview .................................................................................................................................21
Factors affecting the Intake and Digestibility of Grasses .......................................................22 Chemical Composition of Forage Cell Walls ..................................................................22
Cellulose ...................................................................................................................23
Hemicellulose ...........................................................................................................24 Pectin ........................................................................................................................25
β-Glucans .................................................................................................................25
Structural proteins. ...................................................................................................26
Lignin. ......................................................................................................................26 Plant Cell Wall Development and its Impact on Digestion .............................................27
Primary wall phase of development. ........................................................................27 Secondary wall phase of development. ....................................................................28
Anatomical effects on grasses digestibility .....................................................................29
Tissues ......................................................................................................................30 Plant organs ..............................................................................................................31
The Fiber Requirement of Dairy Cattle ..................................................................................34 Classification of Fibrolytic Enzymes......................................................................................36
Classification and Functions of Fibrolytic Enzymes .......................................................37 Cellulases .................................................................................................................38
Hemicellulases (Xylanases) .....................................................................................40 Synergy Between Fibrolytic Enzymes ............................................................................42
Synergy between cellulases ......................................................................................42 Synergy between hemicellulases ..............................................................................44 Synergy between cellulases and hemicellulases ......................................................44
Carbohydrate-Binding Modules ......................................................................................44 Cellulase Kinetics ............................................................................................................45 Product and Substrate Inhibition .....................................................................................47
Cofactors and Coenzymes ...............................................................................................48
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Exogenous Fibrolytic Enzymes in Dairy Cattle Diets ............................................................49 Mode of Action of EFE in Ruminant Diets .....................................................................50
Preingestive effects ..................................................................................................50 Ruminal hydrolytic effects .......................................................................................51
Post-ruminal effects ..................................................................................................54 Microbial cellulose degradation ...............................................................................54
Non-Enzymatic Factors Affecting Efficiency of EFE .....................................................55 Manufacturing process .............................................................................................55 Influence of pH and temperature ..............................................................................57
Specificity of the EFE to the substrate .....................................................................59 Influence of the animal .............................................................................................60 Effects of the method of application of the EFE ......................................................61
3 SCREENING EXOGENOUS FIBROLYTIC ENZYME PREPARATIONS FOR
IMPROVED IN VITRO DIGESTIBILITY OF BERMUDAGRASS HAYLAGE ...............64
Background .............................................................................................................................64
Materials and Methods ...........................................................................................................65 Bermudagrass Substrate ..................................................................................................65
Enzymes ..........................................................................................................................65 EFE effects on In vitro ruminal digestibility (Experiment 1) ..........................................66 EFE Effects on Preingestive DM and Fiber hydrolysis (Experiment 2) .........................68
Proteomic Identification and Quantification of Proteins in Select EFE (Experiment
3) ..................................................................................................................................69
Statistical Analyses ..........................................................................................................69 RESULTS AND DISCUSSION .............................................................................................71
Experiment 1: ..................................................................................................................71 EFE effects on digestibility measures ......................................................................71
Accuracy of predicting digestibility measures from EFE activities .........................73 EFE effects on fermentation measures .....................................................................74
Experiment 2: ..................................................................................................................75
EFE effects on measures of preingestive hydrolysis ................................................75 Prediction of measures of preingestive hydrolysis from enzymatic activities .........78
Experiment 3: Proteomic Identification and Quantification of the Relative Ratio of
Less to More Effective EFE .........................................................................................80
Conclusions.............................................................................................................................82
4 EFFECT OF THE DOSE OF EXOGENOUS FIBROLYTIC ENZYME
PREPARATIONS ON PREINGESTIVE FIBER HYDROLYSIS AND IN VITRO
DIGESTIBILITY OF BERMUDAGRASS HAYLAGE .......................................................92
Background .............................................................................................................................92 Materials and Methods ...........................................................................................................93
Bermudagrass Substrate ..................................................................................................93
Enzymes ..........................................................................................................................94 In Vitro Ruminal Digestibility (Experiment 1) ...............................................................94 Preingestive Fiber Hydrolysis (Experiment 2) ................................................................96
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Statistical Analyses ..........................................................................................................97 Results and Discussion ...........................................................................................................98
Experiment 1: EFE Dose Effects on Measures of in Vitro Digestion and
Fermentation ................................................................................................................98
EFE dose effects on digestibility measures ..............................................................98 EFE dose effects on fermentation measures...........................................................100
Experiment 2: Effects of EFE Dose on Measures of Preingestive Hydrolysis .............104 Conclusions...........................................................................................................................107
5 EFFECT OF ADDING COFACTORS TO EXOGENOUS FIBROLYTIC ENZYMES
ON PREINGESTIVE HYDROLYSIS, IN VITRO DIGESTIBILITY AND
FERMENTATION OF BERMUDAGRASS HAYLAGE ...................................................116
Background ...........................................................................................................................116 Materials and Methods .........................................................................................................117
Bermudagrass Substrate ................................................................................................117 Enzymes ........................................................................................................................118
Screening COF for Synergistic Effects on the Hydrolytic Potential of EFE
(Experiment 1) ...........................................................................................................118
Effects of Adding Increasing Doses of COF to EFE on in Vitro Digestibility
(Experiment 2) ...........................................................................................................119 Statistical Analyses ........................................................................................................121
Results and Discussion .........................................................................................................122 Experiment 1: Effects of Cofactor Addition on Preingestive Hydrolysis .....................122
Experiment 2: Effects of Cofactor Addition to EFE on Digestibility and
Fermentation ..............................................................................................................128
Manganese addition to EFE 11C. ...........................................................................128 Iron addition to EFE 2A or 13D. ............................................................................131
Conclusions...........................................................................................................................134
6 IMPROVING FORAGE DIGESTION AND DAIRY COW PERFORMANCE WITH
FIBROLYTIC ENZYMES ...................................................................................................147
Background ...........................................................................................................................147 Materials and Methods .........................................................................................................148
Location, Housing and Weather ....................................................................................148 Animals and Treatments ................................................................................................149
Enzymatic Activities .....................................................................................................150 Sampling and Analysis ..................................................................................................150
Rumen Degradation Kinetics and Fermentation Measures ...........................................151 Statistical Analysis ........................................................................................................153
Results and Discussion .........................................................................................................155 Conclusions...........................................................................................................................159
7 GENERAL SUMMARY AND RECOMMENDATIONS ...................................................168
LIST OF REFERENCES .............................................................................................................175
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BIOGRAPHICAL SKETCH .......................................................................................................197
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LIST OF TABLES
Table page
2-1 Tissue proportions in organs of different forage types (Wilson, 1993). ............................32
3-1 Form, dose (g/kg of bermudagrass DM), biological source, activities of
endoglucanase, xylanase, exoglucanase, β-glucosidase of exogenous fibrolytic
enzyme (EFE) preparations used in in vitro digestion assays............................................83
3-2 Effects of exogenous fibrolytic enzyme addition on in vitro true dry matter (DMD),
neutral detergent fiber (NDFD), hemicellulose (HEMD) of a 4-wk regrowth of
Tifton 85 bermudagrass haylage (Experiment 1).a ............................................................84
3-3 Descriptive statistics for dependent and independent variables used to develop
regression relationships between activities of exogenous fibrolytic enzymes (EFE)
and measures of a 4-wk regrowth of Tifton 85 bermudagrass haylage. ............................85
3-4 The accuracy of predicting the in vitro digestibility of DM (DMD), NDF (NDFD) of
bermudagrass haylage from various enzyme activity estimates using stepwise
multiple regression (Experiment 1).a .................................................................................86
3-5 Effects of exogenous fibrolytic enzymes on concentrations of total volatile fatty
acids (TVFA), acetate, propionate, butyrate of a 4-wk regrowth of Tifton 85
bermudagrass haylage in buffered-rumen fluid (Experiment 1).a ......................................87
3-6 Effects of exogenous fibrolytic enzymes on DM loss and concentrations of NDF,
hemicellulose (HEM), ADF, cellulose (CEL), lignin (ADL) after preingestive
hydrolysis of a 4-wk regrowth of Tifton 85 bermudagrass haylage (Experiment 2).a .......88
3-7 The accuracy of predicting concentrations of neutral detergent fiber (NDF), water-
soluble carbohydrates (WSC), and ferulic acid (FER) of untreated and enzyme-
treated Tifton 85 bermudagrass haylage (Experiment 2).a ................................................89
3-8 Relative ratio of proteins in EFE 9C to those in 2A as detected by iTRAQ LC-
MS/MS analysisa. The EFE were sourced from both Trichoderma reesei and
Aspergillus sp. and from T. reesei, respectively. ...............................................................90
3-9 Relative ratio of proteins in EFE 11C to those in 2A as detected by iTRAQ LC-
MS/MS analysisa. Both EFE were sourced from Trichoderma reesei. .............................91
4-1 Endoglucanase, xylanase, exoglucanase, β-glucosidase (μmol of sugar released /
min× g) and ferulic acid esterase (nmol of ferulic acid released / min ×g) activities of
exogenous fibrolytic enzyme (EFE) preparations applied to bermudagrass haylage. .....109
4-2 Effects of the dose of exogenous fibrolytic enzymes (EFE) on in vitro true dry matter
(DMD), neutral detergent fiber (NDFD), hemicellulose (HEMD), acid detergent
fiber (ADFD) of a 4-wk regrowth bermudagrass haylage (Experiment 1).1 ...................110
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4-3 Effects of the dose of exogenous fibrolytic enzymes (EFE) on concentrations of total
volatile fatty acids (TVFA), acetate, propionate, butyrate, of the filtrate obtained
from fermentation of a 4-wk regrowth of bermudagrass haylage (Experiment 1)1 .........112
4-4 Effects of the dose of different exogenous fibrolytic enzymes (EFE) on DM loss and
concentrations of NDF, hemicellulose (HEM), ADF after preingestive hydrolysis of
a 4-wk regrowth of bermudagrass haylage (Experiment 2)1 ...........................................114
5-1 Endoglucanase, xylanase, exoglucanase, β-glucosidase (μmol of sugar
released/min/g) and ferulic acid esterase (nmol of ferulic acid released/min/g)
activities of exogenous fibrolytic enzyme (EFE) preparations used. ..............................135
5-2 Effects of adding cofactors (COF) to exogenous fibrolytic enzymes (EFE) on DM
loss (%), concentrations (% of DM) of NDF, hemicellulose (HEM) of a 4-wk
regrowth of bermudagrass haylage (Experiment 1).a .......................................................136
5-3 Effects of treating 4-wk regrowth bermudagrass haylage with exogenous fibrolytic
enzyme (EFE) 11C with or without increasing doses of Mn2+ on in vitro digestibility
of DM (DMD), NDF (Experiment 2).a ............................................................................138
5-4 Effects of treating 4-wk regrowth bermudagrass haylage with exogenous fibrolytic
enzyme (EFE) 11C with or without increasing doses of Mn2+ on concentrations of
total volatile fatty acids (Experiment 2).1 ........................................................................140
5-5 Effects of treating 4-wk regrowth bermudagrass haylage with exogenous fibrolytic
enzyme (EFE) 2A with or without increasing doses of Fe2+ on in vitro digestibility of
DM (DMD), NDF (Experiment 2).a .................................................................................141
5-6 Effects of treating 4-wk regrowth bermudagrass haylage with exogenous fibrolytic
enzyme (EFE) 13D with or without increasing doses of Fe2+ on in vitro digestibility
of DM (DMD), NDF (Experiment 2).a ............................................................................143
5-7 Effects of treating 4-wk regrowth bermudagrass haylage with exogenous fibrolytic
enzyme (EFE) 2A with or without increasing doses of Fe2+ on concentrations of total
volatile fatty acids (Experiment 2).a ................................................................................145
5-8 Effects of treating 4-wk regrowth bermudagrass haylage with exogenous fibrolytic
enzyme (EFE) 13D with or without increasing doses of Fe2+ on concentrations of
total volatile fatty acids (Experiment 2).a ........................................................................146
6-1 Ingredient and chemical composition (mean ± SD) of the Control diet used for the in
situ and lactation study.....................................................................................................161
6-2 Activities of endoglucanase, xylanase, exoglucanase, β-glucosidase (μmol of sugar
released min-1 g-1) of the exogenous fibrolytic enzyme (EFE) preparations mixed
with the dietary ingredients daily. ....................................................................................162
6-3 Effect of addition of fibrolytic enzymes to diet on intake by lactating dairy cows. ........163
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6-4 Effect of dietary treatment with fibrolytic enzymes on milk yield, feed efficiency,
yield and composition of milk fat, protein and lactose, somatic cell counts, body
weight and body condition score of lactating dairy cows. ...............................................164
6-5 Effect of dietary treatment with fibrolytic enzymes on in situ ruminal dry matter
degradation kinetics of a total mixed ration in lactating dairy cows1 ..............................165
6-6 Effect of dietary treatment with fibrolytic enzymes on ruminal fermentation
measures of lactating dairy cows1 ....................................................................................166
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LIST OF FIGURES
Figure page
6-1 Effect of dietary treatment with fibrolytic enzymes on milk yield of lactating dairy
cows. (Treatment × Week, P = 0.035; SEM = 0.59 kg/d) ...............................................167
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LIST OF ABBREVIATIONS
ADF Acid detergent fiber
ADFD Acid detergent fiber digestibility
ADG Average daily gain
ADL Acid detergent lignin
ADLD Acid detergent lignin disappearance
A:P Acetate to propionate ratio
BG β-glucosidase
BH Bermudagrass haylage
BW Body weight
BCS Body condition score
CBM Carbohydrate binding module
CEL Cellulose
CELD Cellulose digestibility
CMC Carboxymethyl cellulose
COF Cofactor
CON Untreated bermudagrass haylage
COU p-coumaric acid
CP Crude protein
DM Dry matter
DMD Dry matter digestibility
DMI Dry matter intake
E.C. Enzyme commission
EFE Exogenous fibrolytic enzymes
EN Endoglucanase
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EX Exoglucanase
FCM Fat corrected milk
FE Ferulic acid esterase
FER Ferulic acid
HEM Hemicellulose
HEMD Hemicellulose digestibility
iTRAQ Isobaric tags for relative and absolute quantification
IVDMD In vitro dry matter digestibility
NDF Neutral detergent fiber
NDFD Neutral detergent fiber digestibility
NDS Neutral detergent solubles
OM Organic matter
SE Standard error
SCC Somatic cell count
TMR Total mixed ration
TVFA Total VFA
VFA Volatile fatty acid
WSC Water-soluble carbohydrates
Wt Weight
XY Xylanase
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Abstract of Dissertation Presented to the Graduate School
of the University of Florida in Partial Fulfillment of the
Requirements for the Degree of Doctor of Philosophy
IMPROVING THE POTENCY AND RELIABILITY OF FIBROLYTIC ENZYMES FOR
ENHANCING FORAGE UTILIZATION BY DAIRY CATTLE
By
Juan Jose Romero Gomez
December 2013
Chair: Adegbola Adesogan
Major: Animal Sciences
The aim was to develop and evaluate the strategic use of in vitro tests to identify the best
exogenous fibrolytic enzymes (EFE) for improving the performance of lactating dairy cows fed
bermudagrass-based rations. Experiment 1 examined the effects of 18 EFE on fiber digestibility
and preingestive hydrolysis of bermudagrass haylage. Compared to the Control, 9, 3, 10 and 8
EFE increased neutral detergent fiber (NDF) digestibility (NDFD), NDF hydrolysis,
saccharification, and ferulic acid (FER) release, respectively. Experiment 2 examined effects of
increasing the dose of the 5 most promising EFE in Experiment 1 (1A, 2A, 11C, 13D and 15D)
on NDFD and preingestive hydrolysis. Increasing the dose of all EFE increased NDFD, NDF
hydrolysis, saccharification and release of FER in an EFE-specific manner. In Experiment 3, 5
cofactors (COF; Mn2+ , Co2+, Fe2+, Ca2+, and Mg2+) were screened to select the best candidates
for synergistically enhancing EFE-mediated increases in saccharification and NDFD of
bermudagrass. Saccharification was increased by adding all COF to EFE 2A and 11C or Mn2+,
Co2+, and Fe2+ to EFE 13D. Increasing the dose of Mn2+ in the presence of EFE 11C
synergistically increased NDFD, whereas increasing that of Fe2+ in the presence of 13D or 2A
decreased or did not affect most measures of digestibility and fermentation. Experiment 4
compared the effects of adding the most promising EFE from the previous experiments (2A or
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XYL) or a mixture of XYL and a Cellulase Plus enzyme (MIX) on the performance of 66
lactating dairy cows fed a ration containing bermudagrass and corn silage, alfalfa-orchardgrass
hay mix, and concentrates for 70 days. Feeding XYL increased dry matter intake relative to
feeding the Control or MIX diets, which had similar intakes. Milk yield was greater or tended to
be greater by cows fed 2A during weeks 3, 6, and 7 and cows fed MIX during weeks 6, 8 and 9
compared to those by Control cows. Therefore EFE treatment increased the performance of
lactating dairy cows. This study validated the use of in vitro NDFD tests to identify EFE that can
increase the performance of lactating dairy cows.
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CHAPTER 1
INTRODUCTION
The fundamental role of the dairy industry in the U.S. is reflected in its $140 billion
contribution to the economic output, $29 billion in household earnings, and its creation of well
over 900,000 jobs in the U.S. (DMI, 2004). A large portion of the total cost of producing milk
from dairy cattle is from purchased feeds (40 to 60 %; Bailey, 2009), with forages representing
38 to 45% of the feed cost (Chahine, 2004). Forages are the main feed source for ruminant
animals and they represent approximately 61% and 83% of the ration of dairy and beef cattle in
the U.S., respectively (Barnes and Nelson, 2003). In the southeastern U.S, warm-season grasses
are the basis of cattle production (Pitman, 2007), but their high fiber content and low digestibility
limit animal productivity and consequently profitability (Hanna and Sollenberger, 2007).
Therefore, improving the quality (nutritive value and intake) of warm-season grasses is a high
priority for the dairy industry in the southeast (Southeast Milk, Inc., 2011). Since bermudagrass
[Cynodon dactylon (L.) Pers.] is the most widely planted warm-season perennial grass for dairy
production in the southeast (10-12 million ha; Newman, 2007), it is an ideal model for testing
strategies to improve the quality of warm-season grasses.
Exogenous fibrolytic enzyme (EFE) application has been proposed as a method to
improve forage quality with studies showing improved fiber digestion and animal performance
due to application of such enzymes (McAllister et al., 2001; Beauchemin and Holtshausen, 2010;
Adesogan et al., 2013). However, use of EFE in ruminant diets is very limited because they have
produced ambiguous results due to the wide array of conditions used to test EFE and the limited
understanding of their mode of action (Beauchemin and Holtshausen, 2010; Adesogan et al.,
2013). The EFE effects are influenced by numerous factors such as dose (Eun et al., 2007) and
activity composition (Eun and Beauchemin, 2007), the prevailing pH and temperature (Arriola et
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al., 2011a), the animal performance level (Schingoethe et al., 1999), the experimental design
(Adesogan et al., 2013), and the fraction and proportion of the diet to which the enzyme is
applied (Krueger et al., 2008a; Dean et al., 2013). Arriola et al. (2011b) showed that adding an
EFE to a corn silage and alfalfa - based total mixed ration (TMR) fed to dairy cows increased
digestibility and increased feed efficiency. However, when a similar enzyme were applied to a
bermudagrass silage-based TMR, none of these performance measures was increased (Queiroz et
al., 2011). Consequently, research was needed to optimize the use of EFE to improve the quality
of bermudagrass. The series of experiments reported in this dissertation were conducted to screen
fibrolytic enzyme candidates to identify those that were ideal for hydrolyzing bermudagrass and
to examine conditions that could optimize the response in order to identify the best candidates
for increasing the performance of dairy cows fed a bermudagrass silage-based diet.
The literature review begins with an overview of the main factors affecting the intake and
digestibility of grasses including cell wall chemical composition, cell wall development and
plant anatomy. Then the fiber requirement of dairy cattle is discussed followed by a description
of fibrolytic enzyme characterization, nomenclature and mode of action. Factors affecting
enzymatic lignocellulose hydrolysis are also reviewed and strategies are proposed for using
fibrolytic enzymes to improve the digestibility of warm-season grasses and the performance of
cattle fed such forages. Chapters 3, 4, 5, and 6 describe experiments involved in a strategic
approach that involves screening and optimization of EFE to determine the best candidates and
doses for improving the utilization of bermudagrass by dairy cattle. Chapter 7 describes an
experiment aimed at validating the strategic approach by testing the effect of selected EF on the
performance of dairy cows fed a bermudagrass silage- based total mixed ration (TMR). In
particular, Chapter 3 describes an experiment designed to screen for the best EFE that will
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improve the in vitro NDF digestibility, fermentation and preingestive hydrolysis of bermudagrass
haylage. Also, regression relationships between enzymatic activities and digestibility measures
were explored in Chapter 3 and a proteomic assay was used to identify and quantify differences
in the composition of the most and least effective EFE at increasing the NDFD of bermudagrass
haylage. Chapter 4 evaluated the effects of the dose of the 5 most promising EFE from Chapter 3
on in vitro fiber digestibility, preingestive fiber hydrolysis and fermentation product
concentrations in order to determine the optimum EFE doses that maximize fiber digestion
efficiently. Chapter 5 describes the effects of adding cofactors (COF) to the EFE selected in
Chapter 3 on digestibility, preingestive hydrolysis and fermentation product concentrations.
Chapter 6 describes the effects of the most promising EFE and a previously effective EFE on the
performance of lactating dairy cows and the kinetics of ruminal degradation of the diet. The main
conclusions, deductions and implications of the studies are summarized in Chapter 6.
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CHAPTER 2
LITERATURE REVIEW
Overview
Exogenous enzyme supplementation is a well-established, widely used and effective
technology for improving feed efficiency and diet options for swine and poultry, which are
typically fed corn-soybean basal diets (Bedford and Partridge, 2010). However, use of
exogenous fibrolytic enzymes (EFE) in ruminant diets is very limited because studies that
examined effects of supplementing diets with EFE have produced equivocal results. This is
partly due to the wide array of conditions used to test EFE and the limited understanding of their
mode of action (Beauchemin and Holtshausen, 2010). Ruminants are very different from poultry
and because they have a 4-compartment stomach and their rumens house an extraordinary
variety of fibrolytic bacteria, fungi and protozoa (Russell, 2002). In most cases, enzyme
activities supplied by commercial EFE products are not novel to the rumen and therefore, EFE
act on the same plant cell wall targets as ruminal endogenous enzymes (Wang and McAllister,
2002). This might explain why EFE have been more effective at improving the productivity of
high-producing cattle rather than those fed at maintenance because the low ruminal pH and high
total tract rate of passage of high-producing cows in early lactation reduces ruminal fiber
digestibility (Mouriño et al., 2001; Cochran et al., 2007; Beauchemin and Holtshausen, 2010).
This problem can be exacerbated in cows fed the high-yielding warm-season grasses that abound
in tropical and subtropical regions because their high fiber contents and low digestibility limit
ruminal forage digestion thereby constraining animal productivity (Hanna and Sollenberger,
2007). The low digestibility of warm season grasses makes them ideal candidate substrates for
studies aimed at improving forage quality and animal performance with EFE that can
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compensate for the low digestibility by introducing exogenous fibrolytic capacity that
supplements the existing endogenous capacity in the rumen.
This review focuses on the factors affecting the intake and digestibility of grasses, the
fiber requirements of dairy cattle, characteristics, functions and nomenclature of fibrolytic
enzymes, the science underlying catalysis by such enzymes and their effects on cell wall
concentration, forage digestibility and the performance of dairy cattle.
Factors affecting the Intake and Digestibility of Grasses
Plant anatomy and chemical composition are the two main plant factors affecting the
voluntary intake and digestibility of grasses (Coleman et al., 2004). Several studies have been
conducted on the effects of the chemical composition of plants on their intake, digestion and
performance by cows. Such studies culminated in publication of the widely consulted and
referenced texts titled ‘Nutrient Requirements of Dairy (NRC, 2001) and Beef Cattle’ (NRC,
2000). Frequently updated databases containing the nutritive value of feeds and plants are also
accessible online (Dairy One, 2013). However, the effect of grass anatomy on voluntary intake
and digestibility has not been studied extensively. This is surprising because lignification of plant
tissues affects their digestibility (Akin, 1989), chewing and rumination time (Coleman et al.,
2004), particle size reduction (Wilson and Kennedy, 1996) and ruminal rate of passage (Kennedy
and Doyle, 1993). The ensuing section will describe the chemical and anatomical composition of
plant cell walls with a focus on their effects on intake and digestibility of plants by ruminants
followed by a review of the importance of fiber in ruminant nutrition.
Chemical Composition of Forage Cell Walls
The plant cell wall is a metabolically active, dynamic compartment with specific and
essential functions such as absorption, transport, and secretion of substances besides its role in
defense against bacterial and fungal pathogens (Evert, 2006). The plant cell wall is mostly
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comprised of structural polysaccharides (cellulose and hemicellulose) and lignin, with protein,
minerals and lipids as minor components (Theander and Westerlund, 1993).
Cellulose
Cellulose is the most abundant organic polymer on earth. It is the main constituent of
plants, where its main role is structural maintenance but it is present also in bacteria, fungi, algae,
and even animals (O’Sullivan, 1997). It is a linear polymer of β-D-glucopyranose linked by 1→4
glycosidic bonds with a wide molecular weight distribution (Aman, 1993). The β-linkages force
the glucose residues to rotate and the repeating unit is anhydro cellobiose (Aman, 1993).
Individual cellulose molecules are extremely large (~15,000 glucopyranose units in native
cellulose cotton) and are arranged into bundles known as microfibrils (Nelson and Cox, 2008).
The extended molecule forms a flat ribbon, which is further stiffened by intra- and
intermolecular hydrogen bonds, which produce a regular crystalline arrangement of the glucan
chain that affects the physical and chemical characteristics of cellulose (Aman, 1993). The
cellulose microfibrils wind together to form fine threads that coil around one another to form
macrofibrils, which have tensile strengths close to steel (Evert, 2006). Depending on hydrogen
bonding, the cellulose chains are highly ordered in some regions where strong hydrogen bonds
hold them together in structures called crystallites, whereas loosely-arranged cellulose molecules
form the amorphous regions (Bhat and Hazlewood, 2001). Pure cellulose is quickly and
completely digested in the rumen (Hatfield et al., 1999a).
There are six polymorphs of cellulose (I, II, III1, III11, IV1, and IV11) that can be
interconverted (O’Sullivan, 1997). The form found in nature is Cellulose I and the other forms
are obtained under experimental conditions. Cellulose II is obtained from cellulose I by
regeneration or mercerization; Cellulose III1 and III11 are formed from cellulose I and II,
24
respectively, by treatment with liquid ammonia; Cellulose IV1 and IV11 are prepared by heating
cellulose III1 and III11, respectively to 206oC in glycerol (O’Sullivan, 1997).
Hemicellulose
Hemicellulose is the second most common polysaccharide in nature. It is always
associated with cellulose in the plant cell wall and can be extracted with alkalis from delignified
walls (Bhat and Hazlewood, 2001). Hemicellulose can constitute more than 30% of the dry
weight of plants (Sun et al., 2004) and it is typically composed of heteropolysaccharides with
variable combinations of sugars and linkages (Hatfield et al., 2007). In grasses, the main portion
of hemicellulose is arabinoxylan, with a backbone of 1,4-linked xylose residues and substitutions
of arabinose, glucuronic acid, and acetic acid that can be attached at the two free OH groups of
C-2 and C-3 of the xylopyranose residue (Aman, 1993; Sun et al., 2004). Ferulic acid and p-
coumaric acid can be found ester linked to the C-5 hydroxyl group of some arabinose
substitutions (Hartley, 1972). Hydroxycinnamates (ferulic acid, p-coumaric acid and sinapic
acid) are structurally related to lignin precursors and they may attach to lignin, playing critical
roles in regulating wall matrix organization (Hatfield et al. 1999a). Hydroxycinnamic acids can
cross-link polysaccharides with other polysaccharides or lignin (Ralph, 1996), and both of these
linkages result in decreased digestibility (Hatfield, 1993). The type and frequency of branch
substitutions varies with species and stage of development (Hatfield et al., 2007). For instance,
corn fiber contains 48-54% xylose, 33-35% arabinose, 5-11% galactose, and 3-6% glucuronic
acid (Saha, 2003). Also, acetyl groups can represent 1-3 % of the total DM in grasses (Theander
et al., 1981). In wheat straw, for every 26, 13, or 18 D-xylopyranosyl residues in the main chain,
there is one uronic acid unit, one L-arabinofuranosyl group and one xylopyranosyl residue,
respectively (Sun et al., 1996). The arabinoxylans can form hydrogen bonds with cellulose and
25
each other (Evert, 2006), but these linkages do not inhibit the rate of digestion of either
polysaccharide in ruminal fluid (Weimer et al., 2000).
Pectin
Pectin is a major component of primary cell walls composed of a range of galacturonic
acid-rich polysaccharides grouped in three major types, homogalacturonan, rhamnogalacturonan-
I and rhamnogalacturonan-II (Willats et al., 2001). These polysaccharides function as hydrating
agents and cementing material for the cellulosic network and make up about one third of the cell
wall of dicotyledonous and monocotyledonous plants (Thakur et al., 1997). One of the main
exceptions occurs in the cell walls of the species in the Graminae family because pectins only
account for 10 µg/g or less of the DM in grasses (Hatfield et al., 1999a).
In most plants, pectin is deposited in the middle lamella between the primary cell wall of
neighboring cells, especially in soft plant tissues under rapid growth and high moisture contents
(Thakur et al., 1997). Pectin forms a gel in the presence of Ca2+ or at low pH (Willats et al.,
2001) and it is rapidly and extensively degraded from cell wall matrices during ruminal
fermentation (Hatfield et al., 1999a). Due to their minor role in grass cell walls, pectins will not
be discussed further.
β-Glucans
Mixed linkage 1,3- and 1,4-β-D-glucans are linear homopolymers of D-glucopyranosyl
residues linked mostly via 2 or 3 consecutive β(1,4) linkages separated by a single β(1,3) linkage
(Izydorczyk and Dexter, 2008). Their occurrence is is limited to immature tissues and endosperm
cell walls of cereals (Hatfield, 1989) and are rapidly and almost completely digested in the
rumen (Grove et al. 2006).
26
Structural proteins.
Primary cell walls typically contain O-glycosylated proteins (Evert, 2006), including
hydroxyproline-rich glycoproteins (extensins) which are glycosylated with arabinose,
arabinobiose, arabinotriose, arabinotetraose, and galactose (O’Neill and York, 2003), the proline-
rich proteins, which are lightly glycosylated (Cassab, 1998), and the glycine-rich proteins, which
are glycoslylated with mannose, arabinose, glucose, xylose and galactose (Matsui et al., 1995).
Structural proteins like extensins, appear to play critical roles in cross-linking wall components,
particularly in primary walls (Hatfield et al., 1999a). Proteins generally make up less than 5 µg/g
of the grass cell wall depending on tissue type and maturity (Hatfield et al., 1999a). Proteins
outside the cell wall matrix are susceptible to rumen digestion, but those within the cell wall may
be completely resistant to rumen microbes and pass intact through the digestive tract.
Lignin.
Lignin is the generic term for a large group of aromatic polymers resulting from the
oxidative combinatorial coupling of 4-hydroxyphenylpropanoids (Vanholme et al., 2010). The
primary precursors of lignin are coniferyl, sinapyl, and p-coumaryl alcohol, which undergo
enzyme-initiated dehydrogenative polymerization during lignin formation (Sarkanen and
Ludwig, 1971). Generally, lignins are classified as guaiacyl (formed predominantly from
coniferyl alcohol), guaiacyl-syringyl (copolymers of coniferyl and sinapyl alcohols) or guaiacyl-
syringyl-p-hydroxyphenyl lignins (formed from all three monomers), according to whether they
are from gymnosperms, woody angiosperms, or grasses, respectively (Evert, 2006). Guaiacyl and
syringyl residues are the most abundant in grasses (Hatfield et al., 2007).
From a nutritional standpoint, lignin is a phenolic-derived macromolecule that interacts
with other wall polymers to provide structural integrity, resistance to degradation, and water
impermeability (Hatfield et al., 1999a). It is the most significant factor limiting the availability of
27
plant cell wall material to ruminants (Van Soest, 1994), but lignin composition does not seem to
directly affect ruminal cell wall digestibility as previously thought (Grabber et al., 1998a; Jung et
al., 1999). The decrease in digestibility observed with increased syringyl- to guaiacyl-type lignin
(Jung and Deetz, 1993) actually was biased by the fact that slowly digestible secondary cell walls
are intrinsically higher in syringil lignin concentration (Jung and Engels, 2002; Grabber, 2005).
The importance of the relative effects of lignin, such as acting as a physical barrier
impeding enzyme digestibility, or as a sequesterer of enzymes by nonspecifically adsorbing
them, is still unclear (Bansal et al., 2009). Also, lignin is a hydrophobic filler that replaces water
in the cell wall (Evert, 2006). The anaerobic rumen environment prevents digestion of lignin by
the rumen microbes (Hatfield et al., 2007). Much of the deleterious effects of lignin on
digestibility result from its interaction with cell wall polymers (Hatfield et al., 1999a) and they
are better discussed under the context of plant cell wall development.
Plant Cell Wall Development and its Impact on Digestion
According to Terashima et al. (1993), the growth and development of the cell wall in
plants can be divided into two phases:
Primary wall phase of development.
The primary phase comprises the increase of the plant cell size mainly due to cell wall
elongation (Jung and Allen, 1995). The primary wall, composed of the first-formed wall layers,
is deposited before and during the growth of the cell (Evert, 2006). After plant cells have reached
mature size, additional development of the primary wall or deposition of a secondary wall
structure can occur (Hatfield et al., 2007). At the primary cell wall development stage cell walls
are composed of cellulose (20-30 %), arabinoxylans (20-40 %), xyloglucans (1-5 %), mixed
linkage glucans (10-30 %), pectin (5 %) and proteins (1 %) (Vogel, 2008).
28
Grasses are also characterized by the presence of ferulic and p-coumaric acids (1-5 %;
Vogel, 2008), which form esters with sugars that reduce polysaccharide digestion (Jung and
Casler, 1991). Lignin deposition is very limited during this initial phase (Evert, 2006). Living
cells with only primary walls may remove cell wall previously acquired in order to differentiate
into other cell types. Consequently only primary cell walls are typically involved in wound
healing and regeneration in plants (Evert, 2006).
Secondary wall phase of development.
The secondary phase of cell wall development mostly initiates after the mature cell has reached
its final size. Secondary cell wall composition in grasses is reported to be 35-45% cellulose, 40-
50% arabinoxylans, 0.1 % pectins, 0.5-1.5 % ferulic and p-coumaric acids, 20 % lignin, 5-15%
silica with minor presence of mixed-linkage glucans, xyloglucans, mannans, and glucomannans
(Vogel, 2008). Secondary cell walls are deposited inside of the primary cell walls and comprise
at least 50% of the cell wall mass in both leaves and stems (Vogel, 2008). The arabinoxylan
found in secondary cell walls has fewer substitutions than in the primary wall allowing for a
stronger interaction with cellulose (Vogel, 2008). Lignin deposition starts at this phase in the
middle lamella - primary wall region and then continuous into the lumen side of the cell wall
(Terashima et al., 1993). Because lignin deposition lags behind polysaccharide aggregation, the
most recently deposited polysaccharides are not yet embedded in lignin; therefore, they are more
digestible (Jung and Allen, 1995). In contrast, the middle lamella - primary wall region is the
most intensely lignified and least digestible region (Jung and Allen, 1995). This partly explains
why microbes digest recalcitrant cells from the inside out (Grant, 2009).
Ferulic acid substitutions act as nucleation sites for the lignification process since radical
coupling to lignin can only occur if ferulates react with monolignols (Ralph et al., 1995).
Hatfield et al. (1999b) suggested that the positioning of ferulates within the wall might regulate
29
lignin formation patterns and control cross-linking within wall matrices. Ferulic acid (Jung,
2003) and p-coumaric acid form ester and ether bonds with lignin, but only form ester bonds
with polysaccharide chains (Ralph and Helm, 1993). Typically, p-coumaric acid is esterified or
etherified to lignin but it does not form cross-linked structures with both ester and ether linkages
(Lam et al., 1991, 1992). In contrast, ferulic acid can be both esterified and etherified and it is
involved in cross-linkages between lignin and arabinose (Lam et al., 1991, 1992). The formation
of diferulates from ferulic acid monomers ester-linked to arabinose in a polysaccharide chain
covalently couples two polysaccharides chains (Hatfield et al., 1999b), plays a vital structural
role in plants and decreases the rate and possibly the extent of polysaccharide degradation
(Grabber et al., 1998b). Furthermore, the crosslinking of arabinoxylans to lignin via ester and
ether linkages with ferulates reduces digestion significantly (Jung and Deetz, 1993; Grabber et
al., 1998 a, b). Grabber (2005) mentioned that each percentage unit increase in lignin
concentration depressed cell wall degradability by two percentage units in his model based on
maize (Zea mays L.) cell walls. Casler and Jung (1999) reported that higher levels of ferulate
cross-linking at the same lignin concentration reduced neutral detergent fibre (NDF) digestibility
of smooth bromegrass (Bromus inermis L.).
Anatomical effects on grasses digestibility
The effect of grass anatomy on voluntary intake and digestibility has not been studied
extensively. This is surprising because the different lignification patterns of plant tissues affect
their digestibility (Akin, 1989), and consequently affect chewing and rumination time (Coleman
et al., 2004), particle size reduction (Wilson and Kennedy, 1996) and ruminal rate of passage
(Kennedy and Doyle, 1993) consequently affecting animal production (Batistoti et al., 2012).
This section describes the anatomy of the nutritionally relevant plant tissues and organs and
explains how they affect forage intake and digestibility, using bermudagrass as the model.
30
Tissues
Epidermis. This is the outermost (dermal) cell layer in plants and it usually consists of
one layer of cells in thickness (Evert, 2006). The epidermis accounts for approximately 26% of
the cross sectional area (CSA) of bermudagrass leaf blades (Wilson, 1993; Rudall, 2007). The
epidermis is thicker and has more lignin in mature grasses (Akin, 1989; Wilson, 1993). The
cuticle covers the epidermis with complex waxes, cutin and phenolic compounds (Gevens and
Nicholson, 2000), which prevent degradation of the cuticle in the rumen (Akin, 1989).
Mesophyll. These are thin-walled cells that comprise photosynthetic chlorenchymatous
tissue internal to the epidermis in leaves but they hardly occur in stems (Rudall, 2007).
Mesophyll cells account for approximately 27% of the leaf blade CSA tissue in bermudagrass
(Akin, 1989). Relative to other cells, they have a loose arrangement that creates a larger exposed
surface area for bacteria colonization and also increases the ruminal rate of passage (Wilson,
1993). Mesophyll cells do not contain lignin, hence they are rapidly digested by cows (Akin,
1989).
Parenchyma bundle sheath. This is a highly specialized group of chlorenchymatous
cells surrounding the vascular tissue in leaves, which accounts for 28% of the leaf blade CSA
tissue in bermudagrass (Akin, 1989). It is called the “Kranz” sheath in C4 grasses (Moore et al.,
2004). With maturity, the thickness of the PBS increases (Wilson, 1990) and their lignin contents
vary with many factors such as stressful conditions (Akin, 1989) and maturity. As PBS CSA
increases, the NDF concentration will increase too (Batistoti, 2012).
Nonchlorenchymatous Parenchyma. These cells account for approximately14% of the
leaf blade tissue CSA in bermudagrass (Wilson, 1993). They are thin-walled in leaves and
usually rapidly and almost entirely digested (Wilson, 1991). However, in the leaf sheath (66 %
CSA) and stems (75 % CSA), the nonchlorenchymatous parenchyma cells can reduce
31
digestibility because they can develop thick secondary walls that can undergo lignification as
they mature (Wilson, 1993).
Sclerenchyma. These cells provide support and protection because they form highly
lignified fibers and sclereids tissues (Rudall, 2007). This tissue only accounts for 10% of the leaf
blade CSA in bermudagrass (Akin 1989). In the leaves, they are present as patches above and
below vascular tissue and in the stems they can form an outer ring that shields
nonchlorenchymatous parenchyma. Sclerenchyma cells can reduce DMI because when
consumed by animals, they increase the time needed for particle size reduction due to their
structural role in the plant (Rudall, 2007).
Vascular tissue. The vascular tissue comprises the xylem and phloem vessels. The
phloem represents only a small fraction of leaf, sheath, and stem CSA and it is rapidly digested
because it has thin-walled cells and lacks lignin (Akin, 1989). In contrast, the cells of the xylem
are thick and heavily lignified in all plant organs (Wilson, 1993).
Middle lamella.This is the intercellular layer between walls of neighboring cells (Rudall,
2007) where lignin formation is initiated and is more concentrated (Jung and Allen, 1995).
Therefore, this layer is one of the greatest constraints to fiber degradation by ruminant microbes
(Coleman et al., 2004).
Plant organs
The two main plant organs of importance in forage grasses fed to ruminants are leaves
and stem. The digestibility of leaf blades and stems is associated with the relative proportion of
tissue types in each organ with digestible or indigestible cell walls (Wilson, 1993; Wilson and
Hatfield, 1997). Since leaves have a high proportion of mesophyll cells, they are much more
digestible than the lignified stem, which is high in nonchlorenchymatous parenchyma cells. In
fact, a high leaf to stem ratio is usually used as an indicator of greater nutritional value of forages
32
(Balasko and Nelson, 2003). The relative proportions of the different tissue types among organs
in C4 and C3 grasses are shown in Table 2-1.
Table 2-1. Tissue proportions in organs of different forage types (Wilson, 1993).
Proportion of tissue in cross sectional area (%)
Cell Type C4 grass
(Panicum maximum)
C3 grass
(Lolium multiflorum)
Blade Sheath Stem Blade Sheath Stem
Epidermis 22 4 2 23 NM 2
Mesophyll 31 7 2 66 86 2
PBS1 24 7 0 5 0 0
Sclerenchyma 2 6 8 1 10 12
NCPAR2 14 66 75 2 NM 75
Vascular tissue
(without
phloem)
6 9 12 3 4 9
Phloem <1 1 1 <1 <1 <1 1PBS= parenchyma bundle sheath, 2NCPAR= non-chlorenchymatous parenchyma.
NM= not mentioned
Leaves. Leaves of grasses are borne on the stem, one at each node, but are projected
alternatively in two rows on opposite sides of the stem (Moser and Nelson, 2003). Each leaf
consists of a sheath, blade, and collar area.
The blade is parallel veined and typically flat and narrow (Moser and Jennings, 2007). It
is the most digestible part because it contains the highest proportion of mesophyll tissue. Blades
of C4 grasses exhibit Kranz anatomy, which has many vascular bundles surrounded by a
specialized, large, parenchyma bundle sheath (Rudall, 2007). In contrast, C3 grasses have
palisade and spongy mesophyll throughout with widely spaced veins (Moser and Jennings,
2007). Consequently, C4 grass leaf blades have a higher proportion of the less digestible, thick-
walled, lignified tissues (parenchyma bundle sheath, sclerenchyma, and vascular tissue) than C3
grasses.
The sheath surrounds the stem above the node where it is attached. It is green and
photosynthetic, but it functions mainly to physically support the blade and to transport materials
33
between the blade and the stem (Moser and Jennings, 2007). The anatomy of leaf sheaths is
intermediate between those of the blade and stem, but they are more like that of the stem. Thus,
sheaths also have lignin concentrations between those of blades and stems and consequently their
digestibility falls between those of blades and stems (Wilman and Altimimi, 1982). Sheath tissue
proportions do not appear to change with maturity (Cherney and Marten, 1982), but there are
marked increases in wall thickness of lignified cells (Wilson, 1976).
Stem. Forage grasses have two distinct forms of vertical stems, both with the shoot apex
at the tip (Moser and Jennings, 2007). In vegetative tillers, stems are very short, consisting of
nodes and basal, non-elongated internodes. This allows the shoot apex to remain near the ground
and escape grazing or cutting (Moser and Jennings, 2007). In reproductive tillers, the dormant
internodes begin to elongate and elevate the future inflorescence and if cut at this point, the tiller
dies (Moser and Jennings, 2007). Stems differ from leaf blades in that their tissue characteristics
change greatly with maturity (Cherney and Marten, 1982), such that stem digestibility can be
similar to or greater than that of leaves when young, but lower than that of leaves when mature
(Hacker and Minson, 1981). When the stem is young, the vascular tissue is in isolated bundles
but as it matures, the bundles link together through lignification of the interfascicular
nonchlorenchymatous parenchyma cells, which form a strong, indigestible tissue (Wilson, 1993).
Eventually, this tissue forms a ring that embraces the entire cortical region of sclerenchyma and
epidermis in most grasses and constitutes a powerful barrier to digestion (Wilson, 1993).
There are three layers of factors that influence the nutritional value of grasses. First, the
organ proportion; second, the tissue proportion within an organ; third, the lignification and
structural carbohydrates present within the different tissues in an organ. The interplay between
these factors creates a high degree of complexity in the determination of the nutritive value of
34
grasses. In the cases were the grass is chopped and mixed, only the chemical composition effects
remain; however, in grazing dairies, grasses are grazed or fed with minor or no physical
processing, leaving the anatomical effects intact. More research is needed on the effects of plant
anatomy on ruminant nutrition. Ruminant nutritionists should work with plant breeders to
develop strategies (e.g. selection, genetic modification) to modify organ and tissue proportions in
relevant species in ways that increase digestibility, without affecting yield and pest resistance.
The Fiber Requirement of Dairy Cattle
Fiber is defined as the complex of dietary nutrients which are relatively resistant to
digestion and are slowly or partially digested by ruminants (Moore and Hatfield, 1994).
Cellulose, hemicellulose and lignin are the major components of fiber and in ruminant nutrition,
these polymers are collectively and chemically classified as neutral detergent fiber (NDF),
whereas acid detergent fiber (ADF) is NDF minus the hemicellulose (Van Soest et al., 1991). For
non-ruminant species, pectins and β-glucans are also included as fiber fractions since they are
not digested by mammalian enzymes (Moore and Hatfield, 1994).
On average, NDF is less digestible than other nutrients (e.g. starch), therefore when NDF
concentration is increased in the diet, the energy density decreases, and voluntary intake is
reduced by rumen fill (NRC, 2001; Mertens, 2007) and the productivity of lactating cows is
limited. However, NDF cannot and should not be eliminated from ruminant diets since ruminants
require adequate dietary fiber for normal and healthy rumen function (Van Soest, 1994). A
myriad of negative outcomes typically occur when diets contain insufficient NDF concentrations
including acidosis, liver abscesses, milk fat depression, laminitis, displaced abomasum among
other conditions. (NRC, 2001). However, NDF concentration is not a good indicator of the type
of fiber that effectively stimulates the ruminal mat formation, rumination, salivation, and ruminal
motility needed for a healthy rumen (VandeHaar, 2005). Instead, physically effective NDF
35
(peNDF), is a term that accounts for the chemical fiber content and particle size of feedstuffs,
resulting in a much better measurement of the adequacy of dietary fiber in dairy cattle diets
(Zebeli et al., 2012). However, particle size has two antagonistic effects on animal performance
(Zebeli et al., 2012). When long forage particles are fed, rumination is stimulated, more saliva is
produced and consequently more buffer is available to prevent deleterious reductions in ruminal
pH, which can decrease ruminal fiber digestion. However, excessively large particle sizes
decrease the gastrointestinal rates of passage and digestion due to a lower particle surface area.
Several methods have been proposed to measure physically effective fiber (Mertens,
1997; Buckmaster et al., 1997). In its latest incarnation, it is measured from a three-screen sieve
and a bottom pan (> 19 mm, 8 to 19 mm, 1.18 to 8mm and < 1.18mm) and the NDF
concentration of each fraction (Kononoff and Heinrichs, 2003). However, the lack of validation
of the accuracy of using this method to measure peNDF has hindered the widespread use of this
concept (NRC, 2001). Thus, the dairy NRC (2001) recommended that under most feeding
situations in U.S., lactating cow diets should contain at least 25% total NDF and 19 % of the DM
must be NDF from forage in order to maintain normal rumen function and prevent milk fat
depression (NRC, 2001). No final suggestion was made for peNDF by the NRC committee.
However, by using models that account for the effects of both peNDF and fermentable starch in
the rumen, Zebeli et al. (2012) recently suggested that a diet containing 31.2% peNDF inclusive
particles exceeding 1.18 mm or 18.5 % peNDF inclusive particles exceeding 8 mm in the diet is
required to maintain a healthy rumen. They noted however, that having more than 14.9% peNDF
inclusive particles exceeding 8 mm will decrease intake and can limit productivity. Thus,
research is needed to develop rations that are energy-dense but don’t require increases in peNDF
above the threshold that leads to lower intake (Zebeli et al., 2012).
36
Forages alone cannot meet the energy requirements of the lactating cow since their lower
digestibility compared to concentrates limit the energy density of the diet. It could be argued that
lactating cows should be fed the least possible amount of forages to maximize their milk
production. However, there are several advantages to feeding forages at high inclusion rates in
the diet (Jung and Allen, 1995). First, forages are necessary to maintain rumen function and
animal health, as previously discussed. Second, there is an economic benefit to using forages
since their production costs are usually lower than those for grain crops. Third, perennial forages
prevent soil erosion and require less pesticide and fertilizer inputs, therefore, they are more
sustainable and environmentally friendly than feeding concentrates in the long run. Thus, it is
necessary to overcome the nutritional limitations of forages and to enhance nutrient availability
from forages for milk production. Although various strategies for increasing forage quality exist
such as plant breeding and post-harvest treatments, most are too expensive, hazardous or
protracted for routine use. Fibrolytic enzymes do not have these limitations and because they
catalyze fiber depolymerization, they are promising additives for achieving the desired increases
in forage quality.
Classification of Fibrolytic Enzymes
Traditionally, enzymes were classified according to the substrate they hydrolyzed by
including or adding the suffix ‘ase’ to the name of the substrate. Subsequently, the International
Union of Biochemistry (IUB) Enzyme Nomenclature System classified enzymes into
oxidoreductases, transferases, lyases, isomerases, ligases and hydrolases and most fibrolytic
enzymes are in the latter group. This system was based on both the type of reaction that enzymes
catalyze and their substrate-specificity (Paloheimo et al., 2010). This system avoids the
confusion caused by of different enzyme names, but it does not provide information about
structural features of enzymes (Henrissat, 1991). Consequently a sequence-based classification
37
was developed, which involved using algorithmic methods to assign sequences to various
families (Withers and Williams, 2013). In this system, fibrolytic enzymes are referred to as
glycoside hydrolases (GH) and they have been classified into more than 100 families, each
containing proteins related by sequence, and consequently by fold (CAZy, 2013). Since the
structure and molecular mechanism of an enzyme are related to its primary structure, this
classification system reflects both structural and mechanistic features (Collins et al., 2005).
Usually, the mechanism used for catalysis, is conserved within each GH family. Overall both
systems provide complimentary information that facilitates understanding of fibrolytic enzymes
especially when one enzyme has more than one activity or when the same activity is present in
different families. Thus, a description of fibrolytic enzymes using both systems will follow to
elucidate their functions. Special focus will be given to Trichoderma spp. and Aspergillus spp.
enzymes since they are the sources of most of the commercially available EFE products (Glass et
al. 2013).
Classification and Functions of Fibrolytic Enzymes
Cellulases and xylanases catalyze stereoselective hydrolysis of the glycosidic bond with
either retention or inversion of the configuration around the anomeric center of the substrate
(Bhat and Hazlewood, 2001). Both reactions are assisted by acid / base catalysis with inversion
occurring by way of a single displacement reaction and retention via double displacement (Bhat
and Hazlewood, 2001). Enzymatic hydrolysis of cellulose is a multi-step reaction that takes place
in a heterogeneous system, in which insoluble cellulose is broken down at the solid–liquid
interface via the synergistic action of endoglucanases and exoglucanases (Andric et al., 2010).
This degradation is accompanied by liquid-phase hydrolysis of soluble intermediates,
cellooligomers and cellobiose, which are hydrolyzed to glucose by the action of β-glucosidase
(Andric et al., 2010).
38
Cellulases
The term “cellulase” refers to a broad group consisting of many fibrolytic enzymes that
hydrolyze cellulose.The following activities are typically included (Paloheimo et al., 2010):
EC 3.2.1.4 (4-β-D-glucan-4-glucanohydrolase; Endoglucanase).This activity catalyzes
the endohydrolysis of internal β-1,4-glycosidic bonds of amorphous and swollen celluloses as
well as cello-oligosaccharides (BRENDA, 2013) but they are generally inactive towards
crystalline cellulose and cellobiose (Bhat and Hazlewood, 2001). End products of endoglucanase
hydrolysis include oligosaccharides of various lengths (Lynd et al., 2002). This activity is present
in GH families 5, 6, 7, 8, 9, 10, 12, 16, 18, 19, 26, 44, 45, 48, 51, 74, 124 and a non-classified
family (CAZy, 2013). As reviewed by Nutt (2006) and Vlasenko et al. (2010), the secretome of
T. reesei contains the following 5 of these families: 1) Endoglucanase type I (GH 7) is the most
abundant accounting for 5-10 % of total cellulase and it also has xyloglucan side-activity. 2)
Endoglucanase Type II (GH 5) is also abundant but has lower activity on substituted cellulose
and β-D-glucans, and has mannase side-activity. 3) EndogluconaseType III (GH 12) lacks a
carbohydrate-binding module (CBM) and can also hydrolyze xyloglucans and xylans. 4)
Endoglucanase Type IV, which no longer exists as it was recently reclassified as auxiliary
activity (AA) 9 and renamed polysaccharide monooxygenase because it acts by an oxidation-
reduction type of reaction (Harris et al. 2010). Although it is no longer regarded as an
endoglucanase, it enhances the activity of the other endoglucanases and seems to require the
presence of cellobiose dehydrogenase for this effect (Bey et al., 2013). 5) Endoglucanase type V
(GH45) is a polyspecific enzyme with lower endoglucanase activity than other families and it
can also hydrolyze glucomannan. The GH45 enzymes have a neutral pH optimum and thus are
widely used in the laundry detergent industry (Koga et al., 2008)
39
EC 3.2.1.91 (4-β-D-glucan cellobiohydrolases, non-reducing end; Exoglucanase II)
and EC 3.2.1.176 (4-β-D-glucan cellobiohydrolases, reducing end; Exoglucanase I).These
cellobiohydrolases act in a processive manner on the reducing or non-reducing ends of cellulose
fibrils, hydrolyzing β-1,4-glycosidic bonds and liberating cellobiose as a major product (Lynd et
al., 2002). Unlike endoglucanases, cellobiohydrolases act on crystalline parts of cellulose chains
(Paloheimo et al., 2010). These enzymes are specific for β-1,4-glycosidic bonds, but are inactive
on cellobiose. Exoglucanase I is found in GH families 7, 9, and 48 and exoglucanase II is present
in GH families 5, 6, 7, and 9. Exoglucanase I is perhaps the most important and best-studied
single enzyme for cellulose hydrolysis because it is secreted copiously by T. reesei and it
comprises up to 60% of the protein in commercial cellulase preparations. However, by itself, it
has little impact on cellulose hydrolysis (Selig et al., 2008). While endoglucanases have an open
substrate-binding cleft or groove, exoglucanases fold to a β-sandwich with extended loops
forming a long, cellulose-binding tunnel, and this partly accounts for their different mode of
action (Paloheimo et al., 2010).
EC 3.2.1.74 (4-β-D-glucan glucohydrolase; Exoglucanase or glucohydrolase).
Glucohydrolases catalyze the hydrolysis of β-1,4-glycosidic bonds in 1,4- β-D-glucans, to
remove glucose units from the non-reducing end (ExplorEnz, 2013). This activity is present in
GH families 1, 3, 5, and 9 (CAZy, 2013). This activity was included among cellulases by
Paloheimo et al. (2010) but Lynd et al. (2002) and (Bhat and Hazlewood, 2001) considered it an
exoglucanase.
EC 3.2.1.21 (β-glucoside glucohydrolases; β-glucosidase).β-Glucosidases can be
classified as either aryl β-D-glucosidases (hydrolyzing aryl-β-D-glycosides exclusively),
cellobiases (hydrolyzing diglucosides and cellooligosaccharides) or β-glucosidases with broad
40
substrate specificities (Bhat and Hazlewood, 2001). This activity is present in GH families 1, 3,
5, 9, 30, 116 and a non-classified family (CAZy, 2013). These enzymes sequentially remove one
glucose unit from either the reducing or the non-reducing end or both ends (Bhat and
Hazlewood, 2001) and they prevent inhibition of cellobiohydrolases (exoglucanases) by their
catalytic end product (cellobiose) (Jalak et al., 2012).
Hemicellulases (Xylanases)
Hemicellulases are composed of a myriad of enzymatic activities that depolymerize grass
arabinoxylans, as well as other types of hemicellulose. The enzymes that depolymerize the xylan
backbone are endoxylanase and β-xylosidase and others, which cleave the substitutions on the
backbone are called accessory enzymes. As described by Collins et al. (2005) hemicellulases
include the following activities:
EC 3.2.1.8 (4-β-D-xylan xylanohydrolase; Endoxylanase). Endoxylanases are
glycosidases that catalyze the endohydrolysis of 1,4-β-D-xylosidic linkages in xylans and yield
xylooligomers (Shallom and Shoham, 2003). Most endoxylanases are specific for unsubstituted
(not branched with acetic acid, glucuronic acid or arabinose) xylosidic linkages of xylans and
release both substituted and unsubstituted xylo-oligosaccharides. In contrast, some
endoxylanases are specific for xylosidic linkages adjacent to substituted groups in the main xylan
chain (Bhat and Hazlewood, 2001). This activity is present in GH families 5, 7, 8, 10, 11 and 43,
and is particularly abundant in GH 10 and 11 (Collins et al., 2005). Usually the presence of
glucuronic acid or arabinofuranose substitutions hinder the binding to and hydrolysis of xylan by
this enzyme, except in some endoxylanases that actually require the substitutions for recognition
and cleavage (Shallom and Shoham, 2003).
EC 3.2.1.37 (4-β-D-xylan xylohydrolase; β-xylosidase).The β-xylosidases are exo-type
glycosidases that catalyze the hydrolysis of 1,4- β-D-xylans (xylo-oligosacharides and
41
xylobiose), and remove successive D-xylose residues from the non-reducing end (BRENDA,
2013).This activity is found in GH families 3,39,43,52, and 54.
EC 3.2.1.55 (α-L-arabinofuranoside, non-reducing end; α-arabinosidase).The α-
arabinosidases catalyze the hydrolysis of terminal non-reducing α-L-arabinofuranoside residues
in α-L-arabinofuranosides, arabinans, arabinoxylans and arabinogalactans, releasing arabinose
(Poutanen, 1988). They are found in GH families 3, 43, 51, 54 and 62. The spatial similarities
between D-xylopyranose and L-arabinofuranose leads to bifunctional xylosidase-arabinosidase
enzymes found in families 3, 43, and 54 (Shallom and Shoham, 2003).
EC 3.2.1.139 (α-glucosiduronase; α-glucuronidase).The α-glucuronidases catalyze the
hydrolysis of α- D-glucuronoside into an alcohol and D-glucuronate (ExplorEnz, 2013). This
activity is exclusively in GH family 67 and has the peculiarity of not hydrolyzing synthetic
substrates (Shallom and Shoham, 2003).
EC 3.1.1.72 (Acetylxylan esterase).The acetylxylan esterases catalyze the deacetylation
of xylans and xylo-oligosaccharides. The enzyme belongs to the carbohydrate esterase (CE)
class, which includes 16 families. Acetylxylan esterases are present in CE families 1, 2, 3, 4, 5,
6, 7, 12, and 15 (CAZy, 2013). Two types of acetylxylan esterase seem to exist, one that acts on
acetylated oligosaccharides and requires the presence of a xylanase and a β-xylosidase to release
acetic acid from polymers and another that does not require other enzymes (Humberstone and
Briggs, 2000). The latter author suggested that since this activity releases acetic acid from
arabinoxylans present in grasses, it could be used to replace silage heterofermentative inoculants
like Lactobacillus buchnerii and thereby reduce dry matter losses caused by the latter during
ensiling while maintaining its beneficial effect on aerobic stability. Acetyl groups can represent
1-3 % of total DM in grasses (Theander et al., 1981).
42
EC 3.1.1.73 (4-hydroxy-3-methoxycinnamoyl-sugar hydrolase; Ferulic acid esterase
and p-coumaric acid esterase).The ferulic acid esterases catalyze the hydrolysis of feruloyl
polysaccharides esterified sugar to release ferulate (ExplorEnz, 2013). Interestingly, no separate
activity for p-coumaric acid esterase activity was found in the ExplorEnz and BRENDA
databases (2013). The activity is likely similar to ferulic acid esterase except that p-coumaric is
released instead. A full profile of the enzyme including its mass, optimal pH and temperature,
and kinetics was published by Borneman et al. (1991) using a bermudagrass cell wall extract, but
not much research has been done on p-coumaric acid esterase. Both enzymes are present in CE
family 1 and in a non-classified family (CAZy, 2013). Ferulic and p-coumaric acid esterases
have been recently the focus of efforts to use them as enhancers of EFE used in ruminant diets
(Beauchemin et al., 2004; Krueger et al., 2008). This is because the ester linkages between
ferulic and p-coumaric acid and arabinose in hemicellulose limit digestibility by ruminal
microbes (Faulds and Williamson, 1994). Though high concentrations of the phenolic acids were
previously considered toxic, recent research has shown that they can be degraded by ruminal
microbes (Jung and Allen, 1995).
Synergy Between Fibrolytic Enzymes
Synergy between cellulases
Cellulase enzyme systems exhibit higher collective activity than the sum of the activities
of individual enzymes, a phenomenon known as synergism. Five forms of synergism have been
reported (Lynd et al., 2002):
Endo-exo synergy. This occurs between endoglucanases and exoglucanases. Randomly
acting endoglucanases generate new cellulose chain ends that serve as starting points for
processive exoglucanases that render the substrate more accessible to endoglucanases (Jalak et
al., 2012).
43
Exo-exo synergy. This occurs between exoglucanases processing from the reducing
(exoglucanase I) and non-reducing ends (exoglucanase II) of cellulose chains. Exoglucanase I
leads to fibrillation, thinning of the cellulose crystal, or narrowing of the crystal end, whereas
exoglucanase II hydrolyzes the cellulose chain less processively, sharpening the crystal trip
(Igarashi et al., 2011). Igarashi et al. (2011) proposed that exoglucanase II creates nicks in
crystalline cellulose, which become starting and end points for exoglucanase I. Therefore, this
can also be considered a type of endo-exo synergism though both enzymes are exoglucanases
(Igarashi et al., 2011).
Exoglucanase - β-glucosidase synergy. This occurs when β-glucosidases relieve
exoglucanases from product inhibition by cellobiose (Jalak et al., 2012)
Endo-endo synergy. This occurs between different types of endoglucanases (Klyosov,
1990). For instance, polysaccharide monooxygenase, previously known as endoglucanase IV,
enhances the activity of other endoglucanases (Bey et al., 2013). This enzyme couples its
reductive activation to the oxidation of cellobioase by cellobiose dehydrogenase (Bey et al.,
2013). This allows polysaccharide monooxygenase e to cleave glycosidic bonds without the
energetically costly step of abstracting a glucan chain from crystalline cellulose (Phillips et al.,
2011)
Swollenin - cellulase synergy. Swollenin is an expansin-like protein that disrupts the
hydrogen bonding between cellulose fibrils and other polysaccharides without producing sugars
from the hydrolysis and thereby reduces cellulose crystallinity (Zhou et al., 2013). In this way,
swollenin allows sliding of cellulose fibers and enlargement of the plant cell wall, and they have
been called amorphogenesis inducers (Saloheimo et al., 2002). They facilitate the action of
44
hydrolytic and oxidative fibrolytic enzymes by facilitating access to the glycosidic linkages by
exoglucanases and endoglucanases (Gourlay et al., 2013)
Synergy between hemicellulases
Efficient and complete hydrolysis of xylan requires the synergistic action of main and
accessory enzymes with different specificities (Coughlan et al., 1993). Two types of synergy
between such enzymes have been described (Coughlan et al., 1993):
Homeosynergy. This occurs between two or more different types of accessory enzymes
or between two or more types of main-chain cleaving enzymes like ferulic acid esterases and α-
L-arabinofuranosidases and endoxylanases and β-xylosidases, respectively.
Heterosynergy. This occurs between accessory and main chain cleaving enzymes (e.g.,
ferulic acid esterases and endoxylanases). Also, synergy between ferulic acid esterase and
swollenin has been demonstrated and it results in greater release of ferulate (Levasseur et al.,
2006). Swollenin seems to have better synergy with xylanases than cellulases (Gourlay et al.,
2013).
Synergy between cellulases and hemicellulases
Hemicellulose is usually more concentrated on the outer surface of cellulose fibers but it
also diffuses into interfibrillar spaces through fiber pores, which act as a physical barrier that
limits accessibility of cellulases to cellulose (Hu et al., 2011). Xylanases alleviate this problem,
improving accessibility of cellulases and consequently increasing the efficiency of the process.
Cellulose access to cellulases is improved by the increased fiber swelling and fiber porosity
caused by the presence of xylanases (Hu et al., 2011).
Carbohydrate-Binding Modules
Carbohydrate-binding modules (CBM), formerly called Carbohydrate-binding domains
are defined as a contiguous amino acid sequence within a carbohydrate-active enzyme with a
45
discrete fold that has carbohydrate-binding activity (CAZy, 2013). The CBM are classified into
70 families based on amino acid sequence similarity (CAZy, 2013). There are CBM that
recognize crystalline and non-crystalline cellulose, chitin, β-1,3-glucans, and β-1,3-1,4-mixed
linkage glucans, xylan, mannan, galactan, and starch (Boraston et al., 2004). Cellulases with a
CBM adsorbed to a greater extent to and increased the rate and extent of crystalline cellulose
hydrolysis when compared with cellulases without CBM (Klyosov, 1990). The CBM affects
binding to the cellulose surface, presumably to facilitate cellulose hydrolysis by bringing the
catalytic domain in close proximity to the substrate (Lynd et al., 2002). Interestingly, ligand
specificity is typically invariant in CBM that recognize crystalline polysaccharides (Boraston et
al., 2004). Many CBM are metalloproteins but the role of metal ions in CBM-ligand interactions
is not well known. However, calcium is reportedly essential for family 35 and 36 CBM
recognition of xylan (Boraston et al., 2004).
Cellulase Kinetics
According to Bansal et al. (2009) the hydrolysis of cellulose, involves more steps than
classical enzyme kinetics due to its heterogeneous nature:
1. Adsorption of cellulases onto the substrate via the CBM.
2. Location of a cleavable bond on the surface of the substrate.
3. Formation of an enzyme-substrate complex.
4. Hydrolysis of the β-glycosidic bond and simultaneous forward sliding of the enzyme
along cellulose chain.
5. Desorption of cellulases from the substrate or repetition of step 4 or 2 / 3 if only the
catalytic domain detaches.
6. Hydrolysis of cellobiose to glucose by β-glucosidase. Additionally, product inhibition
and substrate changes along the course of hydrolysis can affect the steps above.
46
Kinetic models used to describe the enzymatic hydrolysis of cellulose can be divided into
the following five types based on the fundamental approach and methodology used,: Empirical
models, adsorption models, models based on soluble substrates, Michaelis-Menten models, and
fractal models (Aguiar et al. 2013). Empirical models are good at quantifying the effects of
various substrate and enzyme properties on hydrolysis but they cannot be applied at conditions
different to those used for creating them (Bansal et al., 2009). Adsorption models incorporate
adsorbed cellulase concentration into hydrolysis models using in most cases the Langmuir
adsorption isotherm, which has assumptions that may not be valid in all situations (Zhang and
Lynd, 2004). Models based on soluble substrates are limited to the hydrolysis of soluble
cellooligomers (Bansal et al., 2009). Michaelis-Menten kinetic analysis, developed for enzymatic
reactions in ideal aqueous solutions (homogenous reactions) should not be used for analyzing
cellulose hydrolysis due to the heterogeneous nature of cellulose hydrolysis, which occurs with a
space less than 3 dimensions (Wang and Feng, 2010). This is true except for cellobiose
hydrolysis, because cellobiose is soluble (Andric et al., 2010). Fractal kinetics, which occur
when reactions take place in spatially constrained media, have been recently suggested to
analyze cellulose hydrolysis since it can be thought of as a one-dimensional heterogeneous
reaction along a cellulosic fiber (Väljamäe et al. 2003; Bansal et al., 2009). Furthermore, fractal
kinetics must be considered for catalytic reactions involving diffusion of two species (for
bimolecular reactions) on the non-ideal substrate surfaces (with obstacles resulting in partial
segregation; Bansal et al. 2009). The fractal component is correlated with the structural
organization of the substrate. A low fractal component value indicates high levels of de-
polymerization and this concept can be correlated with the extent of cellulose hydrolysis because
this reaction involves a multicomponent water-soluble biocatalytic system acting on a
47
heterogeneous substrate with variable porosity and surface area as obstacles (Aguiar et al., 2013).
The hydrolysis data of chemically-treated sugarcane bagasse was fitted well by a fractal model,
which revealed that alkali washing may not be critical to produce the best substrates for
hydrolysis (Aguiar et al., 2013).
Product and Substrate Inhibition
As with most enzymes, high concentrations of the hydrolytic products of cellulases and
xylanases often inhibit their action. Endoglucanases (Bhat et al., 1989) and most
cellobiohydrolases are inhibited by cellobiose (Wood and McCrae, 1986) but tolerant of glucose
concentrations of up to 100 mM (Bhat et al., 1989). Likewise, xylanases and endoxylanases are
inhibited by high concentrations of xylobiose, but not by xylose (Bhat and Hazlewood, 2001). In
contrast, β-glucosidases are inhibited by glucose and other mono- and disaccharides (Bhat et al.,
1993). The challenge for researchers is to formulate products with ideal proportions of each type
of enzyme to overcome product inhibition and ensure enzyme efficacy.
Glucose directly inhibits β –glucosidases, endoglucanases and exoglucanases (Bhat et al.,
1989; Bhat et al., 1993). However, most of the effects on endoglucanases and exoglucanases are
indirectly from the cellobiose buildup when β–glucosidases are inhibited (Bhat et al., 1993). The
mechanism of glucose product inhibition has been described as mainly competitive, but non-
competitive and mixed inhibition has also been reported (Andric et al., 2010). Since cellobiose is
soluble and because its hydrolysis is simple, β-glucosidase-catalyzed cellobiose hydrolysis and
its product inhibition seem to follow Michaelis-Menten kinetics (Andric et al., 2010).
Cellobiose directly inhibits cellobiohydrolases and endoglucanases (Bhat et al., 1989;
Wood and McCrae, 1986). The mechanism of cellobiose product inhibition has been described
as non-competitive, uncompetitive, competitive, and even as “mixed” inhibition. The variation in
the results obtained on different substrates seem to be related to the differences in the estimation
48
of model parameters used in the different studies rather than substrate type (Andric et al., 2010).
Since T. reesei β-glucosidase is not only more prone to product inhibition than that from A.
niger, but is also scarcer in crude fermentation extracts, addition of surplus β-glucosidase from
A. niger to T. reesei extracts is highly recommended to avoid exoglucanase inhibition (Lynd et
al., 2002).
Interestingly, as substrates, cellulose and cellobiose can cause a pseudo-inhibition of
exoglucanases, endoglucanases, and β-glucosidases (Andric et al., 2010). Cellobiose inhibition
can be solved by adding enough β-glucosidases, as mentioned earlier. Cellulose inhibition is
caused by non-productive binding, especially at high cellulose concentrations (Huang and
Penner, 1991). Furthermore, some β-glucosidases will synthesize cellooligomers by
transglycosylation of cellobiose with glucose (reverse reaction; Andric et al., 2010). Other less
researched inhibitors of cellulases are xylooligomers and xylose (Qing et al., 2010), phenols like
vanillin, syringaldehyde, trans-cinnamic acid, and hydroxybenzoic acid (Ximenes et al., 2010).
Cofactors and Coenzymes
The functional groups of proteins are suited to acid-base catalysis, nucleophilic and
electrophilic catalysis, and in few cases radical initiation but these do not account for all types of
catalytic reactions of enzymes (Broderick, 2001). The most important exceptions are probably
redox reactions and group transfers (Voet et al., 2010). Some enzymes utilize unique properties
of a variety of non-protein molecules and ions to assist in their catalytic reactions (Broderick,
2001). These molecules are known as coenzymes or cofactors. Coenzymes are mostly organic
molecules such as NAD+, typically derivatives of vitamins or bacterial growth factors and they
are usually not covalently bound to the enzyme (Broderick, 2001). So far, only GH of family 4
and 109 require a NAD coenzyme, which remains tightly bound throughout catalysis (Withers
and Williams, 2013). For instance, 6-phospho-β-glucosidase cleaves glycosidic bonds via an
49
elimination mechanism involving NAD (Withers and Williams, 2013). Cofactors are metal ions
required by most enzymes for maintenance of structural integrity (metal-activated enzymes)
and/or catalytic activity (metalloenzymes; Voet et al., 2010). Metal-activated enzymes require a
cation for conformation stabilization in order to achieve maximal activity (Glusker, 2011).
Cofactor cations such as Mn2+, Co2+, Fe2+, Ca2+, and Mg2+ have improved glycoside hydrolase
activity (BRENDA; 2013). In metalloenzymes, cofactors serve as essential substrate templates,
inducers of free radicals, and redox-active cofactors at the enzyme active site (Purich, 2011).
Most of the glycoside hydrolases involved in fiber degradation do not require a cation for
hydrolysis, since they catalyze acid-base reactions (Harris et al., 2010) hence, they are not
metalloenzymes. So far, only a few enzymes mostly grouped in the auxiliary activities family
have been identified as metalloenzymes involved in lignocellulose degradation (CAZy, 2013). In
fact the only cellulolytic enzyme known to require a cofactor is polysaccharide monooxygenase,
which requires copper and catalyzes oxidation-reduction reactions (Quinlan et al., 2011).
Copper-dependent enzymes have also been associated with the de-polymerization of chitin
(Nakagawa et al., 2013).
Exogenous Fibrolytic Enzymes in Dairy Cattle Diets
Use of EFE in dairy cow diets has resulted in equivocal performance responses
(Beauchemin et al., 2004; Adesogan, 2005). In 2002, Wang and McAllister argued that much of
the EFE research had been focused on animal responses to different commercial EFE, yet little
attention had been given to their characteristics and mode of action in ruminant diets. This has
hindered the development of effective EFE for ruminant diets and in fact, product testing of EFE
without a clear understanding of their mode of action has resulted in avoidable setbacks in EFE
development for dairy cattle. Recently, Rosen (2010) analyzed 27 publications on supplementing
EFE to dairy cattle diets with 29 EFE products in a holo-analysis. These EFE were mostly based
50
on β-glucanase, cellulase and xylanase, with minor side-activities of amylase, ferulic acid
esterase, and protease, among others. Dry matter intake and milk yield were enhanced by EFE in
64 % and 63% of the tests and feed conversion ratio in 52%. Milk protein, fat and lactose
concentration were increased in 58, 55, and 51 % of experiments. These results show the
potential of EFE to improve dairy cow performance, but also highlight the variability in the
responses. The following section describes the mode of action of EFE in ruminant diets and
discusses the non-enzymatic factors affecting the response to EFE addition to dairy cattle diets.
Mode of Action of EFE in Ruminant Diets
McAllister et al. (2001) discussed the modes of action of EFE in ruminant diets under the
following categories:
Preingestive effects
This category describes hydrolytic effects of EFE on feeds prior to consumption by
ruminants. When applied to forages or to ruminant diets before feeding, providing the
environment (moisture, pH and temperature) is coducive, EFE initiate fiber hydrolysis via dry
matter (DM) losses (Anderson et al., 2005; Krueger et al., 2008), saccharification (Hristov et al.,
1996; Nsereko et al., 2000) due to partial solubilization of NDF and ADF (Gwayumba and
Christensen, 1997; Krause et al., 1998; Nsereko et al., 2000; Krueger, 2007), and hydrolytic
cleavage of ester linkages that attach phenolic acids in the cell walls to sugars (Anderson et al.,
2005; Krueger et al., 2008). The degree of preingestive hydrolysis achieved depends on the EFE
dose, composition, substrate crystallinity and composition, environmental conditions and the
time that elapses between EFE application and feeding. When applied to a dairy cow total mixed
ration (TMR), EFE act on a significant portion of the diet for several hours before it is consumed
because they are fed at an ad libitum level all day long. Diets of dairy cows in the US typically
have an acidic pH because silage typically represents about half of the diet and has a pH of
51
approximately 4. This acidic pH favors EFE hydrolysis because the optimum pH for most
commercial EFE, which are sourced from Trichoderma spp. and Apsergillus spp. is
approximately 5 (Vicini et al., 2003). However, the optimal temperature for most of such EFE is
50oC (Vicini et al. 2003, Arriola et al., 2011a), therefore the lower ruminal and environmental
temperatures (<40oC) in the US will not allow maximal EFE activity and likely prevent optimal
preingestive hydrolysis by EFE.
Nsereko et al. (2000) demonstrated compelling evidence that applying EFE to feed
causes structural changes that make the feed more amenable to further degradation. In their trial,
EFE application (Multifect Xylanase experimental preparation and Sumizyme X, Monsanto Co.,
St. Louis, MO) improved the digestibility of alfalfa hay (Medicago sativa) in buffered-ruminal
fluid even when substrates were autoclaved and washed to remove enzyme residues and
hydrolysis products, leaving only structural changes as the explanation for the digestibility
improvement. McAllister et al. (2001) reported that EFE application caused appearance of
“digestive pits” in cell walls of barley straw. Collectively, these studies demonstrate that EFE
exert pre-ingestive effects that enhance cell wall utilization.
Ruminal hydrolytic effects
Most of the improvements in forage quality resulting from EFE application were
previously attributed to ruminal effects (Beauchemin et al., 2003), though recent work indicates
that this is not always true (Arriola and Adesogan, 2013). In the rumen, EFE are generally more
stable since some have been shown to be resistant to ruminal proteases. The latter depends on
secondary and tertiary conformation of the protein, glycosylation and the concentrations of
carriers and or stabilizers in the preparation (Hristov et al., 1998; Morgavi et al., 2000a, 2001).
Furthermore, application of EFE to feeds prior to ingestion enhances the adhesion of the EFE to
the substrate, which increases their resistance to proteolysis and prolongs their residence time
52
within the rumen (Fontes et al., 1995). Also, EFE have increased the rate of particle passage in
some (Romero et al., 2013; Feng et al., 1996) though not all cases (Beauchemin et al., 1999;
Yang et al., 1999). According to McAllister et al. (2001), there are two mechanisms by which
EFE influence ruminal fiber utilization:
Direct hydrolysis. The fact that certain EFE can remain active in the rumen indicates
that they can improve digestion through direct hydrolysis of ingested feed within the rumen
(Rode et al., 2001). However, the prevailing EFE application rate will determine the degree to
which the hydrolytic capacity of the rumen is increased by EFE treatment of diets (Beauchemin
and Holtshausen, 2010). At application rates used in typical feeding trials, EFE supply between 5
to 15 % of enzymatic activities present in the rumen (Wallace et al., 2001). Beauchemin and
Holtshausen (2010) cited an unpublished study (Eun and Beauchemin), which showed that
ruminal fluid from cows fed an EFE-treated diet increased the in vitro digestibility of an EFE -
treated and untreated substrate. This indicates that EFE can increase the hydrolytic capacity of
ruminal fluid. The pH and temperature of the rumen (usually 6 and 39oC, respectively) are major
factors determining the activity of EFE. Vicini et al. (2003) reported that two-thirds of enzymatic
activities measured under ideal conditions (pH 5 and temperature 50oC) were lost when enzymes
were assayed at ruminal pH, and a further two-thirds of the reminder were lost at ruminal
temperatures. Therefore, though EFE have the potential to increase the hydrolytic capacity of the
rumen, this effect is often not optimized because many of the EFE activity tests are done under
conditions that overestimate their activity relative to those in the rumen.
Synergism with rumen microbes. Adding EFE to the diet increases the hydrolytic
capacity within the rumen mainly due to increased bacterial attachment (Yang et al., 1999;
Morgavi et al., 2000b; Wang et al., 2001) and stimulation of rumen microbial populations (Wang
53
et al., 2001; Nsereko et al., 2002). Recently, Giraldo et al. (2008) and Gado et al. (2009) showed
that EFE increased microbial growth and production of microbial protein. Furthermore, Morgavi
et al. (2000b) showed that synergism between EFE and rumen microbes enhanced ruminal
cellulose, xylan and corn silage digestion. Synergistic increases in the digestion of wheat straw
NDF were also reported when 28 μg/mL of an EFE (mixture of xylanase, β-glucanases,
CMCase, and amylase) and Fibrobacter succinogenes were applied to the substrate (Wang et al.,
2012). However, a higher EFE dose (280 μg/mL) produced no synergy, probably due to enzyme
crowding of substrate surface (Bommarious et al., 2008). This phenomenon has also been
reported by McAllister et al. (1994) and Morgavi et al. (2004) on alfalfa and corn silage.
Interestingly, in the Wang et al. (2012) experiment, no synergy between the EFE and
Ruminococcus flavefasciens was detected, indicating that the enzymatic activity profile was less
complimentary than that of F. succinogenes (Wang et al., 2012). Applying an EFE to barley
straw prior to incubation increased bacterial colonization of substrate at 4 h but not at 12 or 48 h
(Wang et al., 2012). The initial increase in colonization was attributed to the initial release of
sugars caused by EFE which chemo attracted bacteria and stimulated their growth (McAllister et
al., 1994; Lopez, 2005). Pre-incubating an EFE with barley straw for 24 h at 39oC reduced initial
bacterial colonization (Wang et al., 2012). This may have been because bacterial attachment was
reduced after the extensive hydrolysis of the substrate by the EFE (Hartley and Akin, 1989).
When interpreting these results, it is important to remember that differences in the intestinal flora
between studies are a source of experimental variation that can yield variable results (Bedford
and Apajalahti, 2001). Also, antibiotics can prevent synergy between EFE and ruminal microbes
and reduce the activity of the EFE (Bedford and Apajalahti, 2001).
54
Post-ruminal effects
Adding EFE to ruminant diets can also impact nutrient digestion in the hindgut. In the
abomasum, the glandular compartment in the stomach of ruminants, the pH is closely kept
between 1.6–2.5 (Merchen, 1988). Inactivation of EFE by this low pH and pepsin has been
mentioned as one of the reasons for lower EFE efficacy in ruminants compared to poultry, which
have a higher pH in the proventriculus - gizzard and a faster rate of passage (McAllister et al.,
2001). However, research has shown that a significant portion of EFE activity can be retained
after residence in the abomasum. Hristov et al. (1998) showed that supplementing EFE to the
diet of heifers increased xylanase and endoglucanase activity in the duodenum by 30 and a 5%,
respectively. This along with other reports (Fontes et al., 1995) seems to indicate that xylanases
are more resistant to abomasal conditions than endoglucanases. The increased xylanase activity
in the duodenum decreases duodenal digesta viscosity potentially increasing absorption of
nutrients, as in poultry (Hristov et al., 1998). Increased xylanase activity was also observed in the
large intestine and feces (, 2000) of heifers fed a xylanase EFE. These large intestinal responses
would potentially modify microbial fermentation and alter the microflora distribution and
population (Bedford and Apajalahti, 2001). The latter may result in a greater population of
xylose users, while reducing starch and protein users as in poultry (Bedford and Apajalahti,
2001) as well as an increase in VFA concentration and hence energy supply (Knowlton et al.,
1998).
Microbial cellulose degradation
There are two distinct strategies for degrading cellulose depending on the microbial
requirement for oxygen (Lynd et al. 2002). Since all commercial EFE are produced by aerobic
organisms and the microbes in the rumen are anaerobic, the important differences in their
cellulose degradation strategies are noteworthy.
55
Complexed cellulose systems (Anaerobe strategy). The majority of cellulolytic
anaerobes (bacteria and fungi) have complexed cellulases attached to their cell wall surface
called cellulosomes and consequently they need to attach to the lignocellulosic substrate to be
able to hydrolyze it (McAllister et al., 1994). The anaerobic microorganisms developed this
strategy to be able to efficiently synthesize the needed enzymes without excessive expense of
ATP, which is limited in anaerobic environments (Lynd et al., 2002).
Non-complexed cellulose systems (Aerobe strategy). Due to their aerobic metabolism
and greater ATP production, aerobic cellulose degraders hydrolyze cellulose through the
secretion of copious amounts of enzymes that can be easily recovered from culture supernatants
due to their aerobic metabolism, (Schwarz, 2001). This is the main reason why aerobic
organisms are the preferred commercial EFE sources (Bhat and Hazlewood, 2001). Aerobic
fungi penetrate the lignocellulosic substrate through hyphal extensions, secreting the enzymes in
the cavity created by the hyphae (Ericksson et al., 1990). In this environment loss of enzymes
and products of hydrolysis is likely to be limited (Lynd et al., 2002). However, this is not the
case when EFE from these sources are sprayed on the feed surface and enzymes get diluted on
the large surface of feed particles.
More research is needed to better understand how to achieve and exploit synergy between
these two cellulose digestion strategies in order to enhance fiber utilization in ruminant diets.
Non-Enzymatic Factors Affecting Efficiency of EFE
Many non-enzymatic factors influence the effects of EFE in ruminant diets, and the most
important of these will be discussed in this section.
Manufacturing process
Most EFE preparations are in concentrated, cell-free, spent culture media produced by a
batch fermentation process in submerged or deep-tank bioreactors, mostly with Trichoderma
56
spp., Aspergillus spp., Humicola spp. or Bacillus spp. (Cowan, 1994). These microbial sources
are favored because they secrete large quantities of enzymes, and they are mesophiles (Clarkson
et al., 2001), non-pathogenic and are easy to cultivate on an industrial scale (Paloheimo et al.,
2010). Microbial enzyme sources (production hosts) are classified into wild-type (CMO) and
genetically modified strains (GMO). The CMO produce EFE with multiple activities and their
profiles can be modified by strain development and process optimization. However, enzyme
diversity and expression levels of desired activities can be limited by the host genome.
Therefore, GMO microbes were developed as alternative sources to maximize expression of
desired activities and delete genes coding irrelevant proteins via genetic engineering (Clarkson et
al., 2001; Paloheimo et al., 2010).
The types and activities of EFE produced also depend on growth substrate, temperature,
pH, foaming, aeration, and mixing of the bioreactor (Considine and Coughlan, 1989; Paloheimo
et al., 2010) and these factors can cause batch to batch variations in enzyme activity (Considine
and Coughlan, 1989). Some inducers, like cellulose, need to be added to the culture media in
order to obtain adequate EFE production (Paloheimo et al., 2010) as well as sources of carbon,
nitrogen and phosphorus. Downstream, cells and solids are removed by continuous-flow
centrifugation, filter presses or rotary drum vacuum filters (Gashe, 1992; Paloheimo et al., 2010).
Morgavi et al. (2000a; 2001) reported that stabilizers and preservatives added during
manufacturing increased the survival of EFE in the rumen. Typical stabilizers include NaCl,
glycerol, sorbitol, and propylene glycol and frequently used preservatives include sodium
benzoate, potassium sorbate, and methyl paraben (Paloheimo et al., 2010). Van de Vyver et al.
(2004) and Adesogan (2005) also mentioned that co-factors and natural or artificially-induced
enzyme glycosylation are important for ensuring ruminal stability and function of EFE. These
57
factors may also be important determinants of how long EFE can survive in the gastrointestinal
tract or outdoors, especially when applied to feeds stored for long periods before they are fed.
Influence of pH and temperature
Protein enzymes are polyelectrolytes made up of α-aminoacids often having additional
positive and negative charges on their side chains. Their net charge and catalytically functional
active groups also depends on pH (Purich, 2011). Many enzyme catalyzed reactions involving
proton transfer. Therefore, pH can profoundly influence activity as well as enzyme structure,
catalysis and regulation (Purich, 2011). The effects of variations in pH on the activity of
enzymes is similar to the effects of activators and inhibitors, and the same kinetic methods and
theory can be applied to both (Tipton et al., 2009).
Temperature is the factor affecting activity that is the hardest to interpret since
temperature influences all reactions, including pH-dependent ionization, metal-ligand
interactions, conformation, hydrogen bonding, etc. (Purich, 2011). Most chemical reactions
proceed to a faster velocity as temperature rises since more kinetic energy is imparted to the
reactant molecules resulting in more productive collisions per unit of time (Segel, 1976). The
rate constants in an enzyme-catalyzed reaction will obey approximately Arrhenious law (Laidler
and Peterman, 2009), which indicates that reaction rates often double approximately for every
10oC increase in temperature. However, excessive increases in temperature can also inactivate
enzymes (Laidler and Peterman, 2009). Therefore, most enzymes will permanently lose activities
beyond 70-80 oC, except thermophilic enzymes and some other exceptions (Purich, 2011).
Most commercial EFE products have a pH optimum between 4.0 and 5.0 and a
temperature optimum of 60oC, but great variation exists due to the microbial source(s) of the
EFE (Beauchemin et al., 2004; Svihus, 2010). For instance, T. reesei endoglucanase I had an
optimum pH range on β-glucan of 5.0-7.0 and optimal activity at 65 oC (Paloheimo et al., 2010),
58
whereas for A. niger, the optimum pH of a crude preparation on β-glucan was 4.0-6.0 (Vahjen
and Simon, 1999). In general, fungal cellulases are optimally active between pH 4.0 - 6.0 (Wood,
1985). Endoglucanases, cellobiohydrolases, xylanases, and glucosidases from mesophilic fungi
are optimally active between 40-55oC (Bhat et al., 1989; Coughlan et al., 1993). In addition, the
pH optima range for most xylanases is 4.0-6.0 (de Vries and Visser, 2001). Ruminal conditions
are often very different from these optima, with a relatively constant temperature of 39°C and a
pH approximating 6.0 in North American dairy cattle (Van Soest, 1994). Therefore, many
commercial EFE have suboptimal enzymatic activities in ruminal fluid or in the rumen (Kung et
al., 2002; Vicini et al., 2003). Vicini et al. (2003) measured the pH optima at 50oC and found that
for two EFE sourced from T. reesei , the optimal pH range or value was 5.0-6.0 for xylanases,
4.0-5.5 for endoglucanase, 4.2-5.0 for exoglucanase, 4.5 for β-glucosidase, and 2.6-4.0 for β-
xylosidase. Optimal temperature ranges were 50-60oC for xylanase, 60oC for endoglucanase, and
50oC for exoglucanase. Interestingly, two-thirds of enzymatic activities under ideal conditions
(pH 5 and temperature 50oC) were lost when enzymes were assayed at ruminal pH, and a further
two-thirds of the reminder was lost at ruminal temperatures in the latter study. Arriola et al.
(2011a) also found similar optimal pH and temperatures for the same enzyme activities when 18
EFE from T. reesei, Myceliopthora thermophila, A. niger, Humicola insolens, and A. oryzae
were tested. In the latter study, only 17 % of the18 EFE exhibited optimal endoglucanase and
xylanase activity at pH 6. Therefore, enzyme manufacturers and researchers working on EFE
need to collaborate to develop EFE products that are better suited for ruminant animals. Recent
research in psychrophilic xylanases found that enzymes isolated from Antarctica fungi had up to
10 times more activity than mesophilic enzymes at 5oC and 3 times more at 30oC (Collins et al.,
2005). Similar research should be conducted to identify novel sources of enzymes that act
59
optimally in the rumen. Alternatively, such enzymes or their sources should be genetically
engineered to achieve the same objective.
Stabilizers, preservants and buffers used in EFE commercial preparations can also
influence the enzyme activities (Purich, 2011) and like undisclosed genetic engineering, these
factors may explain differences in activity among enzymes from the same organism.
Specificity of the EFE to the substrate
Enzyme-substrate specificity is a well-known phenomenon (Nelson and Cox, 2008).
Complementing the battery of endogenous fibrolytic enzymes from bacteria, fungi and protozoa
in the rumen (Russell, 2002) with rate limiting exogenous enzymatic activities should be the aim
of EFE addition to ruminant diets. Ideally, to exploit enzyme-feed specificity, each cell wall
polysaccharide should be targeted by one or more appropriate enzymes in EFE preparations.
However, this is difficult to achieve because most EFE are crude or semi crude extracts
comprising many enzymes. Also, the concentration of each cell wall component varies with plant
tissue type and the proportions of each tissue vary with plant species and growth conditions.
When 22 EFE products were tested on alfalfa hay and corn silage, those most effective at
improving the 18-h DMD of alfalfa hay differed from those that were most effective on corn
silage (Colombatto et al., 2003), due to the differences between the composition alfalfa and corn
silage. Alfalfa fiber has more lignin and pectin and less hemicellulose than grasses (Dien et al.,
2006) and corn silage has more starch. Later, Eun and Beauchemin (2008) reported that a meta-
analysis of their EFE studies showed that exoglucanase and endoglucanase activities explained
most of the variation in NDF digestibility of corn silage (55 % for each activity), whereas only
exoglucanase explained most of the variation in NDF digestibility of alfalfa hay (75 %). These
studies reflect enzyme-feed specificity and suggest that cellulases are responsible for 55 to 75%
of the variability in the effects of the tested EFE on digestion of corn and alfalfa silages.
60
Exoglucanases processively cleave cellulose chains at the ends generated randomly by
endoglucanase, to release soluble cellobiose or glucose (Zhang et al., 2006). This enzymatic
depolymerization is the rate-limiting step for cellulose hydrolysis (Zhang et al., 2006).
Interestingly, addition of papain (a cysteine protease) to an EFE synergistically improved
digestibility the corn silage, probably by hydrolyzing structural proteins in the cell wall like
extensins (Eun and Beauchemin, 2007).
Approximately half of the diet of dairy cows is comprised of concentrates particularly
corn and soybean meal, which are starch and protein sources, respectively. Therefore, adding
amylases and proteases could prove useful for improving the utilization of the concentrate
fraction of the diet. Supplementation of a diet with α-amylase from A. oryzae increased milk
yield, reduced milk fat proportion without reducing milk fat yield and tended to increase milk
protein yield in lactating dairy cows (Tricarico et al., 2008). However, adding a protease from
Bacillus licheniformis to the concentrate portion of the diet decreased milk yield and DMI even
though digestibilities of DM, OM, CP, NDF and ADF were increased (Eun and Beauchemin,
2005). The authors attributed the decreased performance to increased ruminal acidosis. Clearly,
more research is needed to develop EFE products with appropriate proportions of cellulase,
amylase, xylanases, and protease activities for improving TMR.
Influence of the animal
Animal factors like low ruminal pH (Mouriño et al., 2000) and high passage rates
(Mertens, 2007) that compromise ruminal fiber digestion are common in high-producing early-
lactation dairy cows. Such cows are typically in negative energy balance and therefore need all
the energy that they can assimilate to cope with the demands of milk production (VanderHaar,
2005). In these critical situations where fiber digestion is depressed and energy requirement is
greatest, response to added EFE is usually greatest (Beauchemin and Holtshausen, 2010).
61
Greater intake, body-weight gain and milk yield responses to EFE treatments have been reported
in early-lactation cows compared to those from their mid- to late-lactation counterparts
(Schingoethe et al., 1999; Knowlton et al., 2002). Consequently, Zheng et al. (2000)
recommended dosing with EFE soon after parturition. This is because currently available EFE
seem to improve only digestibility and animal performance in situations when normal fiber
digestion is compromised.
Effects of the method of application of the EFE
The effects of EFE are influenced by the strategy by which it is applied.
Time of application. Little or no time is required for EFE to attach to substrates
(Beauchemin et al., 2004). Lewis et al. (1996) reported no difference in response to applying
EFE to diets immediately before feeding or 24-h earlier. However, infusing EFE in the rumen
was not effective compared to spraying EFE on the feed (Lewis et al., 1996) suggesting that an
EFE-substrate interaction period was required. Some in vitro studies support the notion that no
interaction period is required (Colombatto, 2000), but others revealed that a 24-h EFE-
pretreatment period was advantageous for improving forage digestibility (Krueger, 2007).
Therefore, more research is needed in this area.
EFE form and application method. Spraying EFE in liquid form on the feed is a more
application method than adding the powder form (Beauchemin et al., 2004) because they don’t
need to be hydrated to be activated and they are more likely to be uniformly distributed in the
feed. Beauchemin et al. (2004) stated that close association between the substrate and EFE is
needed for EFE to bind strongly enough and to prevent its removal and destruction by proteolytic
activity in the rumen. Consequently, spraying is the most widely used method of applying EFE.
Fraction of the diet targeted. Based on their greater cell wall concentration,
theoretically, the forage component of the diet should be the target; however, contradictory
62
reports on the efficacy of this approach exist in the literature. Digestibility and milk production
were improved when an EFE was applied to the concentrate fraction instead of the TMR in two
studies (Rode et al., 1999; Yang et al., 2000). Such counterintuitive results may reflect the
presence of non-fibrolytic activities in the EFE. Dean et al. (2013) reported no differences when
enzymes were applied at ensiling bermudagrass, mixing to the concentrate, bermudagrass silage
or to the TMR but interestingly, EFE have been more effective when applied to dry forages.
Feng et al. (1996) reported that applying an EFE to fresh or wilted smooth bromegrass (Bromus
inermis) had no effect on digestibility but the response increased when the EFE was added to dry
forage. Conversely, Yang et al. (1999) reported no difference between applying EFE to the dry
forage or dry forage plus concentrate. In summary, the studies conducted so far do not give a
clear indication of the best fraction of the diet to treat with EFE. More studies need to be
conducted to compare application of effective EFE to different fractions of the diet.
Rate of enzyme application. Low application rates do not fully exploit the hydrolytic
potential of EFE, especially during short incubation times. Whereas, excessively high application
rates decrease free enzyme binding sites and cause “molecular crowding” of enzymes on
substrate surfaces, which reduces the enzymatic hydrolysis rate (Bommarious et al., 2008). In the
rumen, excessive EFE would compete for available substrate surfaces with other EFE as well as
with ruminal cellulolytic enzymes (Morgavi et al., 2004), and this could decrease fiber
digestibility (Nsereko et al., 2002) and animal performance. Kung et al. (2000) reported that
cows fed a low dose of an EFE tended to produce more milk than those fed a higher dose. Lewis
et al (1996) reported increased FCM yield with an intermediate dose instead of lower or higher
doses of an EFE (2.05 vs. 1.25 and 5.0 mL/kg of DM). In contrast, Schingoethe et al. (1999)
reported no increase in 3.5 % fat corrected milk (FCM) in early lactation cows due to increasing
63
the dose from 0.7 to 1.5 mL/ kg of DM. However, in a holo-analysis, Rosen (2010) did not detect
any effects of EFE dose on voluntary intake, milk yield, and efficiency response variables. These
discrepancies are attributable primarily to differences in the EFE and diets fed to cows in the
studies.
Summary. This review discussed the impact of plant cell wall chemical composition and
anatomy on the digestibility and intake of cattle, the fiber requirement of dairy cattle, the
characteristics, classification, functions and modes of action of fibrolytic enzymes and factors
influencing their ability to overcome the barriers to digestion of forages fed to dairy cattle. In
particular, this review highlighted the variability in the digestibility and animal performance
responses to EFE addition to forages and dairy cow diets and underscored the need for more
research to identify, develop and test more potent and reliable EFE. The review also emphasized
the importance of bermudagrass for dairy production in the Southeast and the critical need to
improve the quality of the forage to increase the level and efficiency of milk production in this
region. The experiments described in Chapters 3, 4, 5, and 6 aimed to address both of the latter
issues. The goal was to improve the potency and reliability of using EFE to improve the
performance of lactating dairy cows fed bermudagrass-based rations. The objectives were to
screen several EFE from multiple companies, to identify the most promising candidates for
improving the NDFD of bermudagrass, to optimize the doses of the latter, to examine if
cofactors could increase their potency and to ultimately test the best candidate in a dairy cow
study to validate its choice. .
64
CHAPTER 3
SCREENING EXOGENOUS FIBROLYTIC ENZYME PREPARATIONS FOR IMPROVED
IN VITRO DIGESTIBILITY OF BERMUDAGRASS HAYLAGE
Background
In the Southeastern US, warm-season grasses are used extensively in cattle production
systems but their high fiber content and low digestibility can limit animal productivity and
consequently profitability (Hanna and Sollenberger, 2007). Improving the quality of warm-
season grasses is a major concern for the dairy industry in the Southeast (Southeast Milk, Inc.,
2011). Bermudagrass (Cynodon dactylon) is the most planted warm-season perennial grass in
southeastern US (10 to 12 million ha; Newman, 2007). Among bermudagrasses, the Tifton 85
cultivar is preferred by southeastern dairy producers because it provides the best combination of
high yields, high quality, and pest resistance and it often is considered a replacement for
expensive imported alfalfa hay (Bernard et al., 2010). Exogenous fibrolytic enzyme (EFE)
application has improved fiber digestion and animal performance in some studies but the results
have not been consistent (Beauchemin and Holtshausen, 2010). Previous studies reported that
adding a fibrolytic enzyme to a corn silage and alfalfa hay - based total mixed ration (TMR)
containing high (48%) or low (33%) concentrate inclusion levels increased NDF digestibility by
6 and 7% and increased feed efficiency by about 6 and 16%, respectively (Arriola et al., 2011b).
However, when the same enzyme mixture was applied to a bermudagrass silage-based TMR,
feed efficiency was unaffected (Queiroz et al., 2011; Bernard et al., 2010). This highlighted the
need for research to develop strategies to increase the hydrolysis of bermudagrass by EFE, in
order to improve its quality as a feed for dairy cattle. This study was the first in a series of
studies (Chapter 3, 4, 5) aimed at improving the potency and reliability of EFE to enhance the
quality of bermudagrass and its utilization by dairy cows. The in vitro studies culminated in an
evaluation of the “best” EFE candidate in a dairy cow study (Chapter 6). The objective of this
65
first study was to identify the most promising EFE preparation for increasing the NDF
digestibility (NDFD) and preingestive fiber hydrolysis of bermudagrass haylage. Additional
objectives were to use proteomic tools to identify differences in the composition of the EFE and
to determine the accuracy of predicting NDF digestibility and measures of preingestive
hydrolysis from EFE activity. The hypothesis was that EFE treatment will increase the in vitro
digestibility, fermentation and preingestive fiber hydrolysis of bermudagrass haylage but the
magnitude of the response will differ with EFE.
Materials and Methods
Bermudagrass Substrate
An established stand of Tifton 85-bermudagrass (Cynodon dactylon) in Alachua County,
Florida was staged in June, 2010 by mowing to a 4-cm stubble and removing the residue. The
field was fertilized subsequently with N (95 kg/ha) and a 4-wk regrowth was harvested on July 7,
2010 by mowing within 1 d to a 4-cm stubble with a CLAAS 3500 mower conditioner (CLAAS
North America, Omaha, NE). The grass was wilted for 2.5 h in the windrow, rolled into round
bales without inoculant addition (John Deere 468 baler, John Deere Co., Moline, IL), wrapped
with 7 layers of 6-mm plastic, and ensiled for 53 d. Ensiled bermudagrass was chosen over hay
since it is more widely used by the Florida dairy industry because humid weather and frequent
rainfall hinders proper drying of hay prior to bailing. Representative haylage samples were cored
from the bales, dried at 60oC for 48 h, and ground to pass a 1-mm screen using a Wiley mill
(Arthur H. Thomas, Philadelphia, PA). The DM, OM, NDF, ADF, ADL, and CP concentrations
of the haylage were 49.4, 93.5, 68.1, 34.2, 3.7, and 18.7%, respectively (DM basis).
Enzymes
In an initial screening study, effects of 18 EFE from 5 companies on in vitro 24-h NDFD
were examined in quadruplicate in each of two in vitro runs (data not shown). The 12 most
66
promising EFE candidates were further evaluated in this study. Their enzymatic activities,
protein concentrations, application doses, and microbial sources are shown in Table 3-1.
Application rates were suggested by the respective manufacturers. Endoglucanase (EN;
enzyme commission, E.C.3.2.1.4), exoglucanase (EX, E.C. 3.2.1.91), xylanase (XY, E.C.
3.2.1.8) and β-glucosidase (BG, E.C. 3.2.1.21) activities were quantified using
carboxymethylcellulose, avicel, oat-spelt xylan, and cellobiose as the substrates, respectively
(Wood and Bhat, 1988). Ferulic acid esterase (FE, E.C. 3.1.1.73) activity was measured using
ethyl ferulate as the substrate (Lai et al., 2009). All activities were measured at 39oC and pH of 6
to mimic ruminal conditions in a lactating dairy cow fed a typical TMR in the US. Protein
concentration was measured using the Bio-Rad Protein Assay (Bradford, 1976) with bovine
serum albumin as the standard (Bio-Rad Laboratories, Hercules, CA).
EFE effects on In vitro ruminal digestibility (Experiment 1)
All EFE were evaluated with a 24 h in vitro ruminal digestibility assay (Goering and Van
Soest, 1970) using the bermudagrass haylage as the substrate. As described by Krueger et al.
(2008), amounts of EFE corresponding to the respective application doses (Table 3-1) were
diluted in 2 mL of 0.1 M citrate–phosphate buffer (pH 6) and added to 0.5 g of substrate (in
quadruplicate) in a 100-ml polypropylene tube fitted with a rubber stopper containing a one-way
gas release valve. The Control treatment consisted of only the buffer. Tubes were tapped gently
to ensure proper mixing of the EFE solution with the substrate. The mixtures were first incubated
at 25°C for 24 h. Subsequently, 52 ml of buffered-ruminal fluid were added and the tubes were
incubated for a further 24 h at 39oC in a forced-air incubator. The fermentations were terminated
by placing tubes on ice. Tube contents were filtered through previously dried (60oC for 48 h) and
weighed 125-mm Whatman no. 451 paper (Fisher Scientific, Pittsburgh, PA) and filtrate samples
67
were retained for further analysis. Residue samples were oven dried at 60oC for 48 h and
weighed. The ruminal fluid collection protocol was approved by the University of Florida
Animal Care Research Committee. The ruminal fluid was filtered through four layers of
cheesecloth prior to inoculation and it was representatively aspirated from two non-lactating,
non-pregnant ruminally-cannulated Holstein cows 3h after consuming a ration of coastal
bermudagrass ad libitum supplemented with corn (0.45 kg), cottonseed hulls (0.46 kg), soybean
meal (0.90 kg), and a vitamin-mineral mix (35.8 g, DM basis). All tubes and artificial saliva
were pre-warmed (39oC) before ruminal fluid addition. Each run was repeated three times. Two
enzyme blank tubes per treatment containing the EFE and no substrate were included to correct
for the effects of EFE. Dried residues were analyzed for NDF, ADF, and ADL (Van Soest, 1991)
sequentially using an ANKOM 200 Fiber Analyzer (ANKOM, Macedon, NY). Hemicellulose
(HEM) was calculated as the difference between NDF and ADF. Cellulose (CEL) was the
difference between ADF and ADL. Residue weights and their NDF, ADF, and ADL
concentrations were used to calculate in vitro true DM, NDF, HEM, and CEL digestibility
(DMD, HEMD, and CELD). The buffered ruminal fluid in each tube was measured for pH
(Accumet XL25 pH meter, Fisher Scientific, Pittsburgh, PA), acidified with 50% H2SO4 (1% v/v
of ruminal fluid sample), and centrifuged at 8,000 × g for 20 min. The supernatant was frozen (-
20oC) and subsequently analyzed for concentrations of VFA (Muck and Dickerson, 1988) using
a Merck Hitachi Elite LaChrome High Performance Liquid Chromatograph (HPLC) system
(Hitachi, L2400, Tokyo, Japan) and a Bio-Rad Aminex HPX-87H column (Bio-Rad
Laboratories, Hercules, CA). Ammonia-N was determined with a Technicon Auto Analyzer
(Technicon, Tarrytoen, NY) and an adaptation of the Noel and Hambleton (1976) procedure that
involved colorimetric N quantification.
68
EFE Effects on Preingestive DM and Fiber hydrolysis (Experiment 2)
The EFE were added to bermudagrass haylage and the mixture was incubated at 25oC for
24 h without ruminal fluid to simulate preingestive hydrolytic effects of the EFE. Enzyme-
substrate mixtures were prepared as described in Experiment 1 without adding buffered-ruminal
fluid. Additional exceptions were that the incubations were in 50-mL culture tubes and sodium
azide (0.02% w/v) was added as an antimicrobial agent to the 2 mL of buffered EFE solution to
prevent substrate hydrolysis by microbes (Krueger et al., 2008). Two blank tubes per treatment
containing no substrate were included to correct for effects of the EFE. After the incubation, 30
mL of double-distilled water was added to each tube. Tubes were shaken for 1 h at 260
oscillations/min with an Eberbach Reciprocating Shaker Model 6000 (Eberbach corporation,
Ann Arbor, MI). Tube contents were filtered subsequently through previously dried (60oC for 48
h)and weighed 125-mm Whatman 451 filter paper (Fisher Scientific, Pittsburh, PA) and filtrate
samples were frozen (-20 oC) for further analysis. Residues were dried at 60oC for 48 h,
weighed, and analyzed for DM, NDF, ADF and ADL as previously described. Residue and
sample dry weights and their concentrations of DM were used to estimate DM losses. Filtrate
samples were thawed and analyzed for water-soluble carbohydrates (WSC; DuBois et al., 1956)
and for ferulic (FER) and p-coumaric acids (COU; Bio-Rad, 2011) using the High Performance
Liquid Chromatograph system described above. Cellobiose, glucose, xylose, and arabinose were
determined in the filtrate of EFE treatments that increased concentrations of WSC. Inositol was
used as an internal standard (Bach-Knudsen and Li, 1991) with an HPX-87P column (Bio-Rad
Laboratories, Hercules, CA) equipped with a refractive index detector. Nanopure water was the
mobile phase.
69
Proteomic Identification and Quantification of Proteins in Select EFE (Experiment 3)
To understand why the EFE had different effects on NDFD, proteomic assays were used
to identify and compare the relative ratio of proteins in the least (9C) and the second (11C) most
effective EFE to the most effective EFE (2A) at improving NDFD. Triplicate samples of EFE
2A, 9C, and 11C were analyzed as described by Silva-Sanchez et al. (2013) using Isobaric tags
for relative and absolute quantitation (iTRAQ) - liquid chromatography-mass spectrometry -
based quantitative proteomics. All analyses were conducted at the Proteomics Division of
University of Florida Interdisciplinary Center for Biotechnology Research. Proteins were
purified following the procedure described by Hu et al. (2013). The identification and analysis of
proteins were performed using ProteinPilotTM Software 4.5 (AB SCIEX, Framingham, MA;
2012). The database was the National Center for Biotechnology Information (NCBI;
http://www.ncbi.nlm.nih.gov; August 7, 2013) for Trichoderma reesei and Aspergillus spp. The
searching parameters were set as iTRAQ peptide label, cysteine alkylation with methyl
methanethiosulfonate, trypsin digestion, identification focus for biological modifications and
BIAS modification The unused score threshold was set to > 1.3 (equivalent to 95% confidence or
better; Silva-Sanchez et al., 2013).
Statistical Analyses
A randomized complete block design with four replicates per treatment and three runs
(blocks) was used to determine effects of EFE preparations on in vitro digestibility and
fermentation measures in Experiment 1.
The model used to analyze digestibility and fermentation data was:
Yijk = µ + Ti + Rj + TRij + Eijk
Where:
µ = general mean
70
Ti = effect of EFE i
Rj = effect of run j
TRij = effect of the EFE i × run j interaction
Eijk = experimental error
A completely randomized design with four replicates per treatment was used to determine
effects of EFE preparations on preingestive DM and fiber hydrolysis in Experiment 2.
The model used to analyze preingestive hydrolysis data was:
Yij = µ + Ti + Eij
Where:
µ = general mean
Ti = effect of EFE i
Eij = experimental error
The GLM procedure of SAS v.9.1 (2012) was used to analyze the data. The least-square
means for the digestibility, fermentation, and preingestive hydrolysis measures were compared to
the corresponding Controls using the Dunnett’s test (Dunnett, 1955). Significance was declared
at P < 0.05 and tendencies at P > 0.05 < 0.10.
Multiple regression relationships between EFE activity and either digestibility data from
Experiment 1 or fiber hydrolysis measures from Experiment 2 were examined using the stepwise
multiple regression procedure of SAS. Model overfitting was prevented by keeping the Mallow’s
C(p) criterion close to the number of regressors plus one. Protein concentration was not used as a
predictor because it was highly correlated with enzymatic activities, potentially causing
multicollinearity.
71
In Experiment 3, a Student’s t-test was used to measure the significance of the relative
ratio of the proteins in EFE 11C or 9C to those in 2A with ProteinPilot Software 4.5 (AB SCIEX,
2012). The degrees of freedom were the number of distinct peptides within the protein evaluated
minus 1 (AB SCIEX, 2012). Quantitation was based on at least three unique peptides (Silva-
Sanchez et al, 2013).
RESULTS AND DISCUSSION
Experiment 1:
EFE effects on digestibility measures
Since both HEM (NDF-ADF) and CEL (ADF-ADL) concentrations are calculated by
difference, changes in ADFD or ADLD will influence the HEMD and CELD values,
respectively. This needs to be borne in mind when interpreting HEMD and CELD results.
Effects of EFE treatment on in vitro true digestibility of DM and fiber in ruminal fluid are
shown in Table 3-2. Compared to the Control, treatment with 6 EFE increased DMD (%, 53.8 to
54.9 vs. 52.0; P < 0.05), 9 increased NDFD (%, 37.8 to 40.4 vs. 35.6; P < 0.05), 9 increased
HEMD (%, 35.3 to 38.0 vs. 33.0 P < 0.05), 5 increased ADFD (%, 41.3 to 42.7 vs. 38.7; P <
0.05), and 7 increased CELD (%, 44.4 to 45.8 vs. 41.8; P < 0.05). The EFE-mediated increases
in NDFD of bermudagrass haylage agree with previous results of EFE on bermudagrass silage
(Dean et al., 2005) and hay (Krueger et al., 2008; Romero et al., 2013). Enzymes seemed to be
more effective at improving the digestibility of the hemicellulose fraction than the ADF and
cellulose fractions. This may have been because most of the EFE had more xylanase activity than
cellulase activity (endoglucanase, exoglucanase, and β-glucosidase, Table 3-1). The untreated
ADF and cellulose fractions were more digestible than the HEM and NDF fractions. Likewise,
after 48-h incubations of bermudagrass (Dean et al., 2008) and bahiagrass (Barton et al., 1976) in
buffered-ruminal fluid, the ADF fraction was more digestible than the NDF fraction. Conversely,
72
Mandebvu et al. (1999) and Romero et al. (2013) reported greater NDF than ADF digestibility
after a 48-h incubation of bermudagrass in buffered-ruminal fluid and noted that the difference
increased with increasing maturity of the forage as did Romero et al. (2013). Usually, the ADF
fraction of forages is less digestible than the hemicellulose fraction. Consequently, ADF has
been used as an indicator of forage digestibility (Van Soest, 1987). However, care must be taken
when interpreting digestibilities of fiber fractions of warm-season grasses because unlike cool-
season grasses, the fiber digestibility of warm-season grasses does not relate to their fiber
fractions (Van Soest, 1994). The extent of digestibility of fiber is limited by extent of
lignification and fiber is only secondarily related to digestibility through its association with
lignin (Van Soest, 1987). Lignin is covalently bound to hemicellulose (Ralph et al., 1995) but
due to different solubilization properties in the detergent system, it is recovered as ADL from the
ADF fraction (Van Soest et al., 1991). Degree of lignification is affected by changes in
temperature, daylength, fertilization, water availability and stress (Van Soest, 1987). In tropical
areas, temperature is seasonally more constant and day length is less variable and consequently
as opposed to the case in cool-season grasses, cellulose and lignin concentrations in warm-season
grasses are not correlated (Van Soest, 1987). Thus, fractions like ADF should not be used as
indicators of indigestible fiber in warm-season grasses (Van Soest, 1987). Most of the enzymes
that increased NDFD increased HEMD except 17D, which only increased ADFD and CELD.
This may have been because of differences in the component proteins of 17D versus those of the
others as well as its low xylanase to cellulase (sum of endoglucanase, exoglucanase and β-
glucosidase) ratio. Although EFE 4A, 13D, and 14D also had a low xylanase to cellulase ratio,
they were sourced from different organisms (M. thermophila, A. oryzae, and A. acuelatus,
respectively). Therefore, EFE 17D probably had other fibrolytic activities more suited for
73
digesting ADF and CEL. Applying EFE to 4A, 5A and 15D improved HEMD but not CELD,
perhaps reflecting the relatively low endoglucanase and exoglucanase activities of these EFE.
Oba and Allen (1999) stated that a 1% unit increase in NDFD of grasses or legumes will result in
increases of 0.17 and 0.25 kg/d in DM intake (DMI) and 4 % fat-corrected milk (FCM) yield,
respectively in dairy cows. If this assumption holds true for bermudagrass, the increase in NDFD
due to application of EFE 2A (%, 40.4 vs. 35.6; P < 0.05) potentially represents increases in
DMI and 4% FCM of 0.8 and 1.2 kg/d in lactating dairy cows, respectively.
Accuracy of predicting digestibility measures from EFE activities
The stepwise multiple regression analysis (Tables 3-3 and 3-4) showed that all
digestibility measures were poorly predicted (R2 < 0.06) by enzyme activity measures. The only
relationships that existed (P < 0.01) were those between β-glucosidase and DMD (P =0.005; R2=
0.05), ferulic acid esterase and NDFD (P = 0.01; R2 = 0.05), ferulic acid esterase and HEMD (P
= 0.05; R2= 0.03), β-glucosidase and ADFD (P = 0.01; R2= 0.06) and endoglucanase plus
exoglucanse and CELD (P =0.02; R2 = 0.06). Eun and Beauchemin (2008) reported that the
DMD and NDFD of corn silage were poorly predicted by endoglucanase (R2= 0.08 and 0.09)
though exoglucanase gave slightly more accurate predictions (R2= 0.12 and 0.34), respectively.
Differences in the ability of exoglucanase to predict the digestibility measures in the latter study
and this one are probably attributable to differences in composition and digestibility of the
forages and EFE used. Nevertheless, the generally poor predictions indicate that enzyme
activities do not accurately predict their effects on bermudagrass digestibility. Since NDF
digestibility is correlated to DMI and milk production (Oba and Allen, 1999), it is unlikely that
enzyme activities can accurately predict such measures of animal performance. As such enzyme
activities should be used for enzyme characterization but not to select the best candidates for
improving forage digestibility or animal performance.
74
EFE effects on fermentation measures
Effects of EFE treatment on ruminal in vitro fermentation measures are shown in Table
3-5. Compared to the Control, 6 EFE had greater total VFA concentration (TVFA, mM, 59.1 to
61.2 vs. 55.4; P < 0.05), 5 had greater acetate concentration (mM, 36.4 to 36.9 vs. 33.5; P <
0.05), 6 had greater propionate concentration (mM, 11.2 to 12.2 vs. 10.4; P < 0.05), 4 had greater
butyrate concentration (mM, 5.04 to 5.23 vs. 4.70; P < 0.05), and 4 had lower acetate to
propionate ratio (A:P, 3.03 to 3.16 vs. 3.24; P < 0.05). In addition, 2 had pH values that were
greater (%, 7.33 to 7.34 vs. 7.28 ; P < 0.05) or lower (%, 7.20 to 7.21 vs. 7.28; P < 0.05), 2 had
greater concentrations of isobutyrate (%, 2.60 to 2.62 vs. 2.17; P < 0.05), and one had greater
concentrations of isovalerate (4A, 2.36 vs. 2.21%; P < 0.05), valerate (17D, 3.21 vs. 2.66%; P <
0.05), and NH3N (5A, 40.2 vs. 38.6, mg/dL; P < 0.05). In most, but not all cases, increased in
vitro digestibility due to EFE treatment led to increased TVFA concentration, reflecting the use
of carbon skeletons from degraded substrate for microbial growth (Owens and Goetsch, 1988).
The increased TVFA concentration is important for dairy cattle since VFA provide about 70% of
the caloric requirements of ruminants (Bergman, 1990). Enzyme treatment seemed to increase
propionate concentration to a greater degree than acetate concentration. For instance, EFE 4A
increased propionate concentration by 17% but only increased acetate and butyrate
concentrations by 9 and 7%, respectively. This response reflects the hydrolysis of fiber fractions
into WSC, which are key fermentable substrates. When levels of sugars available for
fermentation are high in the rumen, a shift in fermentation pattern from acetic to propionic acid
occurs to dispose of excess reducing power (France and Dijkstra, 2005). In most cases, EFE-
mediated increases in propionate proportion resulted in lower A:P ratios. These results agree
with those of Colombatto et al. (2003) on EFE-treated pure cellulose and xylan and Eun and
Beauchemin (2007) on EFE-treated alfalfa hay. Supplementation with soluble sugars also
75
increased propionate concentration and decreased A:P ratio in sheep fed ad libitum diets rich in
easily fermentable carbohydrates (Demeyer, 1991). Increased ruminal propionate concentration
would increase the availability of gluconeogenic substrates in the liver of cows and spare
glycogen, gluconeogenic amino acids and glycerol for other functions. The increased availability
of gluconeogenic substrates could increase glucose availability for lactose synthesis in the
mammary gland, which could potentially increase milk yield (VandeHaar, 2005). Furthermore, a
decreased A:P ratio would reduce H+ loss as methane, which would decrease emission of
greenhouse gasses (Russell, 2002) and reduce energy losses as methane by the cow. In
agreement, Arriola et al. (2011b) and Chung et al. (2012) reported that EFE treatment decreased
the A:P ratio and methane production by lactating dairy cows. The increase in butyrate
concentration due to treatment with EFE 1A, 4A, 13D, and 17D could stimulate cell proliferation
and epithelial growth in the rumen tissues, thus increasing VFA absorption (Gorka et al., 2009).
Also, butyrate provides building blocks for de novo synthesis of fat in the mammary gland
(Mohammed et al., 2011). That EFE 4A increased concentrations of isobutyrate and isovalerate
may have aided fiber digestibility because they are required for optimal growth of ruminal
cellulolytic bacteria (Liu et al., 2009). Enzymes had few and minor effects on NH3N
concentration and pH. This may have been because the in vitro artificial saliva used contained
high concentrations of NH3N and buffers, which may have prevented detection of EFE effects on
these measures.
Experiment 2:
EFE effects on measures of preingestive hydrolysis
Table 3-6 shows effects of EFE application on measures of preingestive cell wall
hydrolysis in Experiment 2. Compared to the Control, 2 EFE increased loss of DM (%, 24.3 to
24.4 vs. 22.0; P < 0.05), 3 increased NDF hydrolysis (%, 62.8 to 64.4 vs. 67.3; P < 0.05), 5
76
decreased concentrations of HEM (%, 29.3 to 32.5 vs. 33.8; P < 0.05), none affected ADF or
cellulose concentrations, and 8 and 6 increased concentrations of FER (μg/g, 210 to 391 vs. 198;
P < 0.05) and COU (μg/g, 170 to 203 vs. 162; P < 0.05), respectively. Furthermore, 10 increased
saccharification (release of WSC%, 2.68 to 5.85 vs. 2.28; P < 0.05) but the specific sugars
released differed with the EFE as evidenced by the fact that 6, 9, 9, and 7 EFE released more
(mg/g) cellobiose (0.24 to 2.25 vs. 0; P < 0.05), glucose (5.3 to 23.0 vs. 4.3; P < 0.05), xylose
(0.11 to 2.15 vs. 0.01; P < 0.05), and arabinose (0.30 to 1.08 vs. 0.12; P < 0.05) than the
respective Controls. Others (Hristov et al., 1996; Krueger et al., 2008) also reported that EFE
application increased the preingestive hydrolysis of barley silage and bermudagrass hay,
respectively. This indicates that several of the EFE can hydrolyze bermudagrass fiber prior to
consumption of the forage, suggesting that adding them to feeds during storage may be
beneficial.
As expected, EFE were more effective at hydrolyzing hemicellulose than cellulose. This
is important because hemicellulose accounts for approximately half of the NDF in tropical
grasses (Van Soest, 1994), and it was the fiber fraction with the least ruminal digestibility in the
Control samples. Since fiber detergent analysis were done in sequence and included fiber-bound
ash except for ADL, it is important to note that some acid detergent soluble ash might have
contributed to increase HEM concentration, although this contribution should have be the same
across treatments. To facilitate further NDF hydrolysis, addition of more endo and exoglucanase
and β-glucosidase activities to the EFE preparations may improve their ability to hydrolyze
cellulose and consequently ADF and digestibility. Hydrolysis of fiber enhances the improvement
of ruminal digestibility by EFE because it creates more attachment points for ruminal bacteria
(Wang et al., 2001). Greater fiber hydrolysis also potentially could increase DMI by reducing gut
77
fill. Furthermore, the EFE-mediated increase in release of WSC (by up to 157%) via cell wall
hydrolysis will increase the supply of soluble sugars that act as gluconeogenic fermentation
substrates and chemo attractants for bacteria, which reduce the lag time prior to feed digestion in
the rumen (Lopez, 2005). Analysis of the composition of the WSC released by fiber hydrolysis
revealed that degradation of cellulose by EFE was evidenced by the release of glucose (up to
435%) and cellobiose (22,400%) from the cell walls. Degradation of hemicellulose by EFE
hydrolysis resulted in much greater concentrations of xylose (21,400%) and arabinose (800%)
than respective values for the Control. These results agree with Anderson et al. (2005) who
reported similar increases in the quantity of sugar released from Tifton 85 bermudagrass hay that
was incubated with cellulases for 72 h.
Release of FER and COU from cell walls were increased by up to 49.4 and 25.3%,
respectively by EFE treatment. Similar results were obtained by Anderson et al. (2005) on Tifton
85 bermudagrass hay. The greater release of FER and COU from bermudagrass would also
improve the accessibility of the substrate to ruminal microbes by removing well-known digestion
barriers (Ralph et al., 1996). Early reports indicating toxicity of ferulic acid to ruminal microbes
have been disproved by the ability of ruminal microbes to metabolize phenolic acids (Jung and
Allen, 1995). Ferulic acid also seems to have potential to improve human health. Soberon et al.
(2012) demonstrated that orally providing pure ferulic acid (150 g single-dose) to dairy cows did
not affect DMI or milk yield and composition but 0.02% of the ferulic acid dosed appeared in
milk 6.5 h after dosing. Some ferulic acid appeared as hippuric acid, which can be transformed to
benzoic acid during cheese manufacturing, enhancing its flavor and stability (Soberon et al.,
2012). Once in the milk, ferulic acid can improve the functionality of milk due to its antioxidant,
anticancer, and antibacterial activities (Soberon et al., 2012).
78
Prediction of measures of preingestive hydrolysis from enzymatic activities
The stepwise multiple regression analysis (Table 3-7) showed that the accuracy of
predicting NDF concentration from EFE activity was moderately good (R2 = 0.62; P < 0.001)
and endoglucanase accounted for more (P < 0.01; R2= 0.33) of the variability in NDF
concentration than other predictors, (exoglucanase and β-glucosidase, P < 0.10; R2= 0.13 and
0.15, respectively). As endoglucanase activity increased, NDF hydrolysis decreased such that a
1% increase in NDF hydrolysis resulted from a 1.3 unit increase in endoglucanase activity. In
contrast, exoglucanase and β-glucosidase had negative correlations with NDF hydrolysis and
much greater changes in the respective activities (481 and 84.7) were required to decrease NDF
hydrolysis by 1%. Water-soluble carbohydrate concentration was accurately predicted (P <
0.001; R2 = 0.95) by a multiple regression model in which endoglucanase accounted for much
more of the variability (P < 0.01; Partial R2= 0.65) than other predictors (P < 0.01, R2 < 0.07).
Endoglucanase and xylanase activities were positively correlated whereas exoglucanase and β-
glucosidase activities were negatively correlated with WSC concentration. A 1% increase in
WSC concentration resulted from a 0.87 unit increase in endoglucanase activity. As for WSC,
FER was predicted accurately (R2 = 0.99; P < 0.001) by a multiple regression model in which
most of the variability was due to xylanase (P < 0.03; Partial R2 = 0.71) followed by
endoglucanase (P < 0.01; Partial R2= 0.17). Other predictors collectively accounted for little (R2
= 0.14) of the variability in FER. As for WSC, endoglucanase and xylanase activities were
positively correlated whereas exoglucanase and β-glucosidase activities were negatively
correlated with FER concentration. It was interesting to note the absence of a relationship
between ferulic acid esterase and FER concentration. This suggests that ethyl ferulate, the
substrate on which FER activity was measured is not an ideal substrate to examine release of
FER from bermudagrass. Surprisingly, xylanase was the best predictor of FER, which suggests
79
that xylanases plays an important role in release of FER from plant cell walls. This is because
xylanases are needed prior to catalysis by esterases that cleave the ester linkages between
arabinose and FER in the cell wall and thereby releasing FER (Faulds, 2010). The role of the
xylanases may be to allow accessibility of esterases to the ester linkages in the cell wall after
releasing the feruloylated oligosaccharides (Faulds, 2010). Ferulic acid esterase as well as
xylanase activity may need to be measured to indicate the potential of EFE to release FER from
grass cell walls. The validity of these postulations should be examined in future research.
This study examined the total release of FER from bermudagrass, rather than the release
of FER that was ester-linked to arabinose or ester / ether-linked to lignin. Theoretically,
commercially available EFE can only release ester-linked ferulic acid from cell walls (Soberon et
al., 2012). Krueger et al. (2008) reported that treatment with an EFE from Humicola sp.
increased the release of ester-linked FER from Pensacola bahiagrass (Paspalum notatum) but not
from Coastal or Tifton 85 bermudagrass. Interestingly, increased release of ether-linked FER
from Tifton 85 bermudagrass also was reported but the reason for this unusual occurrence was
not discussed. Etherase enzymes are required to hydrolyze ether linkages and release ether-
linked FER from cell walls but they are produced rarely by fungi. Mathieu et al. (2013) detected
that no β-etherase activity from 26 fungal strains (including Humicola grisea, Aspergillus sp. and
Trichoderma viride) within three ecological groups (white, brown, and soft rot fungi) and
concluded that extracellular β-etherases are rare and dispensable activities among wood-decaying
fungi. To my knowledge, no other publication has reported β-etherase activity in the secretome
of fungi.
80
Experiment 3: Proteomic Identification and Quantification of the Relative Ratio of Less to
More Effective EFE
The results of the proteomic identification and analysis of the relative ratio of the most
effective (2A) and least effective (9C) EFE at improving NDFD revealed that both EFE had
similar quantities of endo-1,4-β xylanase, β-glucosidase I, amidase, and endoglucanase II with a
(carbohydrate binding module) CBM1 (Table 3-8). These activities catalyze the hydrolysis of 1-
4 β-D-xylosidic linkages in xylans, β-D-glucosides, monocarboxylic acid amide, and internal β-
1,4-glycosidic bonds in cellulose (with mannase side-activity), respectively (CAZy, 2013). The
CBM are amino acid sequences within some carbohydrate-active enzymes that aid the binding
process to carbohydrates, improving catalysis (Boraston et al., 2004). The EFE 2A had 10 times
more endoglucanase III, 17 times more acetylxylan esterase with CBM1, 33 times more xylanase
III, 25 times more β-xylosidase, 7.69 times more polysaccharide monooxygenase with CBM1,
and 3 times more swollenin compared to 9C. These activities (except swollenin and
polysaccharide monooxygenase) catalyze the hydrolysis of internal β-1,4-glycosidic bonds in
cellulose (xylanase side-activity), acetyl groups from xylan, of 1-4 β-D-xylosidic linkages in
xylans, and β-D-xylosides, respectively. Swollenin is an expansin-like protein that reduces
cellulose crystallinity by disrupting the hydrogen bonding between cellulose fibrils and other
polysaccharides without producing detectable quantities of sugars (Zhou et al., 2013). This
facilitates the action of hydrolytic and oxidative fibrolytic enzymes by giving exoglucanases and
endoglucanases access to glycosidic linkages (Gourlay et al., 2013). Polysaccharide
monooxygenase (previously known as endoglucanase IV) enhances the activity of the other
endoglucanases and couples its reductive activation to the oxidation of cellobiose by cellobiose
dehydrogenase (Bey et al., 2013). This allows cellobiose dehydrogenase to cleave glycosidic
bonds without the energetically costly step of abstracting a glucan chain from crystalline
81
cellulose (Phillips et al., 2011).The only advantages 9C had over 2A were that it had 28 times
more rhamnogalacturonan acetylesterase and 20 times more β-galactosidase, which degrade
pectin (rhamnogalacturonan) and xyloglucans, respectively. Evidently, these activities were not
crucial to degradation of the cell walls, partly because pectin is readily digestible in the rumen
(Hatfield et al., 2007) and grasses like bermudagrass have low pectin concentrations (Hatfield et
al., 1999a). More research is needed to evaluate the potential benefits of adding more of these
novel auxiliary proteins to the EFE cocktails used in animal nutrition. These results also suggest
that key EFE proteins for fiber degradation included all or some of the following: xylanase III
(GH10), β-xylosidase (GH3), acetylxylan esterase (CE5) with carbohydrate binding module 1
(CBM1), endoglucanase III (GH12), polysaccharide monooxygenase (AA9) with CBM1 and
swollenin. It is interesting to note that most of the enzyme activities that were more abundant in
EFE 2A (xylanase III, β-xylosidase and acetylxylan esterase) are typically important for
hemicellulose degradation. This study therefore supports the notion that effective EFE products
affect the hemicellulose fraction more than other fiber fractions in bermudagrass (Romero et al.,
2013).
Relative to EFE 11C, which also was very effective at increasing NDFD, EFE 2A, had
14.3 times more xylanase III, 14.3 times more β-xylosidase, 7.7 times more endoglucanase III,
and 1.9 times more polysaccharide monooxygenase, though the differences were less than those
between 2A and 9C (Table 3-9). Nevertheless, 11C had 3.1 times more β-glucosidase I and 8.6
times more xyloglucanase with CBM1 than 2A. These enzymes are responsible for cellobiose
and xyloglucan hydrolysis, respectively. Hence, they partially compensated for lower amounts of
certain important fibrolytic activities in 2A, as evidenced by the increase in NDFD due to adding
11C.
82
Conclusions
Several promising EFE candidates that reduced the fiber concentration of bermudagrass
and increased its digestibility were identified in this study. Increases of up to 4.8% units in
NDFD due to EFE treatment were detected, which could result in production of an extra 1.2 kg/d
of 4% FCM. Application of EFE also increased VFA concentrations, particularly propionate,
which could increase the supply of glucose for lactose synthesis and hence increase milk
production. The improved digestibility seemed to be at least partly explained by hydrolysis of
fiber fractions leading to release of WSC and phenolic compounds from the cell wall. These
results confirm that certain EFE can be used to improve the nutritive value of tropical/subtropical
forages. Regression analyses revealed that enzyme activities accurately predicted pre-ingestive
hydrolysis measures (WSC, FER) and moderately predicted NDF hydrolysis, but poorly
predicted digestibility measures. This indicates that enzyme activity estimates should not be used
to choose the best EFE for improving forage digestibility or animal performance. The proteomics
iTRAQ LC-MS analysis revealed that relative to the most effective EFE, the least effective EFE
at increasing NDFD contained lesser amounts of specific enzymes and auxiliary proteins
necessary for xylan and lignocellulose degradation. This technique could be useful for selecting
the most promising EFE for animal studies, particularly as enzyme activity is poorly related to
digestibility measures, which are closely related to animal performance.
83
Table 3-1. Form, dose (g/kg of bermudagrass DM), biological source, activities of endoglucanase, xylanase, exoglucanase, β-
glucosidase (μmol of sugar released/min/g), ferulic acid esterase activity (nmol of ferulic acid released/min/g), and protein
concentration (mg/g) of exogenous fibrolytic enzyme (EFE) preparations used in in vitro digestion assays.
EFE Form Dose Biological Source Endoglucanase Xylanase Exoglucanase β-glucosidase Ferulic acid
esterase Protein
1A Liquid 2.33 Trichoderma reesei 1,693 1,276 1.68 10.1 2.18 65.3
2A Liquid 2.33 T. reesei 3,624 29,301 0.84 11.7 1.46 111.1
3A Liquid 2.33 T. reesei 2,659 10,234 2.53 15.2 7.35 72.4
4A Liquid 2.33 Myceliopthora
thermophila 416 291 0.77 0.4 1.7 19.1
5A Liquid 2.33 M. thermophila 391 4,269 0.8 0.1 16.4 24.8
9C Liquid 0.1 Aspergillus sp. and T.
reesei 663 2,596 0.88 6.2 3.34 43.0
11C Liquid 10.4 Trichoderma sp. 1,506 1,703 0.97 12.7 6.30 81.1
12C Solid 0.03 Bacillus subtilis 9 1,853 0.44 0.4 0 29.9
13D Liquid 15.6 Aspergillus oryzae 286 86 0.29 1.9 2.35 18.0
14D Liquid 15.6 Aspergillus aculeatus 512 314 3.26 0 2.27 43.8
15D Liquid 15.6 A. oryzae 70 6,499 0.29 0.1 2.57 28.3
17D Liquid 15.6 T. reesei 844 589 1.72 0.9 2.39 113.1
SD 112 212 0.10 0.9 1.27 25.0
84
Table 3-2. Effects of exogenous fibrolytic enzyme addition on in vitro true dry matter (DMD), neutral detergent fiber (NDFD),
hemicellulose (HEMD), acid detergent fiber (ADFD), and cellulose (CELD) digestibility, and lignin disappearance
(ADLD) of a 4-wk regrowth of Tifton 85 bermudagrass haylage (Experiment 1).a
Treatment DMD (%) NDFD (%) HEMD (%) ADFD (%) CELD (%) ADLD (%)
Control 52.0 35.6 33.0 38.7 41.8 7.5
1A 54.4** 39.8** 37.0** 42.5** 45.1** 21.1**
2A 54.8** 40.4** 37.7** 42.7** 45.8** 17.7+
3A 52.7 38.0* 35.7** 40.1 44.4* 4.4
4A 53.8** 38.5** 37.2** 40.3 43.6 10.8
5A 52.4 37.1 35.3* 39.5 42.6 14.5
9C 52.5 36.2 33.4 38.6 42.3 11.2
11C 54.9** 40.0** 38.0** 42.0** 45.5** 13.6
12C 53.9** 38.6** 36.7** 40.8+ 44.6** 8.5
13D 54.1** 38.9** 37.4** 41.3* 45.1** 12.6
14D 52.5 36.3 34.6 38.7 42.5 7.7
15D 53.5+ 38.5** 37.0** 39.6 43.5 10.6
17D 53.2 37.8* 34.8 41.0* 45.4** 8.3
SEM 0.43 0.55 0.60 0.63 0.65 3.21
aMeans within a column differed [+ (P < 0.10), * (P < 0.05) and ** (P < 0.01)] from the Control mean.
85
Table 3-3. Descriptive statistics for dependent and independent variables used to develop
regression relationships between activities of exogenous fibrolytic enzymes (EFE)
and measures of pre-ingestive fiber hydrolysis or in vitro digestibility of a 4-wk
regrowth of Tifton 85 bermudagrass haylage.
Mean Minimum Maximum
EFE Activities (µmol/min applied to 0.5 g bermudagrass DM)
Endoglucanase 2.35 0.00 7.53
Exoglucanase 0.004 0.00 0.024
β-glucosidase 0.01 0.00 0.06
Xylanase 9.00 0.00 48.74
Ferulic acid esterase 1×10-5 0.00 3×10-5
Digestibility measuresa (%)
DMD 53.5 47.6 58.2
NDFD 38.3 29.2 46.7
HEMD 36.3 28.5 45.0
ADFD 40.7 32.5 49.3
CELD 44.3 35.4 52.7
Pre-ingestive hydrolysis measuresb
NDF (%) 65.8 62.8 67.3
WSC (%) 3.4 2.3 5.9
FER (µg/g) 248.2 198 391 aDigestibility of DM (DMD), NDF (NDFD), hemicellulose (HEMD), ADF (ADFD), and
cellulose (CELD),
bWater-soluble carbohydrates (WSC) and ferulic acid (FER).
86
Table 3-4. The accuracy of predicting the in vitro digestibility of DM (DMD), NDF (NDFD), hemicellulose (HEMD), ADF (ADFD),
and cellulose (CELD) of bermudagrass haylage from various enzyme activity estimates using stepwise multiple regression
(Experiment 1).a DMD (%) NDFD (%) HEMD (%) ADFD (%) CELD (%)
Activitya Partial R2 P > F Partial R2 P > F Partial R2 P > F Partial R2 P > F Partial R2 P > F
Endoglucanase
(EN)
n.s.b n.s. n.s. n.s. 0.03 0.03
Endoglucanase2 n.s. n.s. n.s. n.s. n.s
Exoglucanase
(EX)
n.s. n.s. n.s. n.s. 0.02 0.08
Exoglucanase2 n.s. n.s. n.s. n.s. n.s
Β-glucosidase
(BG)
0.05 0.005 n.s. n.s. 0.04 0.02 n.s
Β-glucosidase2 n.s. n.s. n.s. 0.02 0.06 n.s
Xylanase (XY) n.s. n.s. n.s. n.s. n.s
Xylanase2 n.s. n.s. n.s. n.s. n.s
FA Esterase
(FE)
n.s 0.05 0.01 0.03 0.05 n.s. n.s
FA Esterase2 n.s. n.s. n.s. n.s. n.s
Model parameters
Model DMD = 53.17 +
30.462(BG)
NDFD = 37.80 +
49.579(FE)
HEMD = 35.90 +
42.438(FE)
ADFD = 39.83 +
155.897 (BG) – 1863.266
(BG)2
CELD = 43.66 + 0.416
(EN) – 97.921 (EX)
R2 c 0.05 0.05 0.03 0.06 0.06
RMSEd 2.18 3.84 4.16 3.53 3.63
P > F 0.005 0.01 0.05 0.01 0.02 aOnly activities that were correlated (P ≤ 0.10) with DMD, NDF, HEMD, ADFD or CELD were included in the respective multiple
regression models; bn.s.= not significant (P > 0.10); cPartial R2 = Variable contribution to explanation of variation in response
variable; dRMSE= root-mean square error.
87
Table 3-5. Effects of exogenous fibrolytic enzymes on concentrations of total volatile fatty acids (TVFA), acetate, propionate,
butyrate, acetate to propionate ratio (A:P), isobutyrate, isovalerate, valerate, ammonia N (NH3N) and pH in the filtrate after
fermentation of a 4-wk regrowth of Tifton 85 bermudagrass haylage in buffered-rumen fluid (Experiment 1).a
Treatment TVFA
(mM)
Acetate
(mM)
Propionate
(mM)
Butyrate
(mM) A:P
Isobutyrate
(mM)
Isovalerate
(mM)
Valerate
(mM)
NH3N
(mg/dL) pH
Control 55.4 33.5 10.4 4.70 3.24 2.17 2.21 2.66 38.6 7.28
1A 61.2** 36.9** 11.7** 5.05* 3.15* 2.60* 2.29 2.50 38.5 7.21**
2A 58.8+ 36.2** 11.4** 4.87 3.16* 2.16 2.27 2.49 39.8 7.31
3A 56.5 34.3 10.6 4.66 3.23 1.91 2.17 2.79 39.4 7.29
4A 61.0** 36.5** 12.2** 5.05* 3.03** 2.62* 2.36* 2.40 39.9+ 7.31
5A 60.0** 36.7** 11.4** 4.84 3.19 2.35 2.22 2.34+ 40.2* 7.34**
9C 59.1* 35.4+ 10.9 4.80 3.18 2.00 2.21 2.43 38.5 7.26
11C 58.2 35.2 11.1+ 4.85 3.15* 2.07 2.21 2.74 38.1 7.31
12C 58.7+ 35.4+ 11.1 4.87 3.20 2.29 2.29 2.77 38.3 7.33*
13D 59.9** 36.4** 11.2* 5.04* 3.27 2.37 2.33+ 2.53 39.7 7.30
14D 58.4 35.2 11.0 4.81 3.22 2.03 2.22 2.73 38.6 7.28
15D 56.2 34.2 10.6 4.82 3.20 2.07 2.20 2.73 38.4 7.32
17D 60.1** 35.1 11.6** 5.23** 3.09** 2.20 2.25 3.21** 39.4 7.20**
SEM 0.93 0.57 0.20 0.09 0.026 0.119 0.041 0.107 0.41 0.015
aMeans within a column differed [+ (P < 0.10), * (P < 0.05) and ** (P < 0.01)] from the Control mean.
88
Table 3-6. Effects of exogenous fibrolytic enzymes on DM loss and concentrations of NDF, hemicellulose (HEM), ADF, cellulose
(CEL), lignin (ADL), and release of water-soluble carbohydrates (WSC), cellobiose, glucose, xylose, arabinose, ferulic
acid (FER), and p-coumaric acid (COU) after preingestive hydrolysis of a 4-wk regrowth of Tifton 85 bermudagrass
haylage (Experiment 2).a
Treatment
DM
loss
(%)
NDF
(%)
HEM
(%)
ADF
(%)
CEL
(%)
ADL
(%)
WSC
(%)
Cellobiose
(mg/g)
Glucose
(mg/g)
Xylose
(mg/g)
Arabinose
(mg/g)
FER
(μg/g)
COU
(μg/g)
Control 22.0 67.3 33.8 33.5 30.1 3.4 2.28 0.00 4.30 0.01 0.12 198 162
1A 21.7 66.0 33.0 33.1 29.6 3.5 3.39** 0.59** 8.86** 0.29** 0.28 225** 166
2A 24.4** 62.8** 29.3** 33.4 29.8 3.6 5.18** 1.31** 11.39** 2.15** 1.08** 391** 203**
3A 23.2 65.3 32.1** 33.2 29.6 3.6 3.56** 1.05** 10.50** 0.18** 0.52** 241** 171**
4A 22.3 66.1 32.5* 33.6 30.1 3.4 2.94** 0.00 8.22** 0.07+ 0.22 210** 165
5A 22.3 65.9 32.9 32.9 29.5 3.4 2.72** 0.24** 7.70** 0.29** 0.33** 221** 170**
9C 21.5 66.8 33.4 33.4 29.9 3.5 2.68** 0.46** 8.80** 0.11** 0.32* 207+ 164
11C 23.6 64.4* 31.3** 33.1 29.5 3.7 4.54** 2.25** 14.33** 0.36** 1.06** 285** 175**
12C 21.9 66.7 32.8 33.9 30.3 3.5 2.33 n.a. n.a. n.a. n.a. 203 161
13D 22.3 66.3 33.1 33.2 29.6 3.6 2.27 n.a. n.a. n.a. n.a. 200 163
14D 22.3 66.9 33.2 33.7 30.1 3.6 2.89** 0.06 5.33** 0.59** 0.13 207+ 166
15D 21.9 66.6 33.3 33.4 30.0 3.4 3.13** 0.08 4.60 1.13** 0.32* 340** 190**
17D 24.3** 63.7** 31.8** 31.9 28.6 3.3 5.85** 0.04 23.00** 0.37** 0.30* 298** 178**
SEM 0.47 0.6 0.45 0.43 0.39 0.10 0.04 0.04 0.20 0.02 0.05 2.3 1.3
aMeans within a column differed [+ (P < 0.10), * (P < 0.05) and ** (P < 0.01)] from the Control mean; n.a.= not analyzed.
89
Table 3-7.The accuracy of predicting concentrations of neutral detergent fiber (NDF), water-soluble carbohydrates (WSC), and ferulic
acid (FER) of untreated and enzyme-treated Tifton 85 bermudagrass haylage (Experiment 2).a WSC (%) FER (μg/g) NDF (%)
Activitya Partial R2 P > F Partial R2 P > F Partial R2 P > F
Endoglucanase
(EN)
0.65 < 0.01 0.17 < 0.01 0.33 < 0.01
Endoglucanase2 n.s.b 0.02 0.09 n.s.
Exoglucanase
(EX)
0.01 < 0.01 0.02 < 0.01 0.10 < 0.01
Exoglucanase2 n.s. 0.02 n.s. 0.03 0.08
Β-glucosidase
(BG)
0.06 < 0.01 0.07 < 0.01 0.15 < 0.01
Β-glucosidase2 0.06 < 0.01 0.01 < 0.01 n.s.
Xylanase (XY) 0.07 < 0.01 0.71 < 0.01 n.s.
Xylanase2 n.s. 0.001 0.03 n.s.
FA Esterase
(FE)
n.s. n.s. n.s.
FA Esterase2 n.s. n.s. n.s.
Model parameters
Model WSC= 2.466 + 0.873(EN) – 121.627(EX) –
92.188(BG) + 454.768(BG)2 + 0.012(XY)
FER= 203.870 + 51.346(EN) – 1.581(EN)2 -
15287(EX) + 333229(EX)2 – 5447.990(BG)
+ 44889(BG)2 + 1.500(XY) + 0.03 (XY)2
NDF= 66.5 – 1.268(EN) + 480.731(EX) -
10811(EX)2 + 84.663(BG)
R2 c 0.95 0.99 0.62
RMSEd 0.27 6.64 1.07
P > F < 0.001 < 0.001 < 0.001 aOnly activities that were correlated (P ≤ 0.10) with WSC, FER, or NDF were included in the respective multiple regression models; bn.s.= not significant (P > 0.10); cPartial R2 = Variable contribution to explanation of variation in response variable; dRMSE= root-
mean square error.
90
Table 3-8. Relative ratio of proteins in EFE 9C to those in 2A as detected by iTRAQ LC-MS/MS analysisa. The EFE were sourced
from both Trichoderma reesei and Aspergillus sp. and from T. reesei, respectively.
Protein name / Enzymatic activity Class Family Species Ratio S.D.
Proteins with fold changes lower than 0.8 or greater than
1.2b
Xylanase III (E.C. 3.2.1.8) GH 10 T. reesei 0.03 0.002
β-xylosidase (E.C. 3.2.1.37) GH 3 T. reesei QM6a 0.04 0.005
Protein for fungi growth T. reesei QM6a 0.04 0.006
Acetylxylan esterase + CBM1 (E.C. 3.1.1.72) CE 5 T. reesei 0.06 0.004
Endoglucanase III (E.C. 3.2.1.4) GH 12 T. reesei QM6a 0.10 0.013
Polysaccharide monooxygenase + CBM1 (E.C.
3.2.1.4) AA 9 T. reesei QM6a 0.13 0.013
Swollenin (Swo1) T. reesei 0.33 0.026
β-galactosidase (EC 3.2.1.23) GH 35 Aspergillus
fumigatus 19.61 2.927
Rhamnogalacturonan acetylesterase (E.C. 3.1.1.86 ) CE 12 T. reesei 28.23 2.701
Proteins with fold changes between 0.8 and 1.2
Endo-1,4-β –xylanase (E.C. 3.2.1.8) GH 11 T. reesei 0.87 0.189
β-glucosidase I (E.C. 3.2.1.21) GH 3 T. reesei 1.02 0.096
Amidase (E.C. 3.5.1.4) T. reesei QM6a 1.20 0.251
Endoglucanase II + CBM1(E.C. 3.2.1.4) GH 5 T. reesei QM6a 1.20 0.561 aRatios of 9C:2A are mean values from three independent replicates of each EFE, each quantified with at least three unique peptides;
iTRAQ LC-MS/MS= Isobaric tag for relative and absolute quantitation liquid chromatography-mass spectrometry; AA= auxiliary
activity; CE= carbohydrate esterase; GH= glycoside hydrolase bProtein ratios with a P value < 0.05 calculated by Student’s t-test.
91
Table 3-9. Relative ratio of proteins in EFE 11C to those in 2A as detected by iTRAQ LC-MS/MS analysisa. Both EFE were sourced
from Trichoderma reesei.
Protein name / Enzymatic activity Class Family Species Average
Ratio Std. Dev
Proteins with fold changes lower than 0.8 or greater than
1.2b
Xylanase III (E.C. 3.2.1.8) GH 10 Trichoderma
reesei 0.07 0.012
β-xylosidase (E.C. 3.2.1.37) GH 3 Trichoderma
reesei QM6a 0.07 0.013
Endoglucanase III (E.C. 3.2.1.4) GH 12 Trichoderma
reesei QM6a 0.13 0.018
Polysaccharide monooxygenase + CBM1 (E.C.
3.2.1.4) AA 9
Trichoderma
reesei QM6a 0.52 0.074
β-glucosidase I (E.C. 3.2.1.21) GH 3 Trichoderma
reesei 3.10 0.326
Xyloglucanase + CBM1 (E.C. 3.2.1.151) GH 74 Trichoderma
reesei 8.54 2.271
Proteins with fold changes between 0.8 and 1.2
Ribonuclease T3 Trichoderma
reesei QM6a 0.85 0.385
Typ 1 glutamine amidotransferase Trichoderma
reesei QM6a 0.98 0.300
Glucan endo-1,3-β-D-glucosidase (E.C. 3.2.1.39) GH 17 Trichoderma
reesei QM6a 1.07 0.237
aRatios of 11C:2A are mean values from three independent replicates of each EFE, each quantified with at least three unique peptides;
iTRAQ LC-MS/MS= Isobaric tag for relative and absolute quantitation liquid chromatography-mass spectrometry; AA= auxiliary
activity; GH= glycoside hydrolase bProtein ratios with a P value < 0.05 calculated by Student’s t-test.
92
CHAPTER 4
EFFECT OF THE DOSE OF EXOGENOUS FIBROLYTIC ENZYME PREPARATIONS ON
PREINGESTIVE FIBER HYDROLYSIS AND IN VITRO DIGESTIBILITY OF
BERMUDAGRASS HAYLAGE
Background
Warm-season grasses are extensively used for cattle production in the Southeast.
Bermudagrass is the most important of such grasses that is used for cattle production (Newman,
2007) but like other warm-season grasses, the quality of bermudagrass is low (Hanna and
Sollenberger, 2007). Exogenous fibrolytic enzyme (EFE) treatment has been proposed as a
method to improve forage quality and animal performance but results of published studies have
been equivocal (Adesogan, 2005). Various enzyme, animal, feed, and management factors
influence the efficacy of fibrolytic EFE (Beauchemin and Colombatto, 2003; Adesogan et al.,
2013), many of which are challenging to control. One of such factors that is easily controlled is
the dose of the EFE. To our knowledge, only two studies (Dean et al., 2005; Krueger et al., 2008)
have been conducted on effects of the dose of EFE on the nutritive value of bermudagrass. Dean
et al. (2005) reported that 48-h in vitro NDF digestibility (NDFD) increased quadratically with
increasing dose of one of three cellulase-xylanase EFE applied at the point of ensiling to a 5-wk
regrowth of Tifton 85 bermudagrass. Krueger et al. (2008) reported that applying increasing
doses of an EFE with high esterase activity to Coastal or Tifton 85 bermudagrass hay had no
effect on 6, 24, and 48 h in vitro NDFD except for a linear increase in 6 h NDFD of the Tifton 85
cultivar. More studies are needed to examine effects of EFE dose rates on the quality of
bermudagrass hay, silage and haylage due to the important role of these forages in the diets of
dairy and beef cattle in the Southeast. This is because exogenous fibrolytic enzymes can be
ineffective if applied in insufficient or excessive amounts (Sanchez at al., 1996; Beauchemin et
al., 2004). Low doses do not fully exploit the hydrolytic potential of EFE, especially during short
93
incubation times. In contrast, excessively high doses decrease availability of substrates for
catalysis or accessibility of substrates to these sites by crowding the substrate surface, which
reduces the enzymatic hydrolysis rate (Bommarious et al., 2008). In the rumen, competition
between excessively high doses of EFE and ruminal endogenous cellulolytic bacterial enzymes
for substrates can decrease fiber digestibility (Nsereko et al., 2002) and consequently reduce
animal performance (Kung et al., 2000). Therefore, optimization of the EFE dose is critical for
using EFE to improve the digestibility of forages. The objective of this study was to determine
the optimum dose of 5 EFE that were selected as the most promising of 18 candidates from 5
companies at improving the NDFD of bermudagrass haylage (BH, Chapter 3). The hypothesis
was that increasing the dose of each EFE would increase the NDFD of bermudagrass haylage in
a quadratic manner.
Materials and Methods
Bermudagrass Substrate
An established stand of bermudagrass (Cynodon dactylon cv. Tifton 85) in Alachua,
Alachua County, Florida was staged in June, 2010 by mowing to a 4-cm stubble and removing
the residue. The field was fertilized subsequently with N (95 kg/ha) and the grass was allowed to
regrow for 4-wk such that the harvest day was July 7th, 2010. On harvest day, the grass was
mowed within 1 d to a 4-cm stubble with a CLAAS 3500 mower conditioner (CLAAS North
America, Omaha, NE). The grass was wilted for 2.5 h in the windrow and then rolled into round
280-kg round bales without inoculant addition. Bales were wrapped with 7 layers of 6-mm
plastic and ensiled for 53 d. Ensiled bermudagrass was chosen over hay since it is more typically
used in this form by dairy producers due to the high humidity and frequent summer rainfall in
Florida (Staples, 2003). Representative haylage samples were collected as substrate for this
study, dried at 60oC for 48 h and ground to pass the 1-mm screen of a Wiley mill (Arthur H.
94
Thomas, Philadelphia, PA). The haylage had 49.4% of DM and 93.5, 68.1, 34.2, 3.7, and 18.7%
of OM, NDF, ADF, ADL, and CP, respectively (DM basis).
Enzymes
Five previously selected (Chapter 3) commercial and experimental EFE preparations
provided by 3 manufacturers were examined at 4 doses (0, 0.5, 1, and 2×, where 1× is the
manufacturer-recommended doses in this study. Table 4-1 lists the enzymatic activities and
protein concentrations, form, doses, and biological sources of the EFE preparations.
Endoglucanase (enzyme commission, E.C. 3.2.1.4), exoglucanase (E.C. 3.2.1.91), xylanase (E.C.
3.2.1.8) and β-glucosidase (E.C. 3.2.1.21) activities were quantified using carboxymethyl
cellulose, avicel, oat-spelt xylan, and cellobiose as artificial substrates (Colombatto and
Beauchemin, 2004). Ferulic acid esterase (E.C. 3.1.1.73) activity was measured using ethyl
ferulate as the substrate (Lai et al., 2009). All activities were measured at 39oC and a pH of 6 to
mimic conditions in the rumen of a typical US lactating dairy cow. Protein concentration was
measured using the Bio-Rad protein assay (Bradford, 1976) with bovine serum albumin as the
standard (Bio-Rad Laboratories, Hercules, CA).
In Vitro Ruminal Digestibility (Experiment 1)
All EFE were evaluated with a 24-h in vitro ruminal digestibility assay (Goering and Van
Soest, 1970) using bermudagrass haylage as the substrate. As described by Krueger et al. (2008),
EFE were diluted in 0.1 M citrate–phosphate buffer (pH 6) and 2 mL of the solution containing
the requisite EFE dose was applied to 0.5 g of substrate. The Control treatment consisted only of
the citrate–phosphate buffer and the substrate. Treatments were applied in quadruplicate to the
substrate in 100-mL polypropylene tubes capped with a Bunsen rubber stopper fitted with a one-
way gas-release valve. Two blank tubes per treatment, containing no substrate, were included as
EFE blanks. Tubes were tapped gently to ensure proper mixing of EFE solution with the
95
substrate and the suspensions were subsequently incubated at 25oC for 24 h before addition of
buffered-ruminal fluid. The ruminal fluid was representatively collected by aspiration 3 h after
feeding (0800 h) from two non-lactating non-pregnant ruminally-cannulated Holstein cows
consuming a ration consisting of coastal bermudagrass hay ad libitum supplemented with corn
(0.45 kg), cottonseed hulls (0.46 kg), soybean meal (0.90 kg), and a vitamin-mineral mix (35.8 g)
(DM basis). The ruminal fluid collection protocol was approved by the University of Florida,
Institute of Food and Agricultural Sciences, Animal Research Committee. The ruminal fluid
collected was filtered through four layers of cheesecloth and mixed with pre-warmed artificial
saliva (Goering and Van Soest, 1970). Buffered-ruminal fluid (52 mL) was dispensed into pre-
warmed tubes. Tubes were incubated at 39oC for 24 h. Fermentation was terminated by placing
the tubes on ice. Tube contents were filtered through previously dried (60oC for 48h) and
weighed 125-mm Whatman No. 451 paper (Fisher Scientific, Pittsburgh). Filtrate and residues
were collected for further analysis. Residues were dried at 60oC for 48 h, weighed, and analyzed
for NDF, ADF and ADL sequentially (Van Soest, 1991) using an ANKOM 200 Fiber Analyzer
(ANKOM, Macedon, NY). Hemicellulose (HEM) was calculated as the difference between NDF
and ADF and cellulose (CEL) as the difference between ADF, and ADL. Residue weights and
their fiber concentrations were used to calculate true DM, NDF, hemicellulose, ADF and
cellulose digestibility (DMD, HEMD, ADFD and CELD). Filtrate samples were analyzed for pH
using an Accumet XL25 pH meter (Fisher Scientific, Pittsburgh, PA), acidified with 50% H2SO4
(1% v/v) and centrifuged at 8,000 × g for 20 min. The supernatant was frozen (-20oC) and
subsequently analyzed for concentrations of VFA (Muck and Dickerson, 1988) using a Merck
Hitachi Elite LaChrome High Performance Liquid Chromatograph system (Hitachi, L2400,
Tokyo, Japan) fitted with a Bio-Rad Aminex HPX-87H column (Bio-Rad Laboratories,
96
Hercules, CA). Ammonia-N was determined with a Technicon Auto Analyzer (Technicon,
Tarrytoen, NY) and an adaptation of the Noel and Hambleton (1976) procedure that involved
colorimetric N quantification.
Preingestive Fiber Hydrolysis (Experiment 2)
The same EFE doses examined in Experiment 1 were tested in this experiment to
ascertain their effects on preingestive hydrolysis of bermudagrass haylage. Samples were
assayed as described in Experiment 1 except that buffered-ruminal fluid was omitted from the
assay, 50-mL centrifuge tubes were used and sodium azide was added to the 2-mL buffered EFE
solution applied to the substrate (0.02% w/v) to prevent microbial degradation of substrate
(Krueger et al., 2008). Two blank tubes per treatment, containing no substrate, were included as
EFE blanks. After the incubation at 25oC, 15 mL of double distilled water were added and tubes
were shaken for 1 h at 260 oscillations/min with an Eberbach Reciprocating Shaker Model 6000
(Eberbach corporation, Ann Arbor, MI). Tubes were filtered through previously dried (60oC for
48h), and weighed 125-mm Whatman No. 451 filter paper (Fisher Scientific, Pittsburgh) and
filtrate and residue samples were collected. Residues were dried at 60oC for 48 h, weighed and
analyzed for NDF, ADF and ADL as described previously. Residue and sample dry weights and
DM concentrations were used to calculate DM losses. Residue weights and their fiber
concentrations were used to calculate fiber fraction concentrations. Filtrate samples were frozen
(-20oC) and subsequently analyzed for water-soluble carbohydrates (WSC; DuBois et al., 1956)
and ferulic (FER) and p-coumaric acids (COU; Bio-Rad, 2011) using the High Performance
Liquid Chromatograph system and a HPX-87H column described previously.
97
Statistical Analyses
A randomized complete block design with four replicates per treatment and two runs was
used to determine the effects of EFE preparations on digestibility and fermentation measures
(Experiment 1) and substrate DM and fiber disappearance and release of sugars and phenolic
acids (Experiment 2). Run was the blocking factor.
The model used to analyze digestibility, fermentation and preingestive hydrolysis data
was:
Yijk = µ + Di + Rj + DRij + Eijk
Where:
µ = general mean
Di = effect of dose i
Rj = effect of run j
DRij = effect of dose i × run j interaction
Eijk = experimental error
The GLM procedure of SAS v.9.1 (2012) was used to analyze each EFE separately,
because comparing dose rate effects among EFE were not of interest. Polynomial contrasts were
used to determine dose effects and the Fisher’s F-protected least significance difference test was
used to determine the optimal dose. Both of these mean characterization and separation tests
were considered necessary to properly interpret the results because they depict the polynomial
trend and the optimal dose, respectively. The final decision on the optimal dose of the EFE for
future in vitro and animal trials was defined as the least dose that resulted in a greater increase in
NDFD than lower doses and a similar or greater response relative to higher doses. Neutral
detergent fiber digestibility was chosen as the response of choice for selecting the optimal dose
98
because of its correlation with DMI and milk production (Oba and Allen, 1999). Significance
was declared at P < 0.05 and tendencies at P > 0.05 < 0.10.
Results and Discussion
Experiment 1: EFE Dose Effects on Measures of in Vitro Digestion and Fermentation
EFE dose effects on digestibility measures
Since both HEM (NDF-ADF) and CEL (ADF-ADL) concentrations are calculated by
difference, changes in ADFD or ADLD will influence the HEMD and CELD values,
respectively. This needs to be borne in mind when interpreting HEMD and CELD results.
Increasing the dose of EFE 1A linearly increased (P < 0.01) HEMD and had a cubic
effect (P < 0.05) on DMD and NDFD (Table 4-2). Compared to the Control, the 2× dose resulted
in the greatest increase in NDFD and HEMD (7.4 and 11.5%, respectively). This dose also
resulted in the greatest increase in NDFD per unit of EFE 1A used (0.56% NDFD per g of EFE).
Applying EFE 1A did not affect DMD, but the 2 and 3× doses resulted in greater DMD than the
0.5× dose. Applying increasing doses of 1A resulted in cubic effects (P < 0.05) on ADFD and
CELD partly because the respective values decreased to a nadir when the 0.5× and 1× doses
were applied (-8.4 and -10.9%, and -9.8 and -9.5%, respectively; P < 0.05). Dean et al. (2005)
also reported that applying an EFE preparation from T. viride at the point of ensiling Tifton 85
bermudagrass decreased 48-h in vitro ADFD (- 6%). The decrease in ADFD and CELD with
increasing dose of EFE 1A is probably attributable to the declining reactivity of residual
cellulose during enzymatic hydrolysis due to the decrease in surface area and number of
accessible chain ends and/or adsorption of inactive cellulase on the surface of cellulose (Zhang
and Lynd, 2004). Yet, if the EFE dose is too low, the supply of auxiliary enzymes and proteins
like swollenin may be insufficient to remove barriers preventing increases in ADFD and CELD.
99
Applying increasing doses of EFE 2A increased (P < 0.05, quadratic) DMD, NDFD,
HEMD (Table 4-2). The 2× dose resulted in the greatest (P < 0.05) increases in NDFD and
HEMD that differed from lower doses (10.8 and 16.2%, respectively) but the dose that gave the
greatest increase in NDFD per unit of EFE 2A was 0.5× (1.97% NDFD units per g of EFE). The
choice of which dose to use depends on the desired objective. If the intent is to maximize NDFD,
the 2× dose should be selected, whereas the 0.5× dose should be chosen from an efficiency
standpoint. Increasing the dose of EFE 2A did not increase ADFD and CELD. Therefore, the
EFE exerted its hydrolytic effect on HEM rather than CEL, likely due to its high xylanase
activity. Hemicellulose typically represents about half of the fiber concentration in grasses (Van
Soest, 1994), and it was the fiber fraction most effectively hydrolyzed by adding EFE to BH
(Romero et al., 2013).
Increasing the dose of EFE 11C increased ADFD and CELD (P < 0.01, linear), DMD
(P< 0.01, quadratic), and NDFD and HEMD (P < 0.01, cubic) (Table 4-2). The 1× dose resulted
in the greatest increases in NDFD, HEMD and ADFD (16.2, 22.6 and 6.7%, respectively, P <
0.05) whereas that for CELD resulted from the 3× dose (6.2%). The dose resulting in the greatest
increase in NDFD per unit of EFE 11C was 0.5× (0.37% NDFD per g of EFE). As was the case
for EFE 1A and 2A, EFE 11C had its greatest effects on the HEM fiber fraction. However,
unlike 1A and 2A, 11C also increased ADFD and CELD. This is likely because EFE 11C
supplied more exoglucanases than the other EFE. Exoglucanases progressively cleave cellulose
chains at the reducing and nonreducing ends to release cellobiose or glucose after the hydrolytic
cleavage of internal parts of the chain by endoglucanases (Zhang et al., 2006). This enzymatic
depolymerization is the rate-limiting step in cellulose hydrolysis (Zhang et al., 2006).
100
Applying EFE 13D at increasing doses had a cubic effect (P < 0.01) on all digestibility
measures (Table 4-2) and the 0.5× was the optimal dose for increasing DMD, NDFD, HEMD,
and ADFD. This dose resulted in the greatest increases in these digestibility measures (5.14,
13.1, 19.4 and 5.4%, respectively) and elicited the greatest increase in NDFD per unit of EFE
13D (0.59% per g of EFE). Applying 13D did not increase CELD, rather the 2× dose decreased
the response. Hemicellulose was the fiber fraction hydrolyzed to the greatest extent by 13D
despite the relatively low xylanase activity of the EFE.
Increasing the dose of EFE 15D resulted in no effect (P > 0.05) on DMD, quadratic
increases in NDFD (P < 0.10, tendency) and HEMD (P < 0.01) and cubic effects (P < 0.05) on
ADFD and CELD (Table 4-2). The optimal dose for increasing NDFD and HEMD was 0.5×
because the respective responses peaked and plateaued at this dose (5.1 and 9.9%), respectively,
and it resulted in the greatest increase in NDFD per unit of EFE 15D (0.23% per g of EFE).
However, as reported by Dean et al. (2005) for a T. reesei EFE, applying all doses of 15D
decreased ADFD and the 0.5× and 1× doses decreased CELD. This EFE had the poorest effect
on in vitro digestibility perhaps because it had a lower ratio of collective cellulase activities
(endoglucanase, exoglucanase and β-glucosidase) to xylanase activity than other EFE (0.01 vs.
1.43). This might have reduced depolymerization of bermudagrass fiber because synergy
between cellulases and xylanases is needed for lignocellulose degradation (Lynd et al., 2002).
This EFE probably only degraded the most accessible and digestible amorphous cellulose
fractions because its cellulase activity was relatively low. This would have left the more
recalcitrant fractions undegraded, leading to the decreases in ADFD and CELD.
EFE dose effects on fermentation measures
Increasing the dose of EFE 1A had no effect on total VFA (TVFA), acetate, isobutyrate,
valerate, and NH3N concentrations but linearly reduced (P < 0.01) the acetate to propionate
101
(A:P) ratio due to linearly increasing (P < 0.10, tendency) propionate concentration. In addition,
increasing the 1A dose caused cubic responses (P < 0.01) in butyrate and isovalerate
concentrations and pH (Table 4-3). The increased propionate availability at high doses of 1A
would provide more gluconeogenic precursors for the liver, thus improving glucose supply for
lactose synthesis in the mammary gland, such that milk yield may increase (VandeHaar, 2005).
Butyrate concentration was decreased by increasing the dose of 1A (P < 0.05), with the nadir
occurring at doses of 0.5 and 1× (- 13.2 and - 14.8%), respectively. This pattern reflects the
decreases in ADFD and CELD when the same doses of 1A were used because butyrate is one of
the main VFA generated during fiber fermentation (Rinne et al., 1997). Isovalerate concentration
increased with the 0.5 and 3× doses of EFE 1A. This probably indicates that the EFE increased
ruminal protein degradation because isovalerate is a product of leucine deamination (Van Soest
et al., 1994) and pepsin-like proteases were detected in the secretome of EFE from T. reesei
(Adesogan et al., 2013).
Increasing the dose of EFE 2A had no effect on NH3N concentration, a linear increase on
pH (P < 0.01), and a cubic effect on TVFA, acetate, propionate, butyrate, isobutyrate, isovalerate
and valerate concentration and A:P ratio (P < 0.05; Table 4-3). Total VFA concentration was
increased (P < 0.05) by all doses of 2A except 2×. This suggests this EFE has great potential to
increase energy supply in dairy cattle because TVFA provides 70% of the caloric requirements in
ruminants (Bergman, 1990). This increase in TVFA was due mostly to corresponding increases
in concentrations of propionate (P < 0.05; 11.0, 15.6, and 20.2% at 0.5×, 1×, and 3×,
respectively), butyrate (P < 0.05; 12.5 and 9.4% at 0.5×, and 1×, respectively) and acetate (P <
0.05; 7.5% at 3×). The A:P ratio was decreased by increasing doses of 2A because the latter
increased propionate concentrations but did not affect acetate concentration except at the 3×
102
dose. This increased availability of propionate would be a valuable glucose precursor for milk
production in cattle particularly in early lactation when the energy demand for milk production is
critical. Concentrations of isobutyrate and isovalerate were increased by applying 2A at doses of
0.5×, 1×, and 3× (20.1, 21.7, 39.2%) and 0.5×, 1×, 2×, and 3× (7.7, 10.1, 11.0, 18.3%),
respectively, but the 2× dose decreased valerate concentration (- 13.7%). The increase in
concentration of branched-chained VFA suggests that like 1A, EFE 2A also stimulated protein
degradation. This did not result in greater NH3N concentrations probably because the high NH3N
concentration (42.6 mg/dL) in the buffered-ruminal fluid made detection of small changes in
NH3N concentration difficult.
Increasing the dose of EFE 11C had no effect on valerate concentration, increased
butyrate concentration and pH (P < 0.01, quadratic), and had a cubic effect on A:P ratio and
concentrations of TVFA, acetate, propionate, isobutyrate, isovalerate and NH3N (P < 0.05; Table
4-3). Total VFA concentration was increased (P < 0.05) by applying 11C at the 0.5× and 3×
doses (8.2 and 8.6%, respectively), due to corresponding increases in propionate concentration
(P < 0.05) at the 0.5×, 1×, and 3× doses (14.7, 9.2 and 16.5%, respectively). All doses of 11C
similarly decreased the A:P ratio and butyrate concentration was only increased by the 3× dose
(15.3%). As was the case for 1A and 2A, the increased TVFA concentration due to applying 11C
has the potential to increase the supply of energy to lactating cows. Furthermore, the increased
butyrate concentration may induce proliferation of ruminal epithelial growth, thus increasing the
VFA absorption capacity of the rumen (Gorka et al., 2009). Butyrate also provides building
blocks for de novo synthesis of fat in the mammary gland (Mohammed et al., 2011) therefore
applying 11C at the 3× dose may increase milk fat concentration and or yield in lactating dairy
cows. In addition, the 0.5×, 1×, and 3× doses increased isobutyrate concentration by 48.2, 34.4,
103
23.8 and 38.6% and the 0.5×, 1×, 2×, and 3× doses increased isovalerate concentrations by 26.4,
18.8, 15.9, and 18.3% respectively. This increased supply of branch-chained VFA likely
contributed to the corresponding increases in NDFD because ruminal cellulolytic bacteria require
them for optimal growth (Liu et al., 2009).
Increasing the dose of 13D resulted in no effects on NH3N, linear (P < 0.10, tendency)
and quadratic (P < 0.01) increases in concentrations of acetate and valerate, respectively and
cubic effects on other fermentation measures (P < 0.05; Table 4-3). Total VFA concentration
was increased (P < 0.05) by all doses of 13D (8.1, 5.9, 6.3, and 11.4% for 0.5×, 1×, 2×, and 3×,
respectively) largely due to increases (P < 0.05) in propionate concentration at the 0.5×, 2×, and
3× doses (11.0, 9.2 and 16.5%, respectively). Butyrate concentration was only increased by the
3× dose (16.5%) and acetate concentration was unaffected by dose. Consequently, the A:P ratio
was decreased by all doses of 13D. Concentrations of isobutyrate (P < 0.05; 26.5, 17.5, 20.1, and
31.8%) and isovalerate (P < 0.05; 17.3, 17.8, 15.4, and 19.7%) were increased by the 0.5×, 1×,
2× and 3× doses, respectively. However, valerate concentration was only increased by the 1× and
2× doses (P < 0.05; 48.5 and 27.4%, respectively). Therefore, despite having the least activities
and protein concentration of the EFE examined, 13D had important beneficial effects on most
digestibility and fermentation measures.
Increasing the dose of EFE 15D resulted in no polynomial effects on acetate, propionate,
NH3N and pH, linear decreases (P < 0.01) in butyrate concentration and A:P ratio, a quadratic
increase in TVFA concentration (P < 0.10, tendency), and cubic effects on concentrations of
isobutyrate, isovalerate and valerate (P < 0.01, Table 4-3). Only the 0.5× dose increased
concentrations of TVFA (6.59%), acetate (6.97%) and isobutyrate (6.88%) and the 0.5 and 2×
doses increased (P < 0.05) propionate concentration. All doses increased the isovalerate
104
concentration, but butyrate concentration was decreased by the 3× dose (- 9.39%), the A:P ratio
was reduced (P < 0.05) by the 2× and 3× doses (- 3.20 and 4.37%), and valerate concentration
was decreased by all doses except 1×. Beneficial fermentation product responses to applying
EFE 15D were few and small relative to those for the other EFE. This reflects the minor
increases in digestibility caused by this EFE, which would have limited availability of substrates
for microbial fermentation.
Experiment 2: Effects of EFE Dose on Measures of Preingestive Hydrolysis
Increasing the dose of EFE 1A increased DM loss (P < 0.05, nonlinear) and increased
concentrations of FER (P< 0.01, linear), COU and WSC (P < 0.01, nonlinear), and reduced (P <
0.05, nonlinear) concentrations of NDF, HEM, ADF, and CEL (P < 0.05, nonlinear; Table 4-4).
The DM loss response presumably occurred via increased solubility and or particle size
reduction, which increases the substrate surface area exposed to ruminal microbes, and could
thereby improve digestibility (Bansal et al., 2009). Consequently, the increases in digestibility
measures by this EFE may have been caused partly by the increased DM losses. Krueger et al.
(2008) also reported increased DM loss (1.8 percentage unit increase) in bahiagrass treated with
a mixture of pure cellulase, xylanase and ferulic acid esterase from Aspergillus spp.,
Orpinomyces spp. and Clostridium themocelllum, applied at doses of 2, 2 and 1 g/100 g of
bahiagrass DM, respectively. In general, the 3× dose was or was among the most effective doses
at increasing hydrolysis of NDF (- 6.0%), HEM (- 6.0%), and ADF (- 5.8%), CELD (- 6.8%, P <
0.05) and releasing WSC, FER and COU (112.8, 27.2 and 7.2%, respectively; P < 0.05) from
cell walls. Such reductions in fiber concentration could increase voluntary intake because they
reduce negative effects of gut fill on intake (Mertens, 2007). Therefore, by increasing intake,
application of this EFE could increase the supply of nutrients that are critical during early
lactation (VandeHaar, 2005). Since fiber detergent analysis were done in sequence and included
105
fiber-bound ash except for ADL, it is important to note that some acid detergent soluble ash
might have contributed to increase HEM concentration, although this contribution should have
be the same across treatments. The FER and COU released during the hydrolysis resulted from
the action of ferulic and p-coumaric acid esterases. When bound to hemicellulose, these phenolic
compounds decrease the rate and possibly the extent of polysaccharide digestion, especially
when FER is cross-linked to lignin (Grabber et al., 1998b). Therefore, their release from cell
walls by EFE 1A partially explains the increase in digestibility by the EFE. Beauchemin et al.
(2004) and Adesogan (2005) emphasized the importance of including phenolic acid esterases in
EFE products meant to improve forage digestibility, animal productivity and nutrient use
efficiency.
Applying increasing doses of EFE 2A also resulted in increases in DM loss, hydrolysis of
NDF, HEM, ADF, and CEL, and release of WSC, FER, and COU (P < 0.05, cubic; Table 4-4).
As was the case for EFE 1A, the 3× dose was or was among the most effective (P < 0.05) doses
at hydrolyzing NDF (- 11.6%) and HEM (- 18.8%; P < 0.05) and hence increasing DM loss
(21.5%) and concentrations of WSC (242%), FER (129%) and COU (35.5%). The increase in
available sugars due to increasing the 2A dose likely explains the corresponding increases in
propionate concentration by EFE 2A and other EFE in this study. When availability of
fermentable substrates in the rumen is high, the fermentation pattern shifts from one ending in
acetic acid to propionic acid to dispose of excess reducing power (France and Dijkstra, 2005).
Among all the EFE examined in this study, 2A resulted in the greatest reductions in the
hemicellulose fraction, reflecting its greater xylanase activity.
Applying EFE 11C at increasing doses also increased DM loss, hydrolysis of HEM,
NDF, ADF, and CEL, and release of WSC, FER and COU (P < 0.05, nonlinear; Table 4-4). As
106
for EFE 1A and 2A, the 3× dose was or was among the most effective doses at increasing DM
loss (22.7%), and increasing release of WSC (281%), FER (84.0%) and COU (24.3%; P < 0.05).
Krueger et al. (2008) also reported that an EFE rich in esterase activity from Humicola spp.
increased release of ether-linked FER and ester-linked COU from bermudagrass cell walls.
Among the EFE, 11C resulted in the greatest hydrolysis of cellulose, which agrees with the fact
that only this EFE increased ADFD and CELD.
Increasing the dose of 13D linearly increased DM loss and HEM hydrolysis, nonlinearly
increased hydrolysis of NDF, ADF and CEL, and increased saccharification of BH (Table 4-4).
Surprisingly and for unknown reasons, application of all doses of 13D similarly reduced FER but
increased COU. The 3× dose resulted in the greatest DM loss (8.7%) and WSC concentration
(31.3%; P < 0.05) and among the greatest increases in NDF and HEM hydrolysis. In general,
effects of 13D on measures of pre-ingestive hydrolysis were limited relative to those of the other
EFE, reflecting its low enzymatic activities and protein concentration. Nevertheless, it increased
all measures of in vitro digestibility perhaps because it contained important unmeasured
activities and auxiliary proteins (e.g. swollenin, polysaccharide monooxygenase).
Applying increasing doses of EFE 15D linearly increased DM loss and nonlinearly
increased hydrolysis of NDF, HEM, ADF and CEL and release of WSC, FER and COU (P <
0.01; Table 4-4). Application of 15D reduced ADF and CEL concentrations but all doses had the
same effect (P > 0.05). In contrast, the 3× dose resulted in the least NDF (- 5.6%) and HEM (-
7.7%) concentrations, the greatest DM loss (9.9%) and the greatest concentrations of WSC
(62.1%) and FER (92.9%), whereas the 2× and 3× dose resulted in the greatest COU
concentration (20.4%; P < 0.05). In general, 15D seemed more effective at increasing pre-
ingestive hydrolysis than 13D, yet unlike the latter, it had few and relatively small beneficial
107
effects on most in vitro digestibility and fermentation measures. Krueger et al. (2008) also
reported that an EFE from Humicola spp. increased preingestive hydrolysis but did not improve
the NDFD of Tifton 85 bermudagrass hay. These responses suggest that 15D can only hydrolyze
easily accessible fiber fractions but cannot remove the main barriers to fiber digestion by ruminal
microbes. Developing methods to more precisely identify and accurately quantify such barriers
will facilitate development of more effective EFE products.
Conclusions
Increasing the EFE dose resulted in increased pre-ingestive fiber hydrolysis of
bermudagrass haylage as well as increases in in vitro digestion and fermentation but the extent
and nature of the responses differed with the EFE and dose. These results indicate that EFE
doses can be manipulated to increase pre-ingestive fiber hydrolysis and digestion, which would
likely increase intake and supply of critical nutrients to lactating cows. In particular, the
consistent increase in sugar release as doses of each of the EFE increased implies increased
supply of gluconeogenic energy substrates like propionate in the rumen, which could increase
milk production by cows. The highest EFE doses were consistently the most effective at
increasing pre-ingestive hydrolysis but not in vitro digestibility or fermentation. This indicates
that using measures of pre-ingestive hydrolysis such as release of WSC, to choose doses that will
increase animal responses will probably be misleading. Baseline EFE doses examined in the
study were recommended by the manufacturers such that some EFE were applied at higher doses
than others. Based on their ability to increase NDFD, EFE with lower baseline manufacturer-
stipulated doses (1A and 2A) benefited from increasing the dose whereas those with greater
baseline doses (13D and 15D) benefited from reducing the dose. Consequently the optimal dose
for the former was 2× the baseline dose, whereas that for the latter was 0.5× the baseline dose.
The best dose for 11C, which had an intermediate baseline dose was the manufacturer-stipulated
108
dose. Dose rates that enhanced digestibility also typically increased fermentation product
concentrations and reduced the A:P ratio, and hence increased energetic efficiency.
109
Table 4-1. Endoglucanase, xylanase, exoglucanase, β-glucosidase (μmol of sugar released / min× g) and ferulic acid esterase (nmol of
ferulic acid released / min ×g) activities, protein concentration (mg/g), form, dose (g/kg of bermudagrass DM), and
biological source of exogenous fibrolytic enzyme (EFE) preparations applied to bermudagrass haylage.
Parameter EFE
1A 2A 11C 13D 15D SD
Activity
Endoglucanase
1,693
3,624
1,506
286
70
112
Xylanase 1,276 29,301 1,703 86 6,499 221
Exoglucanase 1.68 0.84 0.97 0.29 0.29 0.10
β-glucosidase 10.1 11.7 12.7 1.9 0.1 0.9
Ferulic acid
esterase 2.18 1.46 6.30 2.35 2.57 1.27
Protein concentration 65.3 111 81.1 18 28.3 25.0
Form Liquid Liquid Liquid Liquid Liquid
Dose
0.5× 1.17 1.17 5.2 7.8 7.8
1× 2.33 2.33 10.4 15.6 15.6
2× 4.66 4.66 20.8 31.2 31.2
3× 6.99 6.99 31.2 46.8 46.8
Biological Source Trichoderma
reesei T. reesei T. reesei
Aspergillus
oryzae A. oryzae
110
Table 4-2. Effects of the dose of exogenous fibrolytic enzymes (EFE) on in vitro true dry matter
(DMD), neutral detergent fiber (NDFD), hemicellulose (HEMD), acid detergent fiber
(ADFD), and cellulose (CELD) digestibility, and lignin disappearance (ADLD) of a
4-wk regrowth bermudagrass haylage (Experiment 1).1
Dose DMD
(%)
NDFD
(%)
HEMD
(%)
ADFD
(%)
CELD
(%)
ADLD
(%)
EFE 1A
0 × 48.6abc 35.1ab 31.4a 40.5c 46.1b -14.7a
0.5 × 47.7a 35.0ab 32.5ab 37.1ab 41.6a -4.4b
1 × 47.9ab 34.2a 31.8a 36.1a 41.7a -3.5b
2 × 49.7c 37.7c 35.0c 39.7c 45.6b -7.2ab
3 × 49.0bc 36.5bc 33.8bc 38.7bc 44.0b -11.4ab
Contrast C** C* L** C** C** Q**
SEM 0.40 0.73 0.67 0.76 0.80 3.02
EFE 2A
0 × 48.6a 35.1a 31.4a 40.5ab 46.1a -14.7a
0.5 × 49.7ab 37.4b 34.8b 40.8ab 45.1a 5.9b
1 × 50.1b 38.0bc 34.7b 41.8b 47.9a 2.8b
2 × 50.6b 38.9c 36.5c 41.4b 45.8a 6.1b
3 × 50.4b 38.0bc 36.1bc 39.2a 44.8a 0.8b
Contrast Q* Q** Q** Q* n.s. C**
SEM 0.42 0.67 0.64 0.73 0.76 3.5
EFE 11C
0 × 48.6a 35.1a 31.4a 40.5a 46.1a -14.7a
0.5 × 50.4b 38.9b 35.4b 42.2ab 45.8a 7.4b
1 × 51.4bc 40.8c 38.5c 43.2bc 47.1a 6.1b
2 × 51.9c 40.4bc 38.5c 42.3abc 46.9a 5.0b
3 × 52.2c 41.7c 39.4c 44.3c 49.3b 4.1b
Contrast Q** C** C** L** L** C**
SEM 0.40 0.66 0.58 0.75 0.73 2.93
1Linear (L), quadratic (Q), cubic (C); + (P < 0.10), * (P < 0.05); ** (P < 0.01); Within the same
EFE treatment, means with different superscripts differed (P < 0.05).
111
Table 4-2. Continued 1
Dose DMD
(%)
NDFD
(%)
HEMD
(%)
ADFD
(%)
CELD
(%)
ADLD
(%)
EFE 13D
0 × 48.6a 35.1a 31.4a 40.5a 46.1b -14.7a
0.5 × 51.1b 39.7b 37.5c 42.7b 46.4b 12.8c
1 × 50.7b 38.5b 37.0bc 42.9b 46.0b 15.2c
2 × 49.5a 37.0c 35.7b 39.1a 42.6a -4.5ab
3 × 51.2b 39.1b 37.5c 40.8a 45.3b 4.8bc
Contrast C* C** C** C** C** C**
SEM 0.35 0.49 0.60 0.66 0.52 5.2
EFE 15D
0 × 48.6a 35.1a 31.4a 40.5b 46.1c -14.7ab
0.5 × 49.1a 36.9b 34.5b 38.0a 44.3ab -12.6bc
1 × 49.2a 35.9ab 33.9b 37.9a 43.1a -4.1c
2 × 48.8a 36.2ab 34.3b 38.4a 44.7bc -11.9bc
3 × 48.9a 35.2a 33.3b 37.7a 45.2bc -22.4a
Contrast n.s. Q+ Q** C* C* Q**
SEM 0.39 0.53 0.55 0.58 0.56 3.03
1Linear (L), quadratic (Q), cubic (C); + (P < 0.10), * (P < 0.05); ** (P < 0.01); Within the same
EFE treatment, means with different superscripts differed (P < 0.05).
112
Table 4-3. Effects of the dose of exogenous fibrolytic enzymes (EFE) on concentrations of total volatile fatty acids (TVFA), acetate,
propionate, butyrate, isobutyrate, isovalerate, valerate, NH3N,acetate to propionate ratio (A:P) and pH of the filtrate
obtained from fermentation of a 4-wk regrowth of bermudagrass haylage (Experiment 1)1
Dose TVFA
(mM)
Acetate
(mM)
Propionate
(mM)
Butyrate
(mM) A:P
Isobutyrate
(mM)
Isovalerate
(mM)
Valerate
(mM)
NH3N
(mg/dL) pH
EFE 1A
0 × 60.7a 37.3a 10.9a 5.75c 3.43a 1.89a 2.08a 2.70a 42.6a 6.96a
0.5 × 59.6a 36.7a 10.9a 4.99a 3.37ab 1.84a 2.24b 2.61a 41.8a 7.01b
1 × 59.4a 36.6a 10.9a 4.90a 3.38ab 1.84a 2.21ab 2.82a 42.6a 6.98ab
2 × 61.1a 37.7a 11.4a 5.13ab 3.31bc 1.92a 2.19ab 2.81a 42.9a 6.97a
3 × 61.6a 37.5a 11.6a 5.34b 3.25c 1.92a 2.31b 2.78a 42.7a 6.97a
Contrast n.s. n.s. L+ C** L** n.s. C* n.s. n.s. C*
SEM 1.35 0.77 0.33 0.096 0.032 0.081 0.048 0.138 0.52 0.01
EFE 2A
0 × 60.7a 37.3a 10.9a 5.75a 3.43b 1.89a 2.08a 2.70b 42.6a 6.96ab
0.5 × 63.8bc 37.9a 12.1b 6.47b 3.16a 2.27b 2.24b 2.71b 42.1a 6.97ab
1 × 64.8c 38.7ab 12.6bc 6.29b 3.07a 2.30bc 2.29b 2.59b 40.2a 6.95a
2 × 61.6ab 37.1a 12.0b 5.49a 3.11a 2.10ab 2.31b 2.33a 42.0a 6.98b
3 × 66.5c 40.1b 13.1c 5.74a 3.08a 2.63c 2.46c 2.52ab 41.1a 6.98b
Contrast C** C** C** C** C** C** C** C* n.s. L*
SEM 1.32 0.72 0.35 0.156 0.042 0.127 0.034 0.079 0.62 0.01
EFE 11C
0 × 60.7a 37.3a 10.9a 5.75a 3.43b 1.89a 2.08a 2.70a 42.6ab 6.96a
0.5 × 65.7b 39.3a 12.5b 5.89a 3.16a 2.80b 2.63c 2.57a 43.0abc 7.04b
1 × 63.1ab 37.9a 11.9b 5.73a 3.20a 2.54b 2.47b 2.53a 43.6bc 7.05b
2 × 62.4ab 37.3a 11.8ab 5.75a 3.17a 2.34ab 2.41b 2.56a 42.0a 7.07b
3 × 65.9b 39.0a 12.7b 6.63b 3.09a 2.62b 2.46b 2.62a 44.3c 7.06b
Contrast C** C* C* Q** C** C** C** n.s. C* Q**
SEM 1.38 0.72 0.38 0.158 0.044 0.184 0.052 0.060 0.54 0.02 1Linear (L), quadratic (Q), cubic (C); + (P < 0.10), * (P < 0.05); ** (P < 0.01); Within the same EFE treatment, means with different
superscripts differed (P < 0.05).
113
Table 4-3. Continued 1
Dose TVFA
(mM)
Acetate
(mM)
Propionate
(mM)
Butyrate
(mM) A:P
Isobutyrate
(mM)
Isovalerate
(mM)
Valerate
(mM)
NH3N
(mg/dL) pH
EFE 13D
0 × 60.7a 37.3a 10.9a 5.75a 3.43d 1.89a 2.08a 2.70a 42.6a 6.96a
0.5 × 65.6bc 39.5a 12.1cd 6.02a 3.26b 2.39b 2.44bc 3.26ab 42.9a 7.05b
1 × 64.3b 38.2a 11.4ab 5.95a 3.34c 2.22b 2.45bc 4.01c 43.8a 7.07b
2 × 64.5b 38.9a 11.9bc 5.69a 3.28bc 2.27b 2.40b 3.44bc 42.9a 7.05b
3 × 67.6c 39.7a 12.7d 6.70b 3.13a 2.49b 2.49c 3.22ab 43.0a 7.04b
Contrast C* L+ C* C* C** C* C** Q** n.s. C**
SEM 1.1 0.71 0.24 0.201 0.027 0.105 0.021 0.267 0.56 0.01
EFE 15D
0 × 60.7a 37.3a 10.9a 5.75b 3.43a 1.89a 2.08a 2.70c 42.6a 6.96a
0.5 × 64.7b 39.9b 11.7b 6.06b 3.41a 2.02b 2.34b 2.40b 42.7a 6.94a
1 × 60.8a 37.3a 10.9a 5.75b 3.42a 1.89a 2.30b 2.63c 42.9a 6.97a
2 × 62.3ab 38.2a 11.5b 5.76b 3.32b 1.88a 2.25b 2.44b 43.5a 6.95a
3 × 59.8a 37.0a 11.3ab 5.21a 3.28b 1.85a 2.32b 2.14a 42.6a 6.95a
Contrast Q+ n.s. n.s. L** L** C** C** C** n.s. n.s.
SEM 0.98 0.53 0.19 0.17 0.018 0.019 0.040 0.047 0.43 0.01 1Linear (L), quadratic (Q), cubic (C); + (P < 0.10), * (P < 0.05); ** (P < 0.01); Within the same EFE treatment, means with different
superscripts differed (P < 0.05).
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Table 4-4. Effects of the dose of different exogenous fibrolytic enzymes (EFE) on DM loss and
concentrations of NDF, hemicellulose (HEM), ADF, cellulose (CEL), lignin (ADL)
water-soluble carbohydrates (WSC), ferulic acid (FER), and p-coumaric acid (COU)
after preingestive hydrolysis of a 4-wk regrowth of bermudagrass haylage
(Experiment 2)1
Dose
DM
loss
(%)
NDF
(%)
HEM
(%)
ADF
(%)
CEL
(%)
ADL
(%)
WSC
(%)
FER
(μg/g)
COU
(μg/g)
EFE 1A
0 × 17.2a 71.4e 35.1c 36.3c 32.3c 4.1 2.11a 169a 152a
0.5 × 18.5bc 68.4d 33.8b 34.6ab 30.2b 4.3 2.84b 173b 156b
1 × 18.1b 68.2c 33.4ab 34.8b 30.7b 4.1 3.24c 186c 160c
2 × 18.5bc 67.6b 33.3a 34.4ab 30.0a 4.4 3.87d 202d 162d
3 × 18.9c 67.1a 33.0a 34.2a 30.1a 4.1 4.49e 215e 163d
Contrast C* C** C** C** C* n.s. C** L** Q**
SEM 0.20 0.26 0.17 0.17 0.21 0.21 0.029 1.5 0.7
EFE 2A
0 × 17.2a 71.4e 35.1e 36.3b 32.3c 4.1 2.11a 169a 152a
0.5 × 19.0b 67.0d 32.1d 35.0a 31.0b 3.9 4.03b 246b 177b
1 × 19.1b 65.9c 30.9c 35.0a 30.4a 4.6 5.02c 288c 186c
2 × 20.2c 64.3b 29.4b 34.9a 30.6ab 4.3 6.32d 358d 199d
3 × 20.9d 63.1a 28.5a 34.7a 30.1a 4.6 7.21e 387e 206e
Contrast C* C** C** C** C** n.s. C** C* C**
SEM 0.18 0.30 0.21 0.18 0.23 0.20 0.037 2.4 0.9
EFE 11C
0 × 17.2a 71.4c 35.1d 36.3c 32.3c 4.1 2.11a 169a 152a
0.5 × 18.6b 67.8b 33.0c 34.8b 30.8b 4.0 3.54b 194b 170b
1 × 18.7b 67.3b 32.3b 35.0b 30.9b 4.0 4.48c 225c 174c
2 × 20.6c 64.7a 30.8a 33.9a 29.6a 4.3 6.59d 280d 183d
3 × 21.1d 64.1a 30.4a 33.8a 29.7a 4.0 8.03e 311e 189e
Contrast Q** C* Q** Q** Q** n.s. Q** C** C**
SEM 0.17 0.23 0.22 0.15 0.19 0.10 0.077 1.7 0.7 1Linear (L), quadratic (Q), cubic (C); + (P < 0.10), * (P < 0.05); ** (P < 0.01); n.s. (P > 0.10);
Within the same EFE treatment means with different superscripts differed (P < 0.05).
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Table 4-4. Continued 1
Dose
DM
loss
(%)
NDF
(%)
HEM
(%)
ADF
(%)
CEL
(%)
ADL
(%)
WSC
(%)
FER
(μg/g)
COU
(μg/g)
EFE 13D
0 × 17.2a 71.4d 35.1d 36.3b 32.3b 4.1 2.11a 169a 152a
0.5 × 17.4a 69.6c 34.7cd 34.8a 31.1a 3.7 2.37b 161b 158b
1 × 18.1b 69.3ab 34.6bc 34.8a 31.0a 3.8 2.50c 161b 159b
2 × 18.0b 69.1ab 34.2ab 34.9a 31.0a 3.9 2.61d 164b 159b
3 × 18.7c 68.5a 33.9a 34.5a 30.7a 3.8 2.77e 163b 157b
Contrast L** C* L** C** C** n.s. C** C** Q**
SEM 0.17 0.32 0.18 0.19 0.20 0.10 0.029 1.2 1.1
EFE 15D
0 × 17.2a 71.4c 35.1d 36.3b 32.3b 4.1 2.11a 169a 152a
0.5 × 17.3a 69.0b 33.8c 35.2a 31.5a 3.8 2.78b 241b 170b
1 × 17.9b 68.4b 33.3b 35.1a 31.1a 4.0 3.03c 266c 174c
2 × 18.2b 68.5b 33.2b 35.2a 31.3a 3.9 3.28d 307d 182d
3 × 18.9c 67.4a 32.4a 35.0a 31.0a 3.9 3.42e 326e 183d
Contrast L** C** C** C** C** n.s. C** C** C**
SEM 0.16 0.25 0.17 0.15 0.16 0.07 0.022 2.2 1.0 1Linear (L), quadratic (Q), cubic (C); + (P < 0.10), * (P < 0.05); ** (P < 0.01); n.s. (P > 0.10);
Within the same EFE treatment means with different superscripts differed (P < 0.05).
116
CHAPTER 5
EFFECT OF ADDING COFACTORS TO EXOGENOUS FIBROLYTIC ENZYMES ON
PREINGESTIVE HYDROLYSIS, IN VITRO DIGESTIBILITY AND FERMENTATION OF
BERMUDAGRASS HAYLAGE
Background
Exogenous fibrolytic enzymes (EFE) have been applied to cattle diets to improve
digestibility and animal performance but the results have been equivocal (Adesogan, 2005;
Adesogan et al., 2013). This is because the outcome is influenced by numerous factors including
the EFE dose (Eun et al., 2007) and activity composition (Eun and Beauchemin, 2007), the
prevailing pH and temperature (Arriola et al., 2011a), the animal performance level (Schingoethe
et al., 1999), the experimental design (Adesogan et al., 2013), and the fraction and proportion of
the diet to which the enzyme is applied (Krueger et al., 2008a; Dean et al., 2013). Despite their
well-known effects on the activity of certain enzymes (Voet et al., 2010), effects of cofactor
(COF) addition to EFE on the digestibility of forages are unknown. Cofactors are metal ions
required by most enzymes for maintenance of structural integrity and or catalytic activity (Voet
et al., 2010). Such enzymes can be classified as metal-activated enzymes or metalloenzymes.
Cofactors are not required for metal-activated enzymes but when present, they improve the
conformation stability that maximizes their activities (Glusker, 2011). Metalloenzymes require
COF at their active sites, where they serve as substrate templates, inducers of free radicals, and
redox-active COF (Purich, 2011). Among enzymes involved in lignocellulose degradation, only
a few, mostly those grouped in the Auxiliary Activities family have been identified as
metalloenzymes (Harris et al., 2010; CAZy, 2013). Cofactors like Mn2+, Co2+, Fe2+, Ca2+, and
Mg2+ have increased the activity of metal-activated fibrolytic enzymes (BRENDA, 2013) but to
our knowledge, their effects on EFE used in ruminant nutrition have not been examined. The
productivity of dairy cattle in the southeastern U.S. is limited by the low digestibility of tropical
117
grasses like bermudagrass (Cynodon dactylon), which is the most widely grown perennial grass
in the South (Newman, 2007). Recent research identified 5 EFE that improved the NDF
digestibility (NDFD) of bermudagrass haylage and optimized their dose rates (Chapter 3 and 4).
Whether adding COF to these EFE would synergistically increase their hydrolytic effects is
unknown. The objective of this study was to determine the effects of applying 5 COF to EFE on
the in vitro digestibility, fermentation and preingestive fiber hydrolysis of bermudagrass haylage
(BH). The hypothesis was that adding key COF to the EFE would increase their preingestive
hydrolytic effects. Furthermore, adding key COF at an appropriate dose would synergistically
increase the NDFD and fermentation of EFE-treated BH.
Materials and Methods
Bermudagrass Substrate
An established stand of bermudagrass cv. Tifton 85 in Alachua County, Florida was
staged in June 2010 by mowing to a 4-cm stubble in 1 d with a CLAAS 3500 mower conditioner
(CLAAS North America, Omaha, NE) and removing the residue. The grass was fertilized with N
(95 kg/ha), harvested as a 4-wk regrowth on July 7, 2010 as described above, wilted for 2.5 h in
the windrow and rolled into round bales without inoculant treatment, wrapped with 7 layers of 6-
mm plastic, and ensiled for 53 d. Ensiled bermudagrass was chosen over hay because it is more
frequently used by dairy producers due to the high humidity and frequent summer rainfall in
Florida (Staples, 2003). Representative haylage samples were dried at 60oC for 48 h and ground
to pass a 1-mm screen using a Wiley mill (Arthur H. Thomas Company, Philadelphia, PA). The
chemical composition of BH was 49.4% DM and 93.5, 68.1, 34.2, 3.7 and 18.7% of OM, NDF,
ADF, ADL, and CP, respectively (DM basis). The BH also contained 0.36 and 0.27% water -
soluble Ca and Mg, and 24.1, 64.3, and 0.17 mg/L of water-soluble Fe, Mn, and Co, respectively,
as determined by Inductively Coupled Plasma Spectrometry (Beliciu et al., 2012) after
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microwave digestion (CEM Corp., Matthews, NC)at the Dairy One Forage laboratory, Ithaca,
NY.
Enzymes
Five previously selected (Chapter 3) commercial and experimental EFE preparations
provided by 3 manufacturers were evaluated. Their enzymatic activities, protein concentrations,
forms, mineral concentrations, application rates, and biological sources are listed in Table 5-1.
Endoglucanase (Enzyme commission, E.C. 3.2.1.4), exoglucanase (E.C. 3.2.1.91), xylanase
(E.C. 3.2.1.8) and β-glucosidase (E.C. 3.2.1.21) activities were respectively quantified using
carboxymethyl cellulose, avicel, oat-spelt xylan and cellobiose as substrates (Colombatto and
Beauchemin, 2004). Ferulic acid esterase (E.C. 3.1.1.73) activity was measured using ethyl
ferulate (Sigma-Aldrich Corp, St. Louis, MO) as the substrate (Lai et al., 2009). All activities
were measured at 39oC and a pH of 6 to mimic ruminal conditions. Protein concentration was
measured using the Bio-Rad Protein Assay (Bradford 1976) with bovine serum albumin as the
standard (Bio-Rad Laboratories, Hercules, CA). The mineral concentrations of the EFE were
determined as described for the BH. The EFE application rates were chosen because they had
optimized the NDFD of BH in a previous study (Chapter 4).
Screening COF for Synergistic Effects on the Hydrolytic Potential of EFE (Experiment 1)
An experiment was conducted to select the most promising EFE – COF combinations for
improving hydrolysis of BH cell walls prior to consumption by animals. The selection criterion
was the extent of saccharification (percentage of water-soluble carbohydrates, WSC released
from cell walls) during pre-ingestive hydrolysis of BH. The preingestive hydrolysis procedure
described by Krueger et al. (2008b) was used except that the EFE were diluted in nanopure water
(2 mL) instead of citrate–phosphate buffer to avoid the potential chelating effects of citrate.
Cofactors used were chloride salts of divalent cations of Mn, Co, Fe, Ca, and Mg. These
119
cofactors were chosen because they had increased the activity of fibrolytic enzymes on pure
substrates (BRENDA, 2013). Each COF was added to the EFE solution to achieve a final
concentration of 1 mM (Lai et al., 2009) and sodium azide was added (0.02% w/v) to prevent
substrate degradation by microbes (Krueger et al., 2008b). The EFE – COF solution was added
to 50-mL tubes containing 0.5 g of BH and the tubes were incubated for 24 h at 25oC in
quadruplicate. For each EFE, two blank tubes without substrate were included to correct for
contributions from the EFE. After the incubation, 15 mL of nanopure water was added and the
suspension was shaken (Eberbach reciprocating shaker, Model 6000, Eberbach corporation, Ann
Arbor, MI) for 1 h at 260 oscillations/min, filtered through previously dried (60oC for 48 h) and
weighed 125-mm Whatman 451 filter paper (Fisher Scientific, Pittsburgh) and filtrate samples
were frozen (-20oC). Residues were dried at 60oC for 48 h in a forced draught oven, weighed,
and analyzed sequentially for NDF and ADF (Van Soest et al., 1991) using an ANKOM 200
Fiber Analyzer (ANKOM, Macedon, NY). Amylase was used for NDF determination but no
sodium sulfite was added. The NDF and ADF results were expressed inclusive of residual ash.
Hemicellulose (HEM) was estimated as the difference between NDF and ADF. Residue and
sample weights and DM concentrations were used to calculate DM losses. Filtrate samples were
thawed and analyzed for WSC (DuBois et al., 1956) and ferulic (FER) and p-coumaric (COU)
acids (Bio-Rad, 2011) using a Merck Hitachi Elite LaChrome High Performance Liquid
Chromatograph system (Hitachi, L2400, Tokyo, Japan) and a Bio-Rad Aminex HPX-87H
column (Bio-Rad Laboratories, Hercules, CA).
Effects of Adding Increasing Doses of COF to EFE on in Vitro Digestibility (Experiment 2)
The most promising EFE – COF combinations selected in Experiment 1 (Mn2+ + EFE
11C and FE2+ + EFE 2A or 13D) were evaluated with a 24 h in vitro ruminal digestibility assay
120
(Goering and Van Soest, 1970). The COF were added to the EFE solution at doses of 0, 0.1, 1,
and 10 mM in quadruplicate. The experimental procedure was similar to that for Experiment 1
except that after the 24 h of incubation at 25oC, buffered (Goering and Van Soest, 1970)-ruminal
fluid (52 mL) was added and the suspension was incubated for 24 h at 39oC. The fermentation
was stopped by placing the tubes on ice and contents of tubes were filtered through pre-weighed
125-mm Whatman 451 paper (Fisher Scientific, Pittsburgh). Filtrate samples were immediately
analyzed for pH using an Accumet XL25 pH meter (Fisher Scientific, Pittsburgh, PA), acidified
with 50% H2SO4 (1% v/v of rumen fluid sample), centrifuged at 8,000 × g for 20 min and the
supernatant was frozen (-20oC). Residues were dried at 60oC for 48 h, weighed and stored at
room temperature for subsequent analysis. No sodium azide was used in the EFE solution and
the Control treatment consisted of only nanopure water. For each EFE treatment, two blank tubes
without substrate were included to correct for EFE effects. The ruminal fluid was
representatively collected 3 h after feeding at 0800 h, by aspiration, from two non-lactating non-
pregnant ruminally-cannulated Holstein cows fed coastal bermudagrass hay ad libitum
supplemented with corn (0.5 kg), cottonseed hulls (0.5 kg), soybean meal (1 kg), and a vitamin-
mineral mix (37.5 g). The ruminal fluid collection protocol was approved by the University of
Florida, Institute of Food and Agricultural Sciences Animal Research Committee. Ruminal fluid
was filtered through four layers of cheesecloth prior to use and all tubes and the artificial saliva
were pre-warmed at 39oC before ruminal fluid addition. Dried residues were analyzed for NDF,
ADF and ADL, and hemicellulose and cellulose were calculated as previously described. The
ADL was determined using a modification of the method of Van Soest et al. (1991) for a Daisy
II incubator (ANKOM, Macedon, NY). Residue and original sample weights and their DM and
fiber concentrations were used to calculate true DM, NDF, hemicellulose and cellulose
121
digestibility (DMD, NDFD, HEMD, and CELD, respectively). Filtrate samples were analyzed
for concentrations of volatile fatty acids (VFA; Muck and Dickerson, 1988) with the High
Performance Liquid Chromatograph system described earlier. Ammonia-N was determined with
a Technicon Auto Analyzer (Technicon, Tarrytoen, NY) and an adaptation of the Noel and
Hambleton (1976) procedure that involved colorimetric N quantification.
Statistical Analyses
Experiment 1 was analyzed as a completely randomized design with a 6 (COF) × 6 (EFE)
factorial treatment arrangement and 4 replicates per treatment.
The model used to analyze the preingestive hydrolysis data was:
Yijk = µ + Ti + Cj + TCij + Eijk
Where:
µ = general mean
Ti = effect of EFE i
Cj = effect of cofactor j
TCij = effect of the EFE i × cofactor j interaction
Eijk = experimental error
The GLM procedure of SAS v.9.1 (2012) was used to analyze the data. Fisher’s F-
protected least significance difference test was used to compare COF means within EFE.
Experiment 2 was analyzed as a randomized complete block design with a 2 (EFE) × 4 (COF
dose) factorial treatment arrangement and 2 runs each with 4 replicates / treatment.
The model used to analyze digestibility and fermentation data was:
Yijk = µ + Ti + Dj + TDij + TRik + DRik + TDRijk + Eijk
Where:
µ = general mean
122
Ti = effect of EFE i
Dj = effect of cofactor dose j
Rk = effect of run k
TDij = effect of the EFE i × cofactor dose j interaction
TRik = effect of the EFE i × run k interaction
DRik = effect of the cofactor dose j × run k interaction
TDRijk = effect of the EFE i × cofactor dose j × run k interaction
Eijk = experimental error
Data from each EFE were analyzed separately with the GLM procedure of SAS. The
model included COF dose, EFE and their interaction. Polynomial contrasts were used to
determine dose effects and the Fisher’s least significance difference test was used to compare
least square means across doses. Significance was declared at P <0.05 and tendencies at P >
0.05 < 0.10.
Results and Discussion
Experiment 1: Effects of Cofactor Addition on Preingestive Hydrolysis
Compared to the Control (1.90%), applying each COF alone had no effect (P > 0.05) on
saccharification, but saccharification was increased (P < 0.05) by applying EFE 1A (96.3%
increase), 2A (238%), 11C (83.7%) or 15D (35.3%) alone whereas applying 13D alone was not
effective (Table 5-2). Effects of COF addition to EFE on saccharification depended on the EFE –
COF combination (P < 0.001). Saccharification was increased (P < 0.05) by adding Fe2+ to 1A (
4.83%), by adding Mg2+, Ca2+, Fe2+, Co2+ or Mn2+ to 2A (3.27, 2.64, 9.64, 5.75, and 5.29%
increase, respectively), Mg2+, Ca2+, Fe2+, Co2+ or Mn2+to 11C (13.18 14.61, 23.50, 23.21, and
38.4%, respectively), Fe2+, Co2+ or Mn2+ to 13D (21.88, 14.06, and 13.02%, respectively), and
Fe2+ or Co2+ to 15D (13.62 and 7.39%, respectively).
123
Increases in saccharification due to adding COF to EFE suggest that the EFE contained
metal-activated enzymes since almost all fiber-degrading glycoside hydrolyses catalyze acid-
base reactions and are not metalloenzymes (Harris et al., 2010). The only cellulolytic enzyme
known to require a COF for hydrolysis is polysaccharide monooxygenase, which requires copper
and catalyzes oxidation-reduction reactions (Quinlan et al., 2011). This enzyme was identified as
one of the main activities in EFE 2A and 11C using isobaric tags for relative quantification
(iTRAQ)-based quantitative proteomics combined with mass spectrometry (Adesogan et al.,
2013). To our knowledge, this is the first report of the effects of COF addition to EFE on
saccharification of forage cell walls, though others have investigated effects of COF addition to
fibrolytic enzymes on hydrolysis of pure substrates (Tejirian and Xu, 2010) or steam-pretreated
biofuel biomass (Bin and Hongzhang, 2010).
Singh et al. (1990) reported that activity of exoglucanase II (E.C. 3.2.1.91, which acts on
the non-reducing end of cellulose) from Aspergillus niger was increased 1.82-, 1.51-, 1.40- and
1.85- fold when undisclosed doses of Mn2+, Ca2+, Mg2+, or Co2+ were added, respectively.
Exoglucanases account for up to 80% of the secretome of Trichoderma reesei under cellulose-
inducing conditions (Glass et al., 2013) and this activity is critical for maximum saccharification.
Furthermore, T. reesei endoglucanase activity (E.C. 3.2.1.4) on carboxymethyl cellulose was
increased by adding undisclosed doses of Mn2+, Fe2+, and Co2+, but inhibited by Ca2+ and Mg2+
(Saad and Fawzi, 2004). Also, adding 5 mM of Mn2+to β-glucosidase (E.C. 3.2.1.21) from A.
oryzae resulted in a 1.8-fold increase in activity but adding 5mM of Fe3+resulted in a 7.7-fold
decrease in activity (Riou et al., 1998). Cofactor addition also has increased or inhibited the
activity of hemicellulolytic enzymes. For instance, John and Schmidt (1988) reported that the
activity of β-xylosidase (E.C. 3.2.1.37) from Trichoderma viride was increased by adding 1 mM
124
of Fe2+ or Mn2+, but inhibited by adding 0.1 mM of Ca2+. The EFE tested in this study contained
various fibrolytic activities, hence it is challenging to identify which specific activities benefited
from COF addition. Nevertheless, it is noteworthy that Fe2+ addition consistently improved
saccharification by all the EFE. This may be because Fe2+ increased the activities of β-xylosidase
and endoglucanase, since both of these were likely present in the EFE and they have both been
increased by Fe2+ addition in previous research (John and Schmidt, 1988; Saad and Fawzi,
2004). The greatest increase in saccharification occurred when Fe2+was added to 11C perhaps
because relative to the other EFE, 11C had one of the lowest Fe to protein ratios (0.017 vs.
0.034) and the lowest Fe to endoglucanase ratio (0.0009 vs. 0.003).Adding Co2+ also increased
saccharification by all EFE except 1A. This agrees with previous work indicating that adding
Co2+ increased exoglucanase (Singh et al., 1990) and endoglucanase activities (Saad and Fawzi,
2004), which determine the rate of cellulose digestion (Zhang and Lynd, 2004). The synergistic
increase in saccharification due to adding Co2+ to EFE was greatest for 11C, probably because it
had one of the lowest Co to protein ratios (0.0003 vs. 0.0007).Adding Mn2+ to 2A, 11A and 13D
resulted in 1.05-, 1.38-, and 1.13-fold increases in saccharification. This agrees with studies
showing that adding Mn2+ increased the activity of exoglucanases (Singh et al., 1990),
endoglucanases (Saad and Fawzi, 2004), β-glucosidase (Riou et al., 1998), and β-xylosidase
(John and Schmidt, 1988). That Mn2+ increased the activity of several fibrolytic enzymes
explains why its greatest effects were on 11C and 2A, which had among the greatest total
cellulolytic activities and the lowest Mn to protein ratios (0.004 and 0.004 vs. 0.005),
respectively. Adding Mg2+or Ca2+ only improved saccharification by 2A and 11C reflecting their
low Mg or Ca to protein ratios relative to the other EFE (0.85 and 0.53 vs. 1.14, respectively).
125
Also, because they are macrominerals, the concentrations of these COF in the EFE were at least
ten times greater than those of the other COF, which are microminerals.
Compared to the Control (71.1% NDF), applying each COF alone had no effect (P >
0.05) on NDF hydrolysis. Applying EFE 1A (- 3.94%), 2A (- 8.72%), 11C (- 4.36%) or 15D (-
2.25%) alone increased (P < 0.05) NDF hydrolysis whereas applying 13D alone did not (Table
5-2). Effects of EFE addition on NDF hydrolysis tended (P = 0.066) to be influenced by COF
addition. Theoretically, most WSC released during cell wall saccharification by EFE should have
been generated by NDF hydrolysis, thus NDF values were expected to decrease as WSC
concentrations increased due to saccharification. However, NDF analysis is less precise than
many other chemical components of forages such as WSC as evidenced by the greater coefficient
of variation recommended for NDF assays versus those of other forage chemical components
(Galyean, 2010). The variability in the NDF concentrations in this study (SEM = 0.46% units of
BH DM) was identical to the average increase in WSC due to COF addition, thus preventing
detection of differences in NDF due to saccharification. In theory, using the Uppsala dietary fiber
scheme (Theander et al., 1995) instead of the NDF scheme to evaluate EFE effects on cell wall
saccharification may be more appropriate because it quantifies individual cell wall
polysaccharides rather than detergent soluble or insoluble fiber fractions. However, the Uppsala
dietary fiber results are less precise than those from the NDF scheme (SD of 3.2 vs. 1.3%,
respectively; Mertens, 2003).
Compared to the Control (35.4%), applying each COF alone had no effect on HEM
hydrolysis. Applying EFE 1A (- 5.37%), 2A (- 15.82%), 11C (- 6.78%) or 15D (- 4.52%) alone
reduced (P < 0.05) HEM concentration, whereas applying 13D alone did not (Table 5-2).
126
Since NDF and ADF detergent analysis were conducted in sequence and expressed
inclusive of residual ash, losses of acid detergent-soluble ash may have overestimated HEM
concentrations, but this effect should have been the same across treatments.
Effects of EFE addition on HEM were not affected by COF addition (P = 0.23). In our
previous studies, HEM was the fraction mostly hydrolyzed by EFE (Chapter 3 and 4). Although
the structure of xylan is more complex than cellulose due to its substitutions, which necessitate
more enzymes with different specificities for its hydrolysis, xylan does not form tightly packed
crystalline structures like cellulose and is, thus, more accessible to enzymatic hydrolysis (Saha,
2003).
Compared to the Control (35.7%), applying each COF alone had no effect on ADF
hydrolysis except for Mn2+ (- 1.96%), which did not elicit a corresponding increase in
saccharification. The reason for the latter is unclear. Applying EFE 1A (- 2.52%), 2A (- 1.96%)
or 11C (- 1.96%) alone reduced (P < 0.05) ADF concentration whereas applying 13D and 15D
alone did not (Table 5-2). Effects of COF addition to EFE on ADF hydrolysis depended on the
EFE – COF combination (P < 0.05) yet adding COF did not affect ADF hydrolysis by any EFE.
Compared to the Control (17.3%), applying each COF alone had no effect (P > 0.05) on
DM loss, which estimates particle size reduction. However, applying EFE 1A (6.36%), 2A
(15.60%) or 11C (12.14%) alone increased (P < 0.05) DM loss whereas applying 13D or 15D
alone did not. Effects of COF addition to EFE on DM loss depended on the EFE – COF
combination (P < 0.001). DM loss was increased (P < 0.05) by adding Ca2+ and Fe2+ to 1A (4.89
and 3.80%); Mg2+, Fe2+, and Co2+ to 2A (5.50, 5.50, and 7.50%); Mg2+, Ca2+, Fe2+, Co2+, and
Mn2+ to 11C (8.25, 3.61, 4.64, 4.12, and 4.12%), respectively. No changes were observed when
COF were added to EFE 13D and 15D.
127
Surprisingly, adding Ca2+, Fe2+, or Co2+ alone increased the concentration of FER to 183,
174, and 180 μg/g, respectively, relative to the value for the Control (158 μg/g; P < 0.05), though
adding Mg2+ or Mn2+ had no effect (P > 0.05). The mechanism by which certain COF increased
the FER concentration in the absence of EFE is unknown. Hydration during COF addition may
be implicated and the COF may have stimulated endogenous FER esterases in bermudagrass
haylage or dead silage bacteria, resulting in the increased FER concentrations. Neither microbial
growth nor cross reaction with a reagent can be implicated due to the addition of sodium azide
during the incubation and the use of high performance liquid chromatography to quantify FER,
respectively. In support of the latter suggestion, Sang et al. (2011) reported that 5 mM of Ca2+,
Fe3+, and Co2+ stimulated ferulic acid esterase activity from an unculturable soil bacteria, which
grew in silage contaminated with soil (Pahlow et al., 2003). Adding EFE 1A (179 μg/g), 2A (367
μg/g), 11C (211 μg/g) or 15D (242 μg/g) alone increased the concentration of FER relative to the
value for the Control (158 μg/g; P < 0.05) but 13D did not. Effects of COF addition to EFE on
release of FER depended on the EFE – COF combination (P < 0.001). For instance, adding Mn2+
to 1A increased the response but adding Mg2+ decreased it (194 and 162 μg/g, respectively; P <
0.05), adding Ca2+ or Co2+ to 2A increased the responses but adding Mg2+, Fe2+ or Mn2+
decreased them (414 and 393; 271, 305 and 334 μg/g, respectively; P < 0.05), adding Fe2+ and
Co2+ to 11C decreased the response (192 and 196 μg/g, respectively; P < 0.05) as did adding
Mg2+, Ca2+, Fe2+, Co2+, and Mn2+ to 15D (174, 222, 204, 196 and 225 μg/g, respectively; P <
0.05). No change in FER was detected when COF were added to 13D. Therefore, FER release
was only increased by adding Mn2+ to 1A or by adding Ca2+ or Co2+ to 2A. No published studies
that examined effects of adding Mn2+ to T. reesei ferulic acid esterase were found, but Kanauchi
et al. (2008) reported that Mn2+ inhibited ferulic acid esterase from A. awamori. The increase in
128
FER due to adding Ca2+ to 2A was 1.8 times greater than that detected by adding Ca2+ alone,
whereas that due to adding Co2+ was only 1.2 times greater than that due to adding Co2+ alone.
Thus, Ca2+ was more effective at synergistically increasing the release of FER.
Adding Mg2+, Fe2+, Co2+, or Mn2+ alone increased the COU concentration relative to that
for the Control (170, 168, 178, and 174 vs. 150 μg/g; P < 0.05) as did adding 2A, 11C and 15D
alone (239, 162, 210 vs. 150 μg/g, P < 0.05). However, effects of COF addition to EFE on
release of COU depended on the EFE - COF combination (P < 0.001). The concentration of
COU was decreased (P < 0.05) by adding Mg2+ to 1A (131 μg/g), Mg2+, Fe2+, Co2+, or Mn2+ to
2A (156, 184, 225, and 195 μg/g, respectively), Mg2+ to 13D (129 μg/g), or Mg2+, Ca2+, Fe2+,
Co2+, and Mn2+ to 15D (139, 160, 167, 159, and 176 μg/g, respectively). In contrast, COU
concentration was increased (P < 0.05) by adding Ca2+ to 2A (256 μg/g), Mg2+ or Ca2+ to 11C
(176 and 174 μg/g; P < 0.05), or Ca2+, Fe2+, Co2+, or Mn2+ to 13D (183, 163, and 162 μg/g,
respectively). Hence when added to EFE, Ca2+ synergistically increased release of COU from the
cell wall more consistently than other COF. Yet, McCrae et al. (1994) reported no changes in A.
awamori p-coumaroyl esterase activity when 20 mM of Ca2+ was added perhaps because of their
high dose and the different EFE source.
Experiment 2: Effects of Cofactor Addition to EFE on Digestibility and Fermentation
Manganese addition to EFE 11C.
Since both HEM (NDF-ADF) and CEL (ADF-ADL) concentrations are calculated by
difference, changes in ADFD or ADLD will influence the HEMD and CELD values,
respectively. This needs to be borne in mind when interpreting HEMD and CELD results.
Adding Mn2+ to 11C gave one of the greatest increases in saccharification in Experiment
1, hence effects of this EFE – COF combination on digestibility and fermentation were explored
in Experiment 2. Adding EFE 11C alone increased DMD, NDFD, HEMD, ADFD, and CELD
129
(Table 5-3; P < 0.001). Increasing the rate of Mn2+ application with or without EFE 11C resulted
in linear (P < 0.01) increases in DMD and NDFD, but the increments were greater when 11C
was present (EFE × DOSE interaction; P < 0.05). Adding 10 mM of Mn2+ to 11C resulted in a
3.5% increase in DMD beyond the 6.6% increase caused by adding 11C alone and a 7.3 %
increase beyond the 2.7% increase caused by adding 10 mM of Mn2+ alone. Likewise, adding 10
mM of Mn2+ to 11C resulted in a 8.1% increase in NDFD beyond the 15.5% increase caused by
adding 11C alone and a 17.2 % increase beyond the 6.3% increase caused by adding 10 mM of
Mn2+ alone. Therefore, adding Mn synergistically increased the digestibility of bermudagrass
haylage by EFE 11C.The increases in DMD and NDFD due to adding 0 vs. 10 mM of Mn2+ are
attributable to corresponding increases in ADFD (38.3 vs. 42.9%, P < 0.001) and CELD (43.6
vs. 48.7%, P < 0.001), yet HEMD was unaffected by Mn2+ dose. These results indicate that
cellulolytic activities were stimulated by adding Mn2+ but xylanolytic activities were not.
Manganese increases the activity of exoglucanases, endoglucanases, and β-glucosidases (Singh
et al., 1990; Riou et al., 1998; Saad and Fawzi, 2004), which are all involved in cellulose
depolymerization. In an early study, omission of Mn from an in vitro ruminal-fluid medium
reduced cellulose digestibility (Chamberlain and Burroughs, 1962). In agreement, Arelovich et
al. (2000) reported an increase in in vitro DMD of prairie hay due to adding up to 100 mg/L of
Mn2+ (chloride salt) to ruminal fluid (44.7 vs. 42.0%). Martinez and Church (1970) also reported
that adding over 100 mg/L of Mn2+ (sulfate salt) reduced cellulose digestion by washed
suspensions of rumen microbes but adding 5 to 30 mg/L of Mn2+ optimized the digestion. In this
study, the Mn concentrations in the buffered-ruminal incubation fluid were 3.16, 3.37, 5.20 and
23.5 mg/L for Mn2+ doses of 0, 0.1, 1 and 10 mM in the EFE 11C solution, respectively.
Therefore, the 1 and 10 mM doses were within the range reported to stimulate digestion by
130
ruminal bacteria in the previous study. This may explain why adding 10 mM of Mn2+ to 11C
synergistically increased NDFD in this study.
Applying 11C alone increased concentrations of acetate, butyrate and decreased the
acetate to propionate ratio (A:P; P < 0.01; Table 5-4). An EFE × DOSE interaction (P = 0.002)
was detected for total VFA (TVFA). Increasing the Mn2+ dose in the absence of 11C resulted in a
cubic TVFA response (P < 0.01), in which only the 0.1 mM dose increased TVFA concentration
(4.31%; P < 0.05). However, when EFE 11C was present, TVFA concentration increased
linearly (P < 0.01) with increasing Mn2+ dose. Biswas et al. (2012) and Durand and Kawashima
(1980) also reported that TVFA concentration was increased by adding Mn to the ruminal
incubation fluid used to ferment rice straw and prairie hay-based diets. In this study, the 10 mM
Mn2+ dose increased (P < 0.05) TVFA concentration by 6.1% relative to the value achieved by
adding 11C alone in agreement with corresponding synergistic increases in DMD and NDFD.
Therefore, adding 10 mM of Mn2+ to 11C would likely synergistically increase energy supply to
cows because they derive about 70% of their energy requirements from VFA (Bergman. 1990).
No effect of Mn2+ dose or interaction between 11C and Mn2+ was detected for acetate or
butyrate concentration but increasing the Mn2+ dose had cubic and linear effects on propionate
concentration in the absence or presence of 11C, respectively (EFE × DOSE interaction; P =
0.008). Only the 0.1 mM Mn2+ dose increased propionate concentration in the absence of 11C
(7.35%; P < 0.05). Adding 10 mM of Mn2+ with 11C increased propionate concentration (P <
0.05) by 7.44% beyond the increase due to adding 11C alone. Manganese has stimulated CO2
fixation during succinate production by Ruminococcus flavefaciens (Durand and Kawashima,
1980). This could explain the positive effects of Mn on propionate concentration since succinate
is a precursor of propionate (White et al., 2012).
131
Effects of 11C on the A:P ratio were influenced by adding Mn2+ (P < 0.02). In the
absence of 11C, no Mn2+ dose effects on A:P ratio were evident but in the presence of 11C,
increasing the Mn2+ dose linearly decreased the A:P ratio (P < 0.05). Therefore, adding Mn to
11C would likely increase the beneficial effects of 11C on ruminal energy supply (from
gluconeogenic substrates) and the efficiency of energy utilization in the rumen. These factors
will likely result in increased milk production and reduced ketosis problems in cows
(VandeHaar, 2005) if these in vitro responses occur in cows.
Iron addition to EFE 2A or 13D.
Adding Fe2+ to EFE 2A or 13D resulted in some of the greatest increases in
saccharification in Experiment 1, hence effects of these EFE – COF combination on digestibility
and fermentation were explored in Experiment 2. Adding EFE 2A or 13D alone increased DMD,
NDFD, ADFD, and CELD and 2A also increased HEMD (Tables 5-5 and 5-6; E, P < 0.001).
Increasing the rate of Fe2+ application linearly decreased (P < 0.01) all digestibility measures in
the absence of 2A but had no effect in the presence of 2A (EFE × DOSE interaction; P < 0.05).
Therefore, 2A retained its hydrolytic effect in the presence of Fe2+ and prevented adverse effects
of adding 10 mM of Fe2+alone on DMD (- 6.03%), NDFD (- 14.41%), and HEMD (- 14.24%),
This suggests that certain EFE can be used to reduce adverse effects of Fe toxicity on the
digestion of feeds. More research is required to understand and exploit this response.
Increasing the rate of Fe2+ application linearly decreased (P < 0.01) DMD, NDFD, ADFD
and CELD in the presence or absence of 13D but only decreased HEMD in the absence of the
EFE. Therefore, adding 10 mM of Fe2+ with 13D prevented the beneficial effects of adding 13D
alone on DMD and NDFD (EFE × DOSE interaction; P < 0.05). Harrison et al. (1992) found that
adding 100 mg/L of Fe2+ (chloride salt) to the ruminal incubation fluid reduced (- 10.18%)the
DMD of a fescue-based TMR compared to adding no Fe2+.However, Martinez and Church
132
(1970) reported that adding up to 50 mg/L of Fe3+(chloride salt) had no toxic or negative effects
on cellulose digestion by washed suspensions of rumen microbes and adding 2 to 5 mg/L of Fe3+
increased the response. Durand and Kawashima (1980) also recommended adding 1 to 10 mg/L
of Fe to ruminal fluid to optimize microbial metabolism. In this study, adding 0, 0.1, 1, and 10
mM of Fe2+ to 2A or 13D resulted in mean Fe concentrations in the incubation fluid of 1.86,
2.07, 3.93 and 22.55 mg/L, respectively. Therefore, the results of this study seem to indicate that
a lower dose (22.5 mg/L) than that indicated by Martinez and Church (50 mg/L) can cause
toxicity in ruminal in vitro incubations. Different toxicity outcomes among the latter study and
this one are most likely because washed microbe suspensions were used in the latter study
instead of the whole rumen fluid used in the current experiment. However, the lower Fe doses
used in this study did not cause any digestibility depression and they were within the optimal
range recommended for optimizing microbial metabolism by Durand and Kawashima (1980).
No EFE × DOSE interaction was detected (P > 0.05) for ADFD or CELD due to adding
Fe2+ to 2A or 13D. Yet, unlike lower doses, adding 10 mM of Fe2+ with or without EFE 2A
decreased ADFD (- 5.0%) and CELD (- 7.6%). Similarly, ADFD and CELD were decreased by
adding 10 mM of Fe2+ with or without 13D. Therefore, it seems that EFE 2A and 13D were more
effective at preventing adverse effects of Fe2+ addition on digestion of hemicellulose than on
cellulose digestion. Riou et al. (1998) reported that a β-glucosidase from A. oryzae was inhibited
strongly by 5 mM of Fe3+, which could partly explain the inhibition of cellulose digestion by
high concentrations of Fe in this study. In general, 13D was more susceptible than EFE 2A to the
deleterious effects on digestibility of high Fe concentrations.
Adding increasing doses of Fe2+ alone resulted in nonlinear effects on all fermentation
measures whereas, adding 2A alone increased TVFA, acetate, propionate, butyrate and decreased
133
the A:P ratio (P < 0.01; Table 5-7). Adding as little as 0.1 mM of Fe2+ to 2A prevented the
increase in TVFA caused by treatment with 2A alone, but no further decrease (cubic, P < 0.01)
was detected as the Fe2+ dose increased (EFE × DOSE interaction; P = 0.004). Harrison et al.
(1992) reported that adding 100, 200, and 500 mg/L of Fe2+ increased TVFA concentration but
no further increase occurred when 1000 mg/L were added. However, in that experiment, ferrous
sulfate was used rather than the ferrous chloride used in this study. The form of the iron likely
determines the outcome since it affects its solubility. Sulfate in ferrous sulfate would be rapidly
metabolized to sulfide in the rumen, forming a black precipitate, which complexes with Fe and
makes it less soluble and accessible to rumen microbes, whereas, ferrous chloride is more soluble
and consequently more toxic (Harrison et al., 1992).
No EFE × DOSE interaction was detected for acetate when Fe2+ was added to 2A, but
such interactions were evident for propionate, butyrate, and A:P ratio (EFE × DOSE interaction;
P < 0.001). Increasing the dose of Fe2+ reduced acetate concentration in the presence or absence
of 2A though the decrease was more pronounced when 2A was absent. Increasing the Fe2+ dose
also resulted in nonlinear responses in propionate, butyrate and the A:P ratio in the presence of
2A (P < 0.01).Adding increasing doses of Fe2+ alone resulted in nonlinear effects on all
fermentation measures (Table 5-8) but adding 13D alone did not affect any of the fermentation
measures. Adding increasing doses of Fe2+ to 13D did not affect TVFA (EFE × DOSE
interaction; P < 0.01)but linearly decreased acetate and butyrate concentration and A:P ratio
(EFE × DOSE interaction; P < 0.05). Adding 10 mM of Fe2+ to 13D increased propionate
concentration but lower doses did not (EFE × DOSE interaction; P < 0.01). In summary, adding
Fe2+ to 2A or 13D is not recommended because though they increased saccharification, they did
not influence or mostly decreased beneficial effects of the EFE on measures of digestibility and
134
fermentation in this study. Even the slight increase in propionate concentration and the decrease
in A:P ratio due to adding 10 mM of Fe2+ to 13D are likely outweighed by corresponding
decreases in DMD, NDFD, CELD and ADFD.
Conclusions
Adding 1 mM of each of the COF to EFE 2A or 11C synergistically increased
saccharification of BH as did adding1 mM of Fe2+ to 1A, Mn2+, Co2+, and Fe2+ to 13D, or Co2+or
Fe2 to 15D. The greatest saccharification responses were obtained by adding Mn2+ to11C (38%)
or by adding Fe2+ to 2A or 13D (10 and 21.9%, respectively). Effects of adding increasing doses
of these COF on EFE-mediated changes in vitro digestibility depended on the COF and EFE.
Adding 10 mM of Mn2+ to EFE 11C resulted in a 8.1% increase in NDFD beyond the 15.5%
increase caused by adding EFE 11C alone and a 17.2% increase beyond the 6.3% increase
caused by adding 10 mM of Mn2+ alone. Whereas adding Fe2+ to 2A had no effects on EFE-
mediated digestibility responses, but 2A prevented adverse effects of adding Fe2+ alone on DMD
and NDFD. In contrast, adding Fe2+ to 13D reduced the increases in NDFD caused by adding the
EFE alone. This study shows that adding COF to EFE can synergistically increase, decrease or
not affect the hydrolytic effects of EFE on forage cell walls depending on the specific EFE –
COF combination. More work is required to understand the mechanisms resulting in these
outcomes in order to exploit beneficial effects of COF on EFE.
135
Table 5-1. Endoglucanase, xylanase, exoglucanase, β-glucosidase (μmol of sugar released/min/g) and ferulic acid esterase (nmol of
ferulic acid released/min/g) activities, protein concentration (mg/g), form, application rate (g/kg of bermudagrass DM),
mineral concentration (mg/L of EFE), and biological source of exogenous fibrolytic enzyme (EFE) preparations used.
Parameter EFE
1A 2A 11C 13D 15D S.D.
Endoglucanase 1,693 3,624 1,506 286 70 112
Xylanase 1,276 29,301 1,703 86 6,499 221
Exoglucanase 1.68 0.84 0.97 0.29 0.29 0.10
β-glucosidase 10.1 11.7 12.7 1.9 0.1 0.9
Ferulic acid
esterase
2.18 1.46 6.30 2.35 2.57 1.27
Protein 65.3 111.1 81.1 18 28.3 25.0
Form Liquid Liquid Liquid Liquid Liquid
Application Rate 4.66 4.66 10.4 7.8 7.8
Ca 90.6 108.4 83.1 139.3 91.0
Mg 62.1 94.2 42.6 28.4 24.8
Fe 4.02 4.66 1.40 0.30 0.48
Mn 0.65 0.42 0.31 0.07 0.07
Co 0.033 0.028 0.024 0.026 0.018
Biological Source Trichoderma
reesei T. reesei
T. reesei Aspergillus
oryzae
A. oryzae
136
Table 5-2. Effects of adding cofactors (COF) to exogenous fibrolytic enzymes (EFE) on DM loss
(%), concentrations (% of DM) of NDF, hemicellulose (HEM), water-soluble
carbohydrates (WSC), and ferulic (FER) and p-coumaric acids (COU; μg/g) of a 4-wk
regrowth of bermudagrass haylage (Experiment 1).a
Treatment WSC NDF HEM ADF DM Loss FER COU
No EFE
No COF 1.90ab, z 71.1a, z 35.4a, z 35.7bc, z 17.3ab, z 158a, z 150a, y
Mn2+ 2.06b 70.4a 35.4a 35.0a 17.2a 170ab 174b
Co2+ 2.06b 71.0a 35.3a 35.7c 17.0a 180b 178b
Fe2+ 1.95ab 70.0a 34.9a 35.1ab 17.8b 174b 168b
Ca2+ 1.99b 70.3a 34.7a 35.6ab 17.2a 183b 152a
Mg2+ 1.80a 70.9a 35.2a 35.6bc 17.6ab 158a 170b
SEM 0.059 0.40 0.34 0.20 0.21 5.2 3.8
EFE 1A
1A 3.73ab, w 68.3a, x 33.5a, x 34.8ab, y 18.4bc, y 179b, y 159bc, xy
1A + Mn2+ 3.77abc 67.8a 33.4a 34.4a 18.3bc 194c 165bc
1A + Co2+ 3.88bc 68.5a 33.6a 34.9ab 18.6cd 182bc 166c
1A + Fe2+ 3.91c 67.3a 32.8a 34.5a 19.1de 177b 156bc
1A + Ca2+ 3.72ab 67.1a 32.6a 34.5a 19.3e 180b 155b
1A + Mg2+ 3.67a 68.6a 33.4a 35.2b 18.0abc 162a 131a
SEM 0.058 0.40 0.34 0.20 0.21 5.2 3.8
EFE 2A
2A 6.43a, v 64.9a, w 29.8a, w 35.0ab, y 20.0a, x 367d, v 239d, v
2A + Mn2+ 6.77c 64.9a 29.6a 35.3ab 20.3a 334c 195b
2A + Co2+ 6.80c 63.5a 28.8a 34.7a 21.5b 393e 225c
2A + Fe2+ 7.05d 64.7a 29.0a 35.0ab 21.1b 305b 184b
2A + Ca2+ 6.60b 64.3a 28.8a 35.5b 20.6ab 414f 256e
2A + Mg2+ 6.64bc 64.0a 28.7a 35.3b 21.1b 271a 156a
SEM 0.058 0.40 0.34 0.20 0.21 5.2 3.8
a, b, c, d, e Within EFE treatments, means in a column with different superscripts differed (P <
0.05). w, x, y, z For EFE to which no COF were added, means in a column with different
superscripts differed (P < 0.05)
137
Table 5-2. Continued a
Treatment WSC NDF HEM ADF DM Loss FER COU
EFE 11C
11C 3.49a, x 68.0a, x 33.0a, x 35.0ab, y 19.4c, x 211b, x 162ab, x
11C + Mn2+ 4.83d 67.8a 32.9a 34.9a 18.6b 205ab 170bc
11C + Co2+ 4.30c 68.2a 33.5a 34.7a 18.6b 196a 163ab
11C + Fe2+ 4.31c 68.8a 33.6a 35.2ab 18.5b 192a 156a
11C + Ca2+ 4.00b 67.6a 32.9a 34.7a 18.7b 198ab 174c
11C + Mg2+ 3.95b 68.9a 33.3a 35.6b 17.8a 211b 176c
SEM 0.058 0.40 0.34 0.20 0.21 5.2 3.8
EFE 13D
13D 1.92a, z 70.9a, z 35.4a, z 35.5ab, z 17.4abc, z 150a, z 140b, z
13D + Mn2+ 2.17b 70.2a 34.8a 35.4ab 17.3abc 162a 161c
13D + Co2+ 2.19bc 71.2a 35.5a 35.7b 16.9a 154a 162c
13D + Fe2+ 2.34c 69.9a 34.6a 35.4ab 17.4abc 152a 163c
13D + Ca2+ 2.01ab 70.3a 35.3a 35.0a 17.4abc 148a 183d
13D + Mg2+ 1.94a 69.7a 34.4a 35.3ab 17.6bc 160a 129a
SEM 0.06 0.40 0.34 0.20 0.21 5.2 3.8
EFE 15D
15D 2.57ab, y 69.5a, y 33.8a, y 35.7ab, z 17.4abc, z 242d, w 210e, w
15D + Mn2+ 2.70bc 69.8a 34.6a 35.2a 17.5abc 225c 176d
15D + Co2+ 2.76cd 69.6a 33.8a 35.8b 17.9cd 196b 159b
15D + Fe2+ 2.92d 69.6a 34.0a 35.6ab 17.9cd 204b 167cd
15D + Ca2+ 2.62abc 69.4a 33.6a 35.8ab 18.3d 222c 160bc
15D + Mg2+ 2.51a 69.8a 34.4a 35.4ab 18.0cd 174a 139a
SEM 0.06 0.40 0.34 0.02 0.21 5.3 3.8
a, b, c, d, e Within EFE treatments, means in a column with different superscripts differed (P <
0.05). w, x, y, z For EFE to which no COF were added, means in a column with different
superscripts differed (P < 0.05)
138
Table 5-3. Effects of treating 4-wk regrowth bermudagrass haylage with exogenous fibrolytic
enzyme (EFE) 11C with or without increasing doses of Mn2+ on in vitro digestibility
of DM (DMD), NDF (NDFD), hemicellulose (HEMD), ADF (ADFD) and cellulose
(CELD) and ADL disappearance (ADLD)(Experiment 2).a
Mn2+dose No EFE EFE P value
(mM) EFE DOSE EFE × DOSE
DMD (%)
0 48.1a 51.3d < 0.001 < 0.001 0.047
0.1 48.8ab 50.7cd
1 48.7ab 50.7cd
10 49.4b 53.0e
SEM 0.34 0.34
Contrast1 L** L**
NDFD (%)
0 35.4a 40.8d < 0.001 < 0.001 0.048
0.1 36.6ab 39.9cd
1 36.3ab 39.9cd
10 37.6b 43.7e
SEM 0.58 0.58
Contrast L** L**
HEMD (%)
0 35.1a 40.8b < 0.001 0.231 0.15
0.1 35.2a 39.1b
1 35.0a 39.5b
10 34.9a 42.1b
SEM 0.74 0.80
Contrast None None
1 Linear (L), quadratic (Q), cubic (C); None (P >0.1); * (P < 0.05); ** (P < 0.01);
Means with different superscripts differed (P < 0.05).
139
Table 5-3. Continued 1
Mn2+dose No EFE EFE P value
(mM) EFE DOSE EFE × DOSE
ADFD (%)
0 35.6a 40.9c < 0.001 < 0.001 0.223
0.1 38.0a 40.6c
1 37.7a 41.0c
10 40.4b 45.4d
SEM 0.73 0.73
Contrast None None
CELD (%)
0 41.9a 45.3c < 0.001 < 0.001 0.431
0.1 42.9a 46.2c
1 42.5a 47.4c
10 46.2b 51.3d
SEM 0.71 0.77
Contrast None None
ADLD (%)
0 -19.5a 2.2e 0.271 0.866 < 0.001
0.1 -4.8cde -8.2cd
1 -3.9de -14.9ab
10 -10.6abc -6.5cde
SEM 3.5 3.8
Contrast C** None
1 Linear (L), quadratic (Q), cubic (C); None (P >0.1); * (P < 0.05); ** (P < 0.01);
Means with different superscripts differed (P < 0.05).
140
Table 5-4. Effects of treating 4-wk regrowth bermudagrass haylage with exogenous fibrolytic
enzyme (EFE) 11C with or without increasing doses of Mn2+ on concentrations of
total volatile fatty acids (TVFA), acetate, propionate, and butyrate and acetate to
propionate ratio (A:P) (Experiment 2).1
Mn2+dose No EFE EFE P-value
(mM) EFE DOSE EFE × DOSE
TVFA (mM)
0 60.3ab 61.4abc 0.0025 0.0591 0.0019
0.1 62.9c 62.0abc
1 60.9ab 62.3bc
10 60.2a 65.1d
SEM 0.72 0.77
Contrast1 C** L**
Acetate(mM)
0 36.0a 37.0b 0.0144 0.8468 0.7475
0.1 36.3a 37.0b
1 36.2a 36.7b
10 36.0a 37.7b
SEM 0.59 0.55b
Contrast None None
Propionate(mM)
0 10.8a 11.2ab 0.0086 0.157 0.0079
0.1 11.5bc 11.2ab
1 10.9ab 11.3ab
10 10.8a 12.0c
SEM 0.22 0.22
Contrast C** L*
Butyrate(mM)
0 5.27a 5.63b 0.004 0.7258 0.1509
0.1 5.46a 5.51b
1 5.53a 5.58b
10 5.34a 5.71b
SEM 0.103 0.095
Contrast None None
A : P
0 3.35c 3.21ab 0.0093 0.6125 0.0188
0.1 3.25abc 3.32bc
1 3.31bc 3.24abc
10 3.32c 3.15a
SEM 0.043 0.040
Contrast None L* 1 Linear (L), quadratic (Q), cubic (C); None (P > 0.1); * (P < 0.05); ** (P < 0.01);
Means with different superscripts differed (P < 0.05).
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Table 5-5. Effects of treating 4-wk regrowth bermudagrass haylage with exogenous fibrolytic
enzyme (EFE) 2A with or without increasing doses of Fe2+ on in vitro digestibility of
DM (DMD), NDF (NDFD), hemicellulose (HEMD), ADF (ADFD) and cellulose
(CELD) and ADL disappearance (ADLD) (Experiment 2).a
Fe2+dose No EFE EFE P-value
(mM) EFE DOSE EFE × DOSE
DMD (%)
0 48.1b 50.6c < 0.001 < 0.001 0.004
0.1 48.4b 50.8c
1 48.0b 50.5c
10 45.2a 50.2c
SEM 0.43 0.40
Contrast L** None
NDFD (%)
0 35.4b 39.7c < 0.001 < 0.001 0.004
0.1 35.9b 40.0c
1 35.1b 39.4c
10 30.3a 39.0c
SEM 0.74 0.68
Contrast L** None
HEMD (%)
0 35.1b 39.6c < 0.001 < 0.001 0.007
0.1 34.6b 39.3c
1 33.6b 38.3c
10 30.1a 38.7c
SEM 0.66 0.66
Contrast L** None
1 Linear (L), quadratic (Q), cubic (C); None (P > 0.1); * (P < 0.05); ** (P < 0.01);
Means with different superscripts differed (P < 0.05).
142
Table 5-5. Continued a
Fe2+dose No EFE P-value
(mM) EFE EFE DOSE
ADFD (%)
0 35.6b 40.6d < 0.001 0.009 0.232
0.1 36.2b 40.8d
1 36.6b 41.2d
10 32.4a 39.9c
SEM 0.87 0.87
Contrast L** None
CELD (%)
0 41.9b 44.9d < 0.001 0.001 0.117
0.1 40.9b 45.9d
1 40.7b 45.0d
10 36.5a 43.7c
SEM 0.92 0.92
Contrast L** None
ADLD (%)
0 -19.5a 3.0bc < 0.001 0.005 0.010
0.1 -5.0b -3.9b
1 0.7b 2.4bc
10 -3.0b 11.2c
SEM 3.7 3.7
Contrast C* L*
1 Linear (L), quadratic (Q), cubic (C); None (P > 0.1); * (P < 0.05); ** (P < 0.01);
Means with different superscripts differed (P < 0.05).
143
Table 5-6. Effects of treating 4-wk regrowth bermudagrass haylage with exogenous fibrolytic
enzyme (EFE) 13D with or without increasing doses of Fe2+ on in vitro digestibility
of DM (DMD), NDF (NDFD), hemicellulose (HEMD), ADF (ADFD) and cellulose
(CELD) and ADL disappearance (ADLD) (Experiment 2).a
Fe2+dose No EFE EFE P-value
(mM) EFE DOSE EFE × DOSE
DMD (%)
0 48.1b 49.8d < 0.001 < 0.001 0.043
0.1 48.4bc 49.7d
1 48.0b 49.3cd
10 45.2a 48.4bc
SEM 0.40 0.40
Contrast1 L** L**
NDFD (%)
0 35.4b 38.3d < 0.001 < 0.001 0.047
0.1 35.9bc 38.1d
1 35.1b 37.4cd
10 30.3a 35.9bc
SEM 0.69 0.69
Contrast L** L**
HEMD (%)
0 35.1bcd 37.0d < 0.001 < 0.001 0.014
0.1 34.6bc 37.3d
1 33.6b 36.8d
10 30.1a 36.3cd
SEM 0.72 0.66
Contrast L** None
1 Linear (L), quadratic (Q), cubic (C); None (P > 0.1); * (P < 0.05); ** (P < 0.01);
Means with different superscripts differed (P < 0.05).
144
Table 5-6. Continued a
Fe2+dose No EFE P-value
(mM) EFE EFE DOSE
ADFD (%)
0 35.6b 39.7d < 0.001 < 0.001 0.077
0.1 36.2b 39.0d
1 36.6b 36.7d
10 32.4a 36.3c
SEM 0.88 0.88
Contrast L** Q*
CELD (%)
0 41.9b 44.4d < 0.001 < 0.001 0.710
0.1 40.9b 45.2d
1 40.7b 43.4d
10 36.5a 39.7c
SEM 0.89 0.89
Contrast L** L**
ADLD (%)
0 -19.5a -0.7c 0.655 0.229 0.001
0.1 -5.0bc -15.2ab
1 0.7c -15.3ab
10 -3.0bc -1.3c
SEM 4.7 4.7
Contrast C* None
1 Linear (L), quadratic (Q), cubic (C); None (P > 0.1); * (P < 0.05); ** (P < 0.01);
Means with different superscripts differed (P < 0.05).
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Table 5-7. Effects of treating 4-wk regrowth bermudagrass haylage with exogenous fibrolytic
enzyme (EFE) 2A with or without increasing doses of Fe2+ on concentrations of total
volatile fatty acids (TVFA), acetate, propionate, and butyrate and acetate to
propionate ratio (A : P) (Experiment 2).a
Fe2+dose No EFE EFE P-value
(mM) EFE DOSE EFE × DOSE
TVFA (mM)
0 60.3bc 64.9d < 0.001 < 0.001 0.004
0.1 57.5b 60.9c
1 59.1bc 59.3bc
10 50.8a 59.1bc
SEM 1.10 1.10
Contrast1 C* C*
Acetate(mM)
0 36.0c 37.6f 0.002 < 0.001 0.316
0.1 34.3b 35.6e
1 34.8b 35.2e
10 28.7a 31.5d
SEM 0.69 0.64
Contrast C* L**
Propionate(mM)
0 10.8c 12.2d < 0.001 < 0.001 <.001
0.1 10.1ab 10.8c
1 10.7bc 10.4bc
10 9.4a 11.8d
SEM 0.24 0.24
Contrast C* C**
Butyrate(mM)
0 5.27a 6.06d < 0.001 < 0.001 <.001
0.1 5.15a 5.55b
1 5.55b 5.59b
10 4.66c 5.50b
SEM 0.046 0.046
Contrast C** C**
A : P
0 3.35ef 3.16c < 0.001 < 0.001 <.001
0.1 3.40f 3.30de
1 3.25d 3.36ef
10 3.02b 2.79a
SEM 0.033 0.031
Contrast Q* C* 1 Linear (L), quadratic (Q), cubic (C); None (P > 0.1); * (P < 0.05); ** (P < 0.01);
Means with different superscripts differed (P < 0.05).
146
Table 5-8. Effects of treating 4-wk regrowth bermudagrass haylage with exogenous fibrolytic
enzyme (EFE) 13D with or without increasing doses of Fe2+ on concentrations of
total volatile fatty acids (TVFA), acetate, propionate, and butyrate and acetate to
propionate ratio (A : P) (Experiment 2).a
Fe2+dose No EFE EFE P-value
(mM) EFE DOSE EFE × DOSE
TVFA (mM)
0 60.3bc 61.0c < 0.001 < 0.001 0.003
0.1 57.5b 60.8c
1 59.1bc 61.7c
10 50.8a 59.5bc
SEM 1.14 1.05
Contrast1 C* None
Acetate(mM)
0 36.0d 35.6cd < 0.001 < 0.001 0.005
0.1 34.3c 35.5cd
1 34.8cd 35.8cd
10 28.7a 32.3b
SEM 0.58 0.53
Contrast C* L**
Propionate(mM)
0 10.8bc 10.8c < 0.001 0.29 0.001
0.1 10.1ab 11.1cd
1 10.7bc 11.3cd
10 9.5a 11.6d
SEM 0.26 0.26
Contrast C* None
Butyrate(mM)
0 5.27bc 5.41cd < 0.001 < 0.001 < 0.001
0.1 5.15b 5.43cde
1 5.55de 5.32bc
10 4.66a 5.60e
SEM 0.061 0.068
Contrast C** L*
A:P ratio
0 3.35cd 3.28c < 0.001 < 0.001 0.038
0.1 3.40d 3.28c
1 3.25c 3.25c
10 3.02b 2.78a
SEM 0.043 0.039
Contrast Q* L** 1 Linear (L), quadratic (Q), cubic (C); None (P > 0.1); * (P < 0.05); ** (P < 0.01);
Means with different superscripts differed (P < 0.05).
147
CHAPTER 6
IMPROVING FORAGE DIGESTION AND DAIRY COW PERFORMANCE WITH
FIBROLYTIC ENZYMES
Background
Several studies have examined the efficacy of using exogenous fibrolytic enzymes (EFE
to improve forage quality and ruminant animal performance but the results have been equivocal.
(Adesogan, 2005). Supplementing EFE to nonruminant livestock can improve feed efficiency
and diet flexibility (Bedford and Partridge, 2010), yet EFE use in ruminant diets has been limited
due to inconsistent animal performance responses, which are due partly to the wide array of
conditions under which they are tested and the limited understanding of their mode of action
(Beauchemin and Holtshausen, 2010). In most cases, enzymatic activities supplied by EFE are
not novel to the rumen and therefore EFE act on the same plant cell wall targets as endogenous
ruminal enzymes (Wang and McAllister, 2002). This partly explains why relative to their effects
on dairy cows fed for maintenance, EFE have been more effective at improving the productivity
of high-producing lactating dairy cattle. Such cows typically have depressed ruminal fiber
digestion (Beauchemin and Holtshausen, 2010) due to factors like low ruminal pH and high total
tract rate of passage (Mouriño et al., 2001; Cochran et al., 2007). Hence, dietary addition of EFE
is more likely to result in increased performance by such cattle compared with those fed at
maintenance.
In tropical and subtropical regions, the high fiber content and low digestibility of warm-
season grasses limit dairy cow productivity (Hanna and Sollenberger, 2007), making such
forages and the diets on which they are based good candidates for improvement by EFE. Arriola
et al. (2011b) reported that adding an EFE to a corn silage and alfalfa hay- based total mixed
ration (TMR) for dairy cows increased fiber digestibility by 6%, and increased feed efficiency by
about 16%. However, when the same EFE (Queiroz et al., 2011) and another one (Bernard et al.,
148
2010) were applied to a bermudagrass silage-based TMR, none of the performance measures was
increased. Consequently, research was needed to increase the effectiveness of using EFE to
digest bermudagrass and to improve the performance of cows fed bermudagrass-based diets. A
series of experiments were conducted to screen 18 EFE from 5 companies, select the 5 that gave
the greatest increases in the NDFD of bermudagrass haylage, optimize the dose of the latter,
examine if addition of cofactors synergistically increased the NDFD response, and select the
most promising and economic EFE × dose × cofactor treatment combination for testing in a dairy
cow study. The objective of this experiment was to evaluate the effects of adding the most
promising EFE identified in the previous experiments (2A) and the EFE that increased feed
efficiency in the study of Arriola et al. (2011b; 3A) to the diet on DMI, ruminal fermentation,
kinetics of ruminal digestion, and performance of lactating dairy cattle. The hypothesis was that
application of EFE 2A to the TMR will improve intake, milk yield, kinetics of digestion, and
fermentation in the rumen, whereas applying 3A will improve feed efficiency.
Materials and Methods
Location, Housing and Weather
The study was conducted at the University of Florida Dairy Unit (Hague, FL) from
February to August 2013 (26 wk). Cows were housed in a free-stall open-sided barn fitted with
two rows of fans (1 fan/6 linear meters) for cooling (one facing the feed lane immediately above
the feed bunk and the other immediately above the free stalls). Fans were equipped with low-
pressure water nozzles and both fans and nozzles were activated once ambient temperature
reached 21.1oC. Suspended fluorescent lights provided lighting. During the experiment, the
mean temperature and relative humidity were 21.4oC and 80.9% with minima of 3.2oC and
49.0% and maxima of 28.3oC and 97.0 %, respectively (FAWN, 2013). Stalls were
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approximately 1.14 x 2.31 x 1.21 m and were bedded with sand for alleviation of hoof and leg
stress. Manure in the areas between the feed bunks and the free stalls was flushed out twice daily
by an automated flushing system.
Animals and Treatments
The University of Florida Institute of Food and Agricultural Sciences Animal Research
Committee approved the protocols for this study. In Experiment 1, 66 lactating Holstein cows in
early lactation (21 ± 5 DIM) were grouped by previous milk production, parity (45 multiparous
and 21 primiparous), and previous milking frequency (2 vs. 4× daily) and randomly assigned to 1
of the following 3 treatments: 1) Control (CON, untreated), 2) Xylanase Plus (XYL, 1 mL/kg of
TMR DM, and 3) a 75:25 (v/v) mixture of Cellulase Plus and Xylanase Plus EFE (MIX, 3.4 mL /
kg of TMR DM). The MIX and XYL EFE were called EFE 3A and 2A, respectively in Chapters
3, 4, and 5 and they were sourced from non-recombinant Trichoderma reesei (Dyadic
International, Jupiter FL).
Cows were fed the TMR at 0700 and 1300 h using a separate, 250-kg feed capacity Calan
data ranger (American Calan Inc. Northwood, NH) for each treatment. The EFE were diluted in
deionized water (1:3 v/v) and sprayed with a garden hand-held sprayer on the TMR while it was
being mixed in the data ranger for 5 min. The TMR included 5.0% alfalfa-orchardgrass hay,
9.9% bermudagrass silage, and 35.1% corn silage (Table 6-1). Experimental diets were
formulated to meet the NRC (2001) guidelines for a dairy cow producing 40 kg of milk with
milk fat and protein concentrations of 3.5 and 2.85 %, respectively. Water was provided ad
libitum from an automatic watering system. Cows were fed ad libitum (115% of the previous
day’s intake) individually using Calan gates (American Calan Inc., Northwood, NH). Cows were
fed a common diet during the first 14 d of the experiment, and the last 11 d of this period were
considered the covariate period. Ingredient samples were taken twice a week during the
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experimental period to determine DM concentrations (48 h at 60oC) and the diets and amount of
enzyme applied were adjusted weekly based on changes in the DM concentration of the silages.
Enzymatic Activities
Table 6-2 lists all enzymatic activities and protein concentrations of the EFE.
Endoglucanase (E.C.3.2.1.4), exoglucanase (E.C. 3.2.1.91), xylanase (E.C. 3.2.1.8) and β-
glucosidase (E.C. 3.2.1.21) activities were quantified using carboxymethylcellulose, Avicel, oat-
spelt xylan and cellobiose as substrates (Wood and Bhat, 1988). Ferulic acid esterase (E.C.
3.1.1.73) activity was measured using ethyl ferulate as the substrate (Lai et al., 2009). All
activities were measured at a temperature of 39oC and a pH of 6 to mimic ruminal conditions.
Protein concentration was measured using the Bio-Rad Protein Assay with bovine serum
albumin as the standard (Bio-Rad Laboratories, Hercules, CA).
Sampling and Analysis
Cows were milked twice daily (0800 and 2000 h) and milk production was recorded.
Milk composition was analyzed by ‘in line’ AfiLab milk analyzers (S.A.E. Afikim, Kibbutz
Afikim, Israel) at each milking. Body weight (BW) was recorded twice daily after each milking
by the AfiFarm software system (S.A.E. Afikim). Body condition was scored (BCS) by the same
pair of observers on a 1 to 5 scale (Wildman et al., 1982) at the beginning of the trial and weekly
thereafter.
Duplicate samples of the corn silage, bermudagrass silage, alfalfa hay, concentrates and
orts were collected every 3 to 4 days, composited monthly, and dried at 60ºC for 48 h in a
forced-air oven. Dried residues were sampled bi-weekly composited monthly and ground to pass
the 1-mm screen of a Wiley mill (A.H. Thomas, Philadelphia, PA). Ground samples were
analyzed for DM (105ºC for 16 h) and ash (600ºC for 8 h) concentrations and those of NDF and
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ADF were measured sequentially using the method of Van Soest et al. (1991) in an ANKOM
200 Fiber Analyzer (Ankom Technologies, Macedon, NY). Heat-stable α-amylase was used in
the NDF assay with no sodium sulfite and the results were expressed inclusive of residual ash.
Hemicellulose was calculated as the difference between NDF and ADF. Nitrogen concentration
was determined by the Dumas combustion method (AOAC, 2000) using a Vario MAX CN
Macro Elemental Analyzer (Elementar Analysensysteme GmbH, Hanau, Germany) and CP was
calculated as N × 6.25. Blood samples were collected weekly by coccygeal venipuncture into
vacutainer tubes containing sodium heparin anticoagulant (BD Vacutainer, BD, Franklin Lakes,
NJ). The blood was centrifuged at 2,500 × g for 20 min at 4°C and the plasma was frozen
(−20°C). A Technicon autoanalyzer (Technicon Instruments Corp., Chauncey, NY) was used to
measure plasma glucose (Bran and Luebbe Industrial Method 339-19; Gochman and Schmitz,
1972) and BUN (Bran and Luebbe Industrial Method 339-01; Marsh et al., 1965).
Rumen Degradation Kinetics and Fermentation Measures
In Experiment 2, 3 ruminally-cannulated multiparous Holstein cows in mid-lactation (159
± 47 DIM and 735 ± 8 kg of BW) were assigned to the three treatments used in Experiment 1.
Experiment 2 had a 3 × 3 Latin square design with 23-d periods. Each period had 18 d for
adaptation to the diets followed by 3 d for measuring in situ ruminal degradability of the TMR, 1
rumen recovery day, and 1 d for measuring indices of ruminal fermentation. Cows were housed
under the same conditions described previously. The in situ DM degradability of the TMR was
determined in quadruplicate on d 19 to 21 of each period within cows consuming the same
treatment in that period. Ingredients sampled during each period were dried for 48 h at 60ºC,
ground to pass through the 4-mm screen of a Wiley mill and weighed (5 g of DM) into hot
weighed 10 × 20 cm ANKOM R1020 in situ bags (ANKOM Technology, Macedon, NY), which
were tied approximately 1 cm below the top end with rubber bands. A 2.6 mL solution
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containing nanopure water (CON) with or without the same dose of enzymes used in Experiment
1, was applied to the TMR 24 h before placing the bags in the rumen. Bags were kept at 25ºC
during this pre-incubation period. The bag pore size was 50 µm and the sample size to free bag
surface area ratio was 12.5 mg / cm2. Bags were attached to a rope with clips, placed in the
ventral sac of the rumen for 0, 4, 8, 16, 24, 48, and 72 h and removed simultaneously. Upon
removal from the rumen, bags were washed with cold water manually to remove adherent
particles and bacteria and then washed in a commercial washing machine (Kenmore, Benton
Harbor, MI) using a cool wash cycle without soap. Washed bags were dried for 48 h at 60ºC and
hot-weighed. The model of Mertens (1977) was fitted to the DM degradation data using the
NLIN procedure of SAS, version 9.4 (2013, SAS Inst., Inc., Cary, NC). The model is of the
form:
R(t) = Di × (e – kd × (t - L)) + Io
Where R(t) = Total undegraded residue at any time t (% of DM), Di = potentially
degradable fraction (% of DM), kd = fractional rate of degradation of Di (%/h), t = time incubated
in the rumen in h, L = discrete lag time in h, and Io = undegradable fraction after 72h of
incubation (% of DM). The washout fraction = 100 – Di - Io.
Samples of ruminal fluid were taken through the cannula from different locations in the
rumen by aspiration at 0900 h (just before feeding) and every other hour thereafter until 1900 h
on d 23, for a total of 6 samples per treatment. The samples were immediately filtered with 2
layers of cheesecloth and analyzed for pH using an Accumet XL25 calibrated pH meter (Fisher
Scientific, Pittsburgh, PA). Subsequently, the ruminal fluid (13 mL) was acidified with 130 µL
of a 9.0 M H2SO4 solution and then frozen at -40°C for further analysis. Thawed samples were
centrifuged at 8000 × g for 20 min at 4°C and the supernatant was analyzed for VFA and lactate
153
(Muck and Dickerson, 1988) using a Merck Hitachi Elite LaChrome High Performance Liquid
Chromatograph system (Hitachi, L2400, Tokyo, Japan) and a Bio-Rad Aminex HPX-87H
column (Bio-Rad Laboratories, Hercules, CA). Ammonia-N concentration was measured using
an adaptation of the Noel and Hambleton (1976) procedure that involved colorimetric N
quantification on a Technicon Auto Analyzer (Technicon, Tarrytown, NY).
Statistical Analysis
A completely randomized design was used to analyze the weekly data from Experiment
1. The milking frequency used prior to assignment to the current experiment and its interactions
were not significant for any dependent variable so they were removed from the final model.
The model used to analyze animal performance data was:
Yijk = µ + Ti + Pj + TPij + C(ij)k + Wl + TWil + PWjl + TPWijl + Eijkl
Where:
µ = general mean
Ti = effect of EFE i
Pj = effect of parity j
TPij = effect of the EFE i × parity j interaction
C(ij)k = random effect of cow k nested within treatment and parity (k = 1, 2, 3,…, n)
Wl = effect of week l
TWil = effect of the EFE i × week l interaction
PWjl = effect of the parity j × week l interaction
TPWijl = effect of the EFE i × parity j × week l interaction
Eijkl = experimental error
The GLIMMIX procedure of SAS version 9.4 (2013) was used to analyze the data. A
repeated measures statement with the autoregressive [ar(1)] covariance structure was used for the
154
analysis after examination of the Akaike, finite-population corrected Akaike, and Bayesian
information criteria for various covariance structures for each of the measured parameters. The
covariance structure with the least value for these criteria was chosen except when they had had
similar values, in which case, the simpler model was chosen (Littell et al., 2006). Milk
production and DMI during the last 11d of the training period were used as a covariate for
analyzing the milk production and intake data, respectively.
A Latin square design was used to analyze the data from Experiment 2 and the model
used to analyze ruminal fermentation data was:
Yijk = µ + Ti + Mj + TMij + Ck + Pl + Eijkl
Where:
µ = general mean
Ti = effect of EFE i
Mj = effect of time j
TMij = effect of the EFE i × time j interaction
Ck = random effect of cow k
Pl = effect of period l
Eijkl = experimental error
For the ruminal degradation kinetics, the effect of time (repeated) and interactions with
time were excluded from the model. The autoregressive [ar(1)] covariance structure was used for
all repeated measurements. Fisher’s F-protected Least Significance Difference test was used for
mean separation. Treatment significance was declared at P ≤ 0.05 and tendencies were declared
at 0.1 > P > 0.05.
155
Results and Discussion
Multiparous cows had greater (P < 0.05) voluntary intake of DM (DMI, 28.8 vs. 26.9
kg/d), OM (OMI, 26.8 vs. 25.0 kg/d), CP (CPI, 4.7 vs. 4.4 kg/d), NDF (NDFI, 9.1 vs. 8.5 kg/d),
HEM (HEMI, 4.3 vs. 4.0 kg/d), and ADF (ADFI, 4.8 vs. 4.5 kg/d) than primiparous cows but no
parity × treatment and parity × treatment × week interactions were detected (P > 0.1).
On average, the cows in this study had greater DMI (27.8 kg/d) than those reported in the
studies of Bernard et al. (2010; 24.5 kg/d), Queiroz et al. (2011; 22.0 kg/d), and Dean et al.
(2013; 21.3 kg/d), partly because of differences in the compositions of the diets and the BW and
lactation stages of the cows. Compared to feeding CON and MIX, feeding XYL increased mean
DMI (28.6 vs. 27.4 and 27.4 kg/d; P =0.048), OMI (26.7 vs. 25.5 and 25.5 kg/d; P = 0.031) and
CPI (4.7 vs. 4.5 and 4.5 kg/d; P = 0.021), respectively but feeding MIX did not affect (P > 0.10)
any measure of intake (Table 6-3). Arriola et al. (2011b) reported that applying the same MIX
EFE to diets containing 33 or 48% concentrate had no effect on DMI, CPI, NDFI, and ADFI by
cows in early lactation cows. Feeding XYL instead of MIX may have increased DMI in this
study because of its greater xylanase activity and the relatively high HEM concentration of the
TMR relative to those in the study of Arriola et al (2011b). The XYL EFE (2A) also had
degraded HEM effectively in previous experiments (Chapter 2). Greater DMI are especially
beneficial for dairy cows because milk production is limited by intake of digestible energy
(Beauchemin and Holtshausen, 2010). Dietary addition of EFE also has increased the DMI of
corn silage - (Gado et al., 2009) and barley-alfalfa silage –based (Beauchemin et al., 2000) TMR
when fed to early and mid-lactation cows, respectively. Furthermore, Lewis et al. (1999)
reported increased DMI at three different enzyme application rates with early lactation cows fed
a TMR in which the alfalfa hay and silage were treated with a T. reesei enzyme (Cornzyme,
Finnfeeds Int.).
156
Despite being applied at 3.4× the XYL dose and containing 25% of XYL (v/v), the MIX
treatment did not improve DMI as did XYL. This could be due to the fact that MIX provided
only 85% of enzyme activities in XYL on an applied basis. Furthermore, previous in vitro results
(Chapter 1) showed that each of the component enzymes of MIX (1A and 2A) was more
effective at increasing in vitro NDFD than MIX, and yet XYL gave the greatest increase of the
three treatments. These responses suggest that the doses and or ratios of key fibrolytic enzymes
in MIX were not ideal for increasing DMI. No effects on DMI of lactating dairy cows were
reported when a similar MIX preparation from the same company was applied at a slightly lower
dose (2.8 mL/kg) due to greater component activities to TMR containing 22% of 4 or 7-wk
regrowths of bermudagrass silage (Queiroz et al., 2011). Similarly, Bernard et al. (2010) reported
that adding an EFE from T. reesei (Promote NET, Cargill, Minnetonka, Mn) to TMR consisting
of basal alfalfa or bermudagrass silage (12% of TMR) had no effect on DMI. Likewise, Dean et
al. (2013) reported no increase in DMI of mid lactation cows when the same enzyme was applied
at ensiling, or at feeding (to concentrate, bermudagrass, or TMR) when diets contained 35%
bermudagrass silage (DM basis). Therefore, enzymes have not improved the DMI of dairy cattle
fed diets containing moderate to high levels of bermudagrass. The increased DMI observed in
this study is attributable to the high xylanolytic capacity of XYL, which is reflected in its high
xylanase activity, high xylanase to endoglucanase ratio (8:1) as well as its ability to increase the
saccharification of bermudagrass cell walls into xylose by 21,400% compared to the 1,700%
increase obtained with an equal dose of MIX (Chapter 3).
Mean milk yield in this study (40.7 kg/d) was similar to that of Bernard et al. (2010; 41.4
kg/d) and greater than those reported by Queiroz et al. (2011; 38.4 kg/d) and Dean et al. (2013;
157
32.0 kg/d). These differences are due to variations in the diet composition and the BW and
lactation stages of the cows in the studies.
Cows fed XYL had (P < 0.05) or tended to have (P < 0.1) greater milk yield than those
fed the Control diet during wk 3, 6, and 7and those fed MIX had greater milk yield than Control
cows during wk 6, 8 and 9 (Figure 6-1; treatment × week interaction, P = 0.04; Table 6-4).
Application of EFE also tended (P = 0.09) to increase mean yield of 3.5% FCM yield (40.7,
41.8, and 41.0 kg/d for CON, XYL, and MIX, respectively). Yang et al. (1999) and Schingoethe
et al. (1999) reported that EFE (Promote, Bioavance Technologies Inc., NE and Experimental
product, Finnfeeds Int., UK, respectively) application to the alfalfa hay and corn silage in TMR
increased milk yield of cows in mid (25.6 vs. 23.7 kg/d) and early lactation (27.3 vs. 25.4 FCM
kg/d), respectively. In the latter study, the responses occurred after 4 wk of feeding and were
maintained for 8 wk. In the current study, increases in milk production due to feeding XYL were
evident after 3 wk of feeding and persisted till wk 7, though differences at wk 4 and 5 were
numerical (P = 0.11 and 0.34, respectively). Increases in milk production due to feeding MIX
were evident (P < 0.01) at wk 6 to 9, except for wk 7 (P = 0.30). Reasons why the response to
MIX occurred later than that for XYL are unclear but they may be because XYL had more
xylanase and could consequently improve forage and diet quality earlier. Nevertheless, data from
both EFE indicate that beneficial responses of feeding EFE may not be evident for 3 to 6 wk
after feeding the EFE is initiated.
Cows fed XYL produced 2.8% more (P = 0.07, tendency) milk fat than those fed the
Control or MIX diets, which had similar milk fat yields. Applying EFE had no effect (P > 0.1)
on milk component concentrations, milk protein or lactose yield, feed efficiency, BW, BW
change, or BCS. Bernard et al. (2010) and Queiroz et al. (2011) reported no change in milk
158
composition or yield of milk components, feed efficiency, or BW change due to application of
EFE. However, Dean et al. (2013) detected trends for increased milk fat and protein
concentration and decreased feed efficiency when an EFE was applied to a TMR containing 35
% bermudagrass. However, no effect was evident when the EFE was applied just prior to feeding
to the bermudagrass silage or concentrate or to the bermudagrass silage at the point of ensiling.
Therefore, most of the studies in which EFE were applied to bermudagrass-based TMR detected
no effects on milk component concentrations or yields. Studies in which EFE were applied to
diets lacking bermudagrass such as the alfalfa hay - corn silage- based TMR of Arriola et al.
(2011b) and the barley-alfalfa silage – based TMR of Holtshausen et al. (2011) reported that EFE
treatment increased feed efficiency but did not affect milk yield or composition. Treatment with
EFE decreased DMI in the study of Holtshausen et al. (2011) but had no effect on DMI in that of
Arriola et al. (2011b). That XYL treatment increased DMI and milk production in this study but
not those cited above is probably due to the greater xylanase activity of XYL and the greater
hemicellulose concentration of the diet used in this study. These factors would have led to
greater fiber hydrolysis by XYL, which would reduce rumen fill, enhance DMI and thereby
increase the supply of nutrients required for milk production (Beauchemin and Holtshausen,
2010).
Applying EFE had no effect (P > 0.1) on the lag phase (h), washout fraction, potentially
degradable fraction, undegradable fraction, or fractional degradation rate of DM (Table 6-5).
Similar responses were reported in in situ ruminal degradation studies that examined effects of
applying EFE to the TMR (Arriola et al., 2011b; Holtshausen et al., 2011), compared different
EFE application strategies (Dean et al., 2013) or that applied EFE to bermudagrass hay (Krueger
et al., 2008; Romero et al., 2013).
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Fibrolytic enzyme application did not affect the ruminal pH or concentrations of
ammonia-N, total VFA, acetate (A), propionate (P), butyrate, isobutyrate, isovalerate, valerate,
or the A:P ratio (P > 0.1; Table 6-6). Previous in vitro studies (Chapters 3, 4, and 5) revealed
increased total VFA and propionate concentration and a decreased A:P ratio when XYL was
applied to bermudagrass haylage. Lack of a similar response in this study may be attributable at
least partly to application of XYL to the TMR in this study versus bermudagrass haylage in
previous studies. Since bermudagrass silage only represented 10% of the TMR used in this study,
beneficial effect of the EFE on bermudagrass silage may have been obscured by the fermentation
of the whole TMR. Many studies reported that dietary EFE had no effect on total VFA
concentration (Yang et al., 1999; Kung et al., 2002; Sutton et al., 2003). However, Arriola et al.
(2011b) reported that application of an EFE increased total VFA concentration and decreased the
A:P ratio in ruminal fluid of lactating dairy cows. In that study, total VFA concentration was
118.6 and 133.1 mM and A:P was 2.94 and 2.59 for the Control and enzyme-treated TMR
containing a similar level of concentrates (48%) to that in this study, respectively. The
comparatively greater total VFA concentration (156 mM) and the lower A:P ratio (2.5) in this
study are likely attributable to differences in DMI and the compositions of the diets and EFE in
the studies.
Conclusions
Application of XYL to the TMR increased DMI, OMI and CPI, and also increased milk
yield during wk 3, 6, and 7 as did MIX during wk 6, 8, and 9 and these responses were more
evident during peak versus early lactation. Furthermore, enzyme application tended to increase
FCM and fat yield. No EFE effects on milk component concentrations and yield of milk protein
and lactose, feed efficiency, BW, BW change and BCS were detected. Enzyme treatment did not
affect DM degradation kinetics in the rumen or pH and concentrations of VFA or ammonia-N in
160
ruminal fluid. Application of the XYL EFE to a bermudagrass-based TMR increased DMI and
milk production, implying that this EFE can be used to increase the performance of lactating
dairy cows fed diets containing up to 10% of bermudagrass in the southeast. This study also
validated the strategic approach to EFE evaluation that involved using in vitro tests to identify
EFE that would increase the performance of lactating dairy cows.
161
Table 6-1. Ingredient and chemical composition (mean ± SD) of the Control diet used for the in
situ and lactation study
Item Control NDF (% of DM) CP (% of DM)
Ingredient (% of DM)
Corn silage 35.1 40.7 ± 5.7 6.6 ± 0.6
Corn grain 20.6
Soybean meal 10.3
Bermudagrass silage 9.9 65 ± 2.2 13.0 ± 2.2
Citrus pulp 6.2
Whole cottonseed 5.2
Alfalfa-
orchardgrass hay
5.0 49.6 ± 3.5 12.0 ± 1.3
AminoPlus1 4.1
Mineral mix2 3.7
Chemical
DM (%) 64.8
OM (% of DM) 93.9
Ash (% of DM) 6.7
CP (% of DM) 16.5
NDF (% of DM) 32.9
ADF (% of DM) 17.4
Hemicellulose (% of DM)3 15.5
NFC (% of DM)4 39.3
NEL (Mcal/kg of DM) 1.59
1Ag Processing Inc. (Omaha, NE) 2 Mineral mix contained 26.4% CP, 5.06% Ca, 10.7% Na, 6.8% K, 4.1% Mg, 0.26% S, 1.6% P,
417 mg/kg of Mn, 665 mg/kg of Zn, 229 mg/kg of Cu, 2166 mg/kg of Fe, 24 mg/kg of Co, 14
mg/kg of I, 7.1 mg/kg of Se, 116,511 IU of vitamin A/kg, and 1164 IU of vitamin E/kg (DM
basis). 3 Hemicellulose was calculated as the difference between NDF and ADF. 4 Calculated as NFC = 100 – [CP+ ash + fat (NRC, 2001, values) + NDF]
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Table 6-2. Activities of endoglucanase, xylanase, exoglucanase, β-glucosidase (μmol of sugar released min-1 g-1) and ferulic acid
esterase (nmol of ferulic acid released min-1 g-1) and protein concentrations (mg g-1) of the exogenous fibrolytic enzyme
(EFE) preparations mixed with the dietary ingredients daily.
EFE1 Endoglucanase Xylanase Exoglucanase β-glucosidase
Ferulic acid
esterase
Protein
XYL 3,624 29,301 0.84 11.7 1.46 111.1
MIX 2,659 10,234 2.53 15.2 7.35 72.4
S.D. 107 221 0.03 0.9 0.29 25
1 XYL (Xylanase Plus) and MIX (75:25 Mixture of Cellulase Plus and XYL) were produced by Dyadic (Jupiter, FL) from non-
recombinant Trichoderma reesei.
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Table 6-3. Effect of addition of fibrolytic enzymes to diet on intake by lactating dairy cows.
Intake (kg/d) Treatment1,2
SEM
P value
CON XYL MIX TRT Week TRT × Week
DM 27.4b 28.6a 27.4b 0.41 0.048 < 0.001 0.526
OM 25.5b 26.7a 25.5b 0.36 0.031 < 0.001 0.513
CP 4.5b 4.7a 4.5b 0.06 0.023 < 0.001 0.450
NDF 8.7 9.1 8.7 0.20 0.212 < 0.001 0.530
Hemicellulose 4.1 4.3 4.1 0.09 0.202 < 0.001 0.480
ADF 4.6 4.8 4.6 0.11 0.227 < 0.001 0.488
1 CON= Control; XYL= Xylanase Plus; MIX= 75:25 Mixture of Cellulase Plus and XYL. Xylanase Plus and Cellulase Plus were
produced by Dyadic (Jupiter, FL) from non-recombinant Trichoderma reesei. 2 Means in the same row with different superscripts differed (P < 0.05).
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Table 6-4. Effect of dietary treatment with fibrolytic enzymes on milk yield, feed efficiency, yield and composition of milk fat, protein
and lactose, somatic cell counts, body weight and body condition score of lactating dairy cows.
Treatment2
SEM P value
Measure CON XYL MIX TRT Week TRT × Week
Milk yield
(kg/d)
40.0 41.2 40.8 0.46 0.152 < 0.001 0.035
3.5% FCM
(kg/d)
40.7 41.8 41.0 0.38 0.086 < 0.001 0.234
Milk protein
(%)
2.99 2.98 2.96 0.021 0.594 0.002 0.738
Milk fat (%) 3.60 3.62 3.53 0.038 0.176 < 0.001 0.378
Milk lactose
(%)
4.84 4.82 4.84 0.013 0.409 < 0.001 0.551
Milk protein
(kg/d)
1.19 1.22 1.20 0.013 0.190 < 0.001 0.473
Milk fat (kg/d) 1.44 1.48 1.44 0.015 0.068 < 0.001 0.464
Milk lactose
(kg/d)
1.93 1.98 1.97 0.026 0.383 0.001 0.103
SCC (× 1,000 /
mL)
119 68 97 32.0 0.517 0.250 0.489
Feed Efficiency
(FCM / DMI)
1.47 1.45 1.49 0.024 0.363 < 0.001 0.777
BW (kg) 601 606 596 3.6 0.198 < 0.001 0.655
BW change
(kg/d)
0.4 0.4 0.4 0.09 0.935
BCS (1 to 5) 3.1 3.2 3.2 0.05 0.608 < 0.001 0.972 1 CON= Control; XYL= Xylanase Plus; MIX= 75:25 Mixture of Cellulase Plus and XYL. Xylanase Plus and Cellulase Plus were
produced by Dyadic (Jupiter, FL) from non-recombinant Trichoderma reesei. 2 Means in the same row with different superscripts differed (P < 0.05).
165
Table 6-5. Effect of dietary treatment with fibrolytic enzymes on in situ ruminal dry matter degradation kinetics of a total mixed ration
in lactating dairy cows1
Measure Treatment2 SEM P-value
CON XYL MIX TRT
Lag phase, h 0.40 1.59 1.24 0.333 0.139
Washout fraction, %
of DM
37.9 38.4 39.4 0.56 0.276
Potentially
degradable DM
fraction, %
48.5 48.8 48.8 1.16 0.978
Fractional
degradation rate of
DM, h-1
0.064 0.068 0.058 0.0048 0.414
Undegradable DM
fraction, % after 72h
of incubation
13.6 12.7 11.9 1.16 0.616
1 CON= Control; XYL= Xylanase Plus; MIX= 75:25 Mixture of Cellulase Plus and XYL. Xylanase Plus and Cellulase Plus were
produced by Dyadic (Jupiter, FL) from non-recombinant Trichoderma reesei. 2 Means in the same row with different superscripts differed (P < 0.05).
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Table 6-6. Effect of dietary treatment with fibrolytic enzymes on ruminal fermentation measures of lactating dairy cows1
Measure Treatment2 SEM P-value
CON XYL MIX TRT Time TRT × Time
pH 6.10 6.10 6.07 0.135 0.979 < 0.001 0.250
Ammonia-N
(mg/dL)
9.7 8.8 10.1 1.20 0.733 < 0.001 0.583
Total VFA (mM) 155.0 149.4 164.7 6.66 0.292 0.002 0.386
Acetate (mM) 87.1 81.5 88.6 2.44 0.126 < 0.001 0.205
Propionate (mM) 35.5 34.0 40.8 2.46 0.164 0.001 0.451
Butyrate (mM) 17.9 17.5 18.8 1.01 0.659 < 0.001 0.519
Isobutyrate (mM) 1.0 1.1 1.1 0.07 0.740 < 0.001 0.362
Isovalerate (mM) 5.5 6.0 5.7 0.60 0.825 0.027 0.778
Valerate (mM) 7.3a 9.3a 9.0a 1.79 0.691 0.008 0.696
A:P ratio 2.6 2.5 2.3 0.11 0.212 < 0.001 0.991
1 CON= Control; XYL= Xylanase Plus; MIX= 75:25 Mixture of Cellulase Plus and XYL. Xylanase Plus and Cellulase Plus were
produced by Dyadic (Jupiter, FL) from non-recombinant Trichoderma reesei. 2 Means in the same row with different superscripts differed (P < 0.05).
167
Figure 6-1.Effect of dietary treatment with fibrolytic enzymes on milk yield of lactating dairy cows. (Treatment × Week, P = 0.035;
SEM = 0.59 kg/d); abc, xyz: means with different letters at the same week differed at P < 0.05 and 0.10, respectively.
CON= untreated TMR; MIX= TMR treated with enzyme Mixture (75:25 Mixture of Cellulase Plus and XYL); and XYL=
TMR treated with Xylanase Plus enzyme. Cows were at 35 + days in milk during wk1. Xylanase Plus and Cellulase Plus
were produced by Dyadic (Jupiter, FL) from non-recombinant Trichoderma reesei.
38.5
39
39.5
40
40.5
41
41.5
42
42.5
1 2 3 4 5 6 7 8 9 10
Mil
k y
ield
(k
g/d
)
Week
CON
MIX
XYL
a, x a
a
b, y
b
b
ab, x
ab ab x
y
xy
x
y
xy
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CHAPTER 7
GENERAL SUMMARY AND RECOMMENDATIONS
In the southeastern U.S, warm-season grasses are the basis of cattle production (Pitman,
2007), but their high fiber content and low digestibility limit animal productivity and
consequently profitability (Hanna and Sollenberger, 2007). Therefore, improving the nutritional
quality of warm-season grasses is a high priority for the dairy industry in the region (Southeast
Milk, Inc., 2011). Since bermudagrass is the most widely planted warm-season perennial grass
for dairy production in the southeast (10-12 million ha; Newman, 2007), it is an ideal model for
testing strategies to improve the quality of such grasses. Although some studies have shown that
EFE application to forages and diets improved forage digestibility and animal performance,
respectively (Beauchemin and Holtshausen, 2010; Adesogan et al., 2013), use of EFE in
ruminant diets is very limited. This is because their use has produced equivocal animal
performance results due to the wide array of conditions under which they have been tested and
limited understanding of their mode of action (Beauchemin and Holtshausen, 2010; Adesogan et
al., 2013). Effects of EFE are influenced by numerous factors such as the dose (Eun et al., 2007)
and activity composition (Eun and Beauchemin, 2007), the prevailing pH and temperature
(Arriola et al., 2011a), the animal performance level (Schingoethe et al., 1999), the experimental
design (Adesogan et al., 2013), and the fraction and proportion of the diet to which the enzyme is
applied (Krueger et al., 2008a; Dean et al., 2013). Arriola et al. (2011b) showed that adding an
EFE to a corn silage and alfalfa - based total mixed ration (TMR) fed to dairy cows increased
digestibility and increased feed efficiency. However, when the same enzyme was applied to a
bermudagrass silage-based TMR, none of these performance measures was increased (Queiroz et
al., 2011). Consequently, research was needed to optimize the use of EFE to improve the quality
of bermudagrass. A series of experiments was conducted to screen fibrolytic enzyme candidates
169
to identify those that were ideal for hydrolyzing bermudagrass and to examine conditions that
could optimize the response in order to identify the best candidates for increasing the
performance of dairy cows fed a bermudagrass silage-based diet.
The first study (Chapter 3) aimed to identify the best 5 EFE among 12 that improved the
in vitro NDF digestibility and fermentation (Experiment 1) and preingestive hydrolysis
(Experiment 2) of bermudagrass haylage. In addition, regression relationships between
enzymatic activities and measures of digestibility and preingestive hydrolysis were explored. A
proteomic assay was used to identify and quantify differences in the composition of the most-
and least-effective EFE at increasing the NDFD of bermudagrass haylage (Experiment 3). In
Experiment 1, Compared to the Control, 6 EFE-treated substrates had greater DMD, 9 had
greater NDFD, 5 had greater, 6 had greater total VFA concentration, and 4 had lower acetate to
propionate ratio. In Experiment 2, 3 EFE increased NDF hydrolysis, 10 increased
saccharification, and 8 increased the release of ferulic acid from cell walls. Regression analyses
revealed that enzyme activities accurately predicted preingestive hydrolysis measures (WSC, R2
= 0.95; FER, R2 = 0.99) and moderately predicted NDF hydrolysis (R2 = 0.65), but poorly
predicted digestibility measures (R2 <0.10). This indicates that enzyme activity estimates should
not be used to choose the best EFE for improving forage digestibility or animal performance.
The proteomics iTRAQ LC-MS analysis in Experiment 3 revealed that relative to the most
effective EFE, the least effective EFE at increasing NDFD contained lesser amounts of specific
enzymes and auxiliary proteins necessary for xylan and lignocellulose degradation. Five
promising EFE candidates that reduced the fiber concentration of bermudagrass and increased its
digestibility were identified (EFE 1A, 2A, 11C, 13D and 15D).
170
The second study (Chapter 4) aimed to examine the effects of the dose of the 5 most
promising EFE from Chapter 3 on in vitro fiber digestibility and fermentation (Experiment 1)
and preingestive fiber hydrolysis (Experiment 2) and to determine the optimum EFE doses that
maximize the NDFD of bermudagrass haylage efficiently. In Experiment 1, increasing the EFE
dose nonlinearly increased the DMD and NDFD of 1A, 13D, 2A and 11C. Doses with the
highest increases in NDFD were 2×, 2×, 1×, 0.5× and 0.5X for 1A, 2A, 11C, 13D and 15D,
respectively, where 1× was the EFE manufacturer-recommended dose. Increasing the dose of
2A, 11C, and 13D nonlinearly increased total VFA and propionate concentrations and decreased
the acetate to propionate ratios of 2A, 11C, and13D as did 1A and 15D (linear). In Experiment 2,
increasing the dose of all EFE increased NDF hydrolysis, saccharification and ferulic acid
release from cell walls and these responses were generally greatest with the 3× EFE dose. This
study revealed that increasing the dose of some EFE increased NDFD, fermentation and
preingestive hydrolysis but the optimal dose varied between EFE. Furthermore, the dose that
optimized preingestive hydrolysis differed from those that optimized NDFD, indicating that
preingestive hydrolysis is not an accurate indicator of effects of EFE on forage fiber digestibility.
The third study (Chapter 5) aimed to examine the effects of adding 5 cofactors (COF;
Mn2+ , Co2+, Fe2+, Ca2+, and Mg2+) to the 5 EFE selected in Chapter 3 on preingestive hydrolysis
(Experiment 1). The effect of increasing the COF dose on in vitro digestibility and fermentation
of bermudagrass haylage was examined in Experiment 2 using the best EFE-COF combinations
from Experiment 1 (Experiment 2). In Experiment 1, saccharification was increased by adding
all COF to EFE 2A and 11C and by adding Mn2+, Co2+, and Fe2+ to EFE 13D. In Experiment 2,
increasing the dose of Mn2+ with or without 11C linearly increased NDFD but the response was
greatest when10 mM of Mn2+ was added to 11C. This response reflected a synergistic increase in
171
NDFD that exceeded those due to adding the EFE or COF alone. However, increasing the dose
of Fe2+ in the absence of 2A linearly decreased NDFD but no effect was detected in the presence
of 2A. This indicates that certain EFE can prevent adverse effects of metal toxicity on forage
digestibility. However, increasing the Fe2+ dose decreased NDFD in the presence or absence of
13D, although when 13D was present NDFD was never lower than control. Therefore adding
some COF to EFE can synergistically enhance their hydrolytic effects but others may reduce
them.
The fourth study (Chapter 6) aimed to examine the effects of the most promising EFE
from the previous experiments (EFE 2A or XYL) and a previously effective EFE (MIX) on the
performance of dairy cows in early-lactation (Experiment 1) and the kinetics of ruminal
degradation of the diet (Experiment 2). In Experiment 1, application of XYL to the TMR
increased DMI, OMI and CPI, and also increased milk yield during wk 3, 6, and 7 as did MIX
during wk 6, 8, and 9 and these responses were more evident during peak versus early lactation.
Furthermore, enzyme application tended to increase FCM and fat yield. No EFE effects on milk
component concentrations, milk protein and lactose yields, feed efficiency, BW, BW change and
BCS were detected. Enzyme treatment did not affect in situ ruminal degradation kinetics or pH
and concentrations of volatile fatty acids or ammonia-N in ruminal fluid. In summary,
application of the EFE improved the productivity of cows fed a TMR that included warm-season
grasses.
It is noteworthy that the dose of XYL (1 g/kg TMR) was 3.4× less than that of MIX (3.4
g/kg). Since XYL costs 0.011 $/g and MIX costs 0.007 $/g, the costs of treating the TMR with
XYL and MIX were 0.011 and 0.024 $/kg TMR, respectively. Based on a ration cost of 0.35
$/kg TMR and a milk price of 0.57 $/kg (USDA, 2013) and using the intake and milk yield
172
values obtained in this experiment both XYL and MIX had negative margin over feed costs (-
0.05 and - 0.20 $, respectively). The XYL EFE would only be profitable if the cost was reduced
to 0.009 $/g or if the milk price increased to 0.62 $/kg. Mean milk prices have increased by
200% during the last 50 yr in Florida (0.15 $/kg in 1960 to 0.49 $/kg in 2012; Arriola and De
Vries, 2013). Therefore, it is plausible that milk prices will increase to the point that will make
application of XYL justifiable economically in the future. More research to improve the
efficiency of manufacturing EFE is needed in order to decrease the associated costs and thereby
make EFE use more attractive to the dairy industry. With lower EFE costs, higher doses of XYL
could become more practical and under this scenario, in vivo experiments should be conducted
to examine if higher doses of XYL continue to produce an economic increase in milk yield.
This study showed that application of a XYL to a bermudagrass-based TMR increased
DMI and milk production, implying that this EFE can be used to increase the performance of
lactating dairy cows in the Southeast provided the cost is decreased. Future studies should
examine the effect of applying EFE to TMR containing higher levels (> 10%) of bermudagrass
than that examined in Chapter 5.
The main conclusions from this series of studies are that a strategic approach to EFE
evaluation can be used to identify effective EFE that would increase the performance of lactating
dairy cows. The results from the studies indicate that EFE that have sufficient types and
quantities of enzymatic activities to hydrolyze extensively hemicellulose and improve NDFD
under ruminal conditions are good candidates for increasing forage digestibility and the
performance of lactating dairy cows. This study also showed that proteomic assays can identify
the key enzymes and auxiliary proteins necessary for optimizing ruminal forage digestion and
the performance of lactating dairy cows. Future research should up regulate the genes
173
responsible for producing these key enzymes and proteins and down regulate or inhibit the
expression of genes for those that are irrelevant or less important for xylan and lignocellulose
degradation by genetically engineering the microbial sources of the EFE. This will improve the
efficacy of EFE products used in ruminants and could potentially reduce the costs of enzymes by
increasing their potency and efficiency.
Recent research and investment in EFE for cellulosic ethanol production could
potentially benefit animal nutrition since many of the types of biomass used for bioethanol
production are forages. In addition to contributing to reducing enzyme costs as described above,
cell wall characterization assays used in the biofuel industry could improve our understanding of
effects of EFE application to forages and diets on xylan and lignocellulose degradation because
they quantify individual monosaccharaides components of fiber rather than estimate fiber
fractions from crude extracts like NDF, ADF and ADL. However, these techniques need to be
refined to make them less expensive, more precise and more amenable for evaluating large
numbers of samples.
Future research should also investigate the efficacy of using EFE as silage additives
because EFE-mediated saccharification will increase the availability of fermentable substrates,
which could increase the rate of acidification of silages and reduce DM losses. This is
particularly important for warm-season grasses, which typically lack sufficient sugars for rapid
acidification or legumes, which have high buffering capacities that reduce acidification.
Furthermore, the optimal pH for most of the xylanases and cellulases produced by T. reesei and
A. oryzae, which are the most widely used sources of commercial EFE, is 4-5, which is the
typical pH of silage. Future research will also need to determine if improvements in silage
174
acidification and fermentation due to EFE application culminate in improved performance
responses, when such silages are fed to lactating dairy cows.
Another aspect worthy of future research is to investigate the potential use of acetyl xylan
esterase to release acetate from the hemicellulose matrix. This could potentially reduce the
growth of spoilage-causing and pathogenic fungi and increase the aerobic stability of silage
because of the antifungal nature of the released acetate. If this approach is successful, it would
decrease the need to increase the aerobic stability of silage with expensive acids like propionic
acid or heterofermentative inoculants, which produce acetate from lactate but also slightly
increase losses of DM as CO2 from silages.
Exogenous fibrolytic enzyme technology has considerable potential to improve the
performance of dairy and beef cattle and small ruminants, because such EFE can be used to
improve the quality of forages that ruminants rely on to produce much needed human food.
175
LIST OF REFERENCES
Adesogan, A. T. 2005. Improving forage quality and animal performance with fibrolytic
enzymes. Pages 91-109 in Florida Ruminant Nutrition Symposium Proc., University of
Florida – IFAS. Gainesville, FL.
Adesogan, A. T., Z. X. Ma, J. J. Romero, and K. G. Arriola. 2013. Improving cell wall digestion
and animal performance with fibrolytic enzymes. J. Anim. Sci.: In press.
Aguiar, R. S., M. H. L. Silveira, A. P. Pitarelo, M. L. Corazza and L. P. Ramos. 2013. Kinetics
of enzyme-catalyzed hydrolysis of steam-exploded sugarcane bagasse. Bioresour.
Technol. 147: 416-423.
Akin, D. E., 1989.Histological and physical factors affecting digestibility of forages. Agron. J.
81: 17-25.
Aman, P. 1993. Composition and structure of cell wall polysaccharides in forages. Pages 183-
200 in Forage Cell Wall Structure and Digestibility. H. G. Jung, D. R. Buxton, R. D.
Hatfield and J. Ralph, eds. ASA, CSSA and SSSA., Madison, WI.
Anderson, W. F., J. Peterson, D. E. Akin and W. H. Morrison. 2005. Enzyme pretreatment of
grass lignocellulose for potential high-value co-products and an improved fermentable
substrate. Pages 303-310 in Twenty-sixth Symposium on Biotechnology for Fuels and
Chemicals. B. H. Davison, B. R. Evans, M. Finkelstein and J. D. McMillan, eds. Humana
Press, Chattanooga, TN.
Andric, P., A. S. Meyer, P. A. Jensen, and K. Dam-Johansen. 2010. Reactor design for
minimizing product inhibition during enzymatic lignocellulose hydrolysis: I. Significance
and mechanism of cellobiose and glucose inhibition on cellulolytic enzymes. Biotechnol.
Adv. 28(3):308-324.
Arelovich, H. M., F. N. Owens, G. W. Horn and J. A. Vizcarra. 2000. Effects of supplemental
zinc and manganese on ruminal fermentation, forage intake, and digestion by cattle fed
prairie hay and urea. J. Anim. Sci. 78(11):2972-2979.
Arriola, K. G., J. J. Romero and A. T. Adesogan. 2011a. Effects of pH and temperature on
fibrolytic enzyme activities of various commercial exogenous enzyme preparations.
ADSA-ASAS Joint Annual Meeting, New Orleans, Louisiana, USA. J. Anim. Sci. 89(E-
Suppl. 1):554. (Abstr).
Arriola, K. G., S. C. Kim, C. R. Staples, and A. T. Adesogan. 2011b. Effect of fibrolytic enzyme
application to low- and high-concentrate diets on the performance of lactating dairy
cattle. J. Dairy Sci. 94(2):832-841.
Arriola, K. G., and A. T. Adesogan. 2013. Effect of fibrolytic enzyme application on the
digestibility of corn silage, alfalfa hay, two concentrates, and complete diets under
simulated ruminal and preruminal conditions. Anim. Nutr. Feed Techn. 13:263-276.
176
Arriola K. G., and A. De Vries. 2013. Florida Dairy Industry Statistics: Economic Measures.
AN287. IFAS Extension, Gainesville. Accessed November 29,
2013.http://edis.ifas.ufl.edu/pdffiles/DS/DS13100.pdf.
AOAC. 2000. Official Methods of Analysis. Association of Official Analytical Chemists. USA.
Bach-Knudsen, K. E. and B. W. Li. 1991. Determination of oligosaccharides in protein-rich
feedstuffs by gas-liquid chromatography and high-performance liquid chromatography. J.
Agr. Food Chem. 39(4):689-694.
Bailey, K. 2009. Managing income over feed costs. Agricultural Research and Cooperative
Extension. Accessed November 10, 2013.
http://pubs.cas.psu.edu/FreePubs/pdfs/ua458.pdf.
Balasko, J. A., and C. J. Nelson. 2003. Grasses for northern areas. Pages 125-148 in Forages: An
Introduction to Grassland Agriculture. M. Collins, ed. Iowa State Press, IA.
Bansal, P., M. Hall, M. J. Realff, J. H. Lee, and A. S. Bommarius. 2009. Modeling cellulase
kinetics on lignocellulosic substrates. Biotechnol. Adv. 27(6):833-848.
Barnes, R. F. and C. J. Nelson. 2003. Forages and grasslands in a changing world. Pages 3-25 in
Forages: An Introduction to Grassland Agriculture. R. F. Barnes, C. J. Nelson, and K. J.
Moore, eds. Blackwell Publishing Co., Ames, IA.
Barton, F. E., H. E. Amos. D. Burdick, and R. L. Wilson. 1976. Relationship of chemical
analysis to in vitro digestibility for selected tropical and temperate grasses. J. Anim. Sci.
43(2):504-512.
Batistoti, C., B. Lemp, L. Jank, M. Moris, A. C. Cubas, R. A. Gomes, and M. V. B. Ferreira.
2012. Correlations among anatomical, morphological, chemical and agronomic
characteristics of leaf blades in Panicum maximum genotypes. Anim. Feed. Sci. Tech.
171:173-180.
Beauchemin, K. A., W. Z. Yang, and L. M. Rode. 1999. Effects of grain source and enzyme
additive onsite and extent of nutrient digestion in dairy cows. J. Dairy Sci. 82:378-390.
Beauchemin, K. A., L. M. Rode, M. Maekawa, D. P. Morgavi, and R. Kampen. 2000. Evaluation
of a nonstarch polysaccharidase feed enzyme in dairy cow diets. J. Dairy Sci. 83:543-
553.
Beauchemin, K. A., D. Colombatto, D. P. Morgavi, and W. Z. Yang. 2003. Use of exogenous
fibrolytic enzymes to improve feed utilization by ruminants. J. Anim. Sci. 81:37-47.
Beauchemin, K. A., D. Colombatto, and D. P. Morgavi. 2004. A rationale for the development of
feed enzyme products for ruminants. Can. J. Anim. Sci.84: 23–36.
177
Beauchemin, K. A. and L. Holtshausen. 2010. Developments in enzyme usage in ruminants.
Pages 206-230 in Enzymes in Farm Animal Nutrition, 2nd ed. M. R. Bedford and G. G.
Partridge, eds. CAB International. Oxforshire, UK.
Bedford M. R. and J. Apajalahti. 2001. Microbial interactions in the response to exogenous
enzyme utilization. Pages 299-314 in Enzymes in Farm Animal Nutrition. M. R. Bedford
and G. G. Partridge, eds. CAB International. Oxforshire, UK.
Bedford, M. R. and G. G. Partridge. 2010. Feed enzymes, the future: Bright hope or regulatory
minefield? Pages 304-312 in Enzymes in Farm Animal Nutrition, 2nd ed. M. R. Bedford
and G. G. Partridge, eds. CAB International. Oxforshire, UK.
Bergman, E. N. 1990. Energy contributions of volatile fatty acids from the gastrointestinal tract
in various species. Physiol. Rev. 70(2):567-590.
Bernard, J. K., J. J. Castro, N. A. Mullis, A. T. Adesogan, J. W. West, and G. Morantes. 2010.
Effect of feeding alfalfa hay or Tifton 85 bermudagrass haylage with or without a
cellulase enzyme on performance of Holstein cows. J. Dairy Sci. 93(11):5280-5285.
Bey, M., S. Zhou, L. Poidevin, B. Henrissat, P. M. Coutinho, J.-G.Berrin, and J.-C. Sigoillot.
2013. Cello-oligosaccharide oxidation reveals differences between two lytic
polysaccharide monooxigenases (Family GH61) from Podospora anserina. Appl.
Environ. Microb. 79(2):488-496.
Bhat, M. K., S. I. McCrae., and T. M. Wood. 1989. The endo-(1→4)-β- -glucanase system of
Penicillium pinophilum cellulase: Isolation, purification, and characterization of five
major endoglucanase components. Carbohyd. Res. 190:279-297.
Bhat, M. K., J. Solomon, and M. Ramesh. 1993. Purification and characterization of an
extracellular β-glucosidase from the thermophilic fungus Sporotrichum thermophile and
its influence on cellulase activity. J. Gen. Microbiol. 139:2825-2832
Bhat, M. K., and G. P. Hazlewood. 2001. Enzymology and other characteristics of cellulases and
xylanases. Pages 11-60 in Enzymes in Farm Animal Nutrition. M. R. Bedford and G. G.
Partridge, eds. CABI Publications. Wallingford, UK.
Bin, Y. and C. Hongzhang. 2010. Effect of the ash on enzymatic hydrolysis of steam-exploded
rice straw. Bioresour. Technol. 101(23):9114-9119.
Bio-Rad. 2011. Overview and strategies for bio-organic molecule purification. Accessed
November 10, 2013. http://www.bio-
rad.com/webroot/web/pdf/lsr/literature/Bulletin_1928.pdf.
Biswas, P. K. and B. Purnendu. 2012. In vitro evaluation of a diet supplemented with Mn on
nutrient digestibility and rumen fermentation pattern in cattle. Explor. Anim. Med. Res.
1(2):161-166.
178
Bommarious, A. S., A. Katona, S. E. Cheben, A. S. Patel, A. J. Ragauskas, K. Knudson, and Y.
Pu. 2008. Cellulase kinetics as a function of cellulose pretreatment. Metab. Eng.
10(6):370-381.
Boraston, A., D. N. Bolam, H. J. Gilbert, and G. J. Davies. 2004. Carbohydrate-binding modules:
fine-tuning polysaccharide recognition. Biochem. J. 382:769-781.
Borneman, W. S., L. G. Ljungdahl, R. D. Hartley, and D. E. Akin. 1991. Isolation and
characterization of p-coumaroyl esterase from the anaerobic fungus Neocallimastix strain
MC-2. Appl. Environ. Microb. 57(8):2337-2344.
Bradford, M. 1976. A rapid and sensitive method for the quantitation of microgram quantities of
protein utilizing the principle of protein-dye binding. Anal. Biochem. 72:248–254.
BRENDA. 2013. The Comprehensive Enzyme Information System. Accessed November 10,
2013. http://www.brenda-enzymes.org/.
Broderick, J. B. 2001. Coenzymes and Cofactors. Pages 1-11 in Encyclopedia of Life Sciences.
John Wiley & Sons, Ltd. New York, NY.
Buckmaster, D. R., A.J. Heinrichs., R.A. Ward, and B.P. Lammers. 1997. Characterizing
effective fiber with particle size and fiber concentration interactions. Pages 53-54 in Proc.
of the 18th International Grassland Congress. ID: 886. Saskatoon, Saskatchewan,
Canada.
Casler, M. D., and H. J. G. Jung. 1999. Selection and evaluation of smooth bromegrass clones
with divergent lignin and etherified ferulic acid concentration. Crop. Sci. 39:1866-1873.
Cassab, G. I. 1998. Plant cell wall proteins. Annu. Rev. Plant Physiol. Plant Mol. Biol. 49:281-
309.
CAZy. 2013. Carbohydrate Active Enzymes. Accessed November 10, 2013.
http://www.cazy.org/.
Chahine, M. 2004. Forages in total mixed rations. University of Idaho Extension. Accessed
November 10, 2013.
http://www.extension.uidaho.edu/forage/Proceedings/2004%20Proceedings%20pdf/chahi
ne.pdf. http://www.cazy.org/.
Chamberlain, C. C., and W. Burroughs. 1962. Effect of Fluoride, Magnesium and Manganese
Ions on in Vitro Cellulose Digestion by Rumen Microorganisms. J. Anim. Sci. 21(3):428-
432.
Cherney, J. H., and G. C. Marten. 1982. Small grain crop forage potential: II. Interrelationships
among biological, chemical, morphological, and anatomical determinants of quality.
Crop Sci. 22:240-243.
179
Chung, M., A. Shilton, B. Guieysse, and C. Pratt. 2012. Contribution of milk and feed wastes to
methane emissions from New Zealand dairy farm. Chemeca 2012: Pages 937 in Quality
of life through chemical engineering. Wellington, New Zealand.
Clarkson, K., B. Jones, R. Bott, B. Bower, G. Chotani and T. Becker. 2001. Enzymes: Screening,
expression, design and production. Pages 315-352 in Enzymes in Farm Animal Nutrition.
M. Bedford and G. Partridge, ed. CABI Publishing, Oxford, UK.
Cochran, W. G. 2007. Sampling techniques. John Wiley & Sons. New York, NY.
Cochran R.C., W.K. Coblentz, and E.S. Vanzant. 2007. Animal methods for evaluating forage
quality. Pages 541-552 in Forages: The Science of Grassland Agriculture. R.F. Barnes,
C.J. Nelson, K.J. Moore and M. Collins, ed. Blackwell Publishing. Ames, IA.
Coleman S. W., J. E Moore, and J. R. Wilson. 2004. Quality and utilization. Pages 267-308 in
Warm Season (C4) Grasses. L. E. Moser, B. L. Burson and L. E. Sollenberger, eds. ASA,
CSSA and SSSA. Madison, WI.
Collins, T., C. Gerday, and G. Feller. 2005. Xylanases, xylanase families and extremophilic
xylanases. FEMS Microbiol. Rev. 29(1):3-23.
Colombatto, D. 2000. Use of enzymes to improve fibre utilization in ruminants. A biochemical
and in vitro rumen degradation assessment. Ph.D. Diss., Univ. of Reading, U.K.
Colombatto, D., D. P. Morgavi, A. F. Furtado, and K. A. Beauchemin. 2003. Screening of
exogenous enzymes for ruminant diets: Relationship between biochemical characteristics
and in vitro ruminal degradation. J. Anim. Sci. 81: 2628-2638.
Considine, P. J., and M. P. Coughlan. 1989. Production of carbohydrate-hydrolyzing enzyme
blends by solid-state fermentation. Pages 273-281 in Enzyme Systems for Lignocellulose
Degradation. M.P. Coughlan, ed. Elsevier Applied Science, New York.
Coughlan, M. P., M. G. Tuohy, E. X. F. Filho, J. Puls, M. Clayessens, M. Vrsanská, and M. M.
Hughes.1993. Enzymological aspects of microbial hemicellulases with emphasis on
fungal systems. Pages 53-84 in Hemicellulose and Hemicellulases. M. P. Coughlan and
G. P. Hazlewood, eds. Portland Press, London.
Cowan, W. D. 1994. Factors affecting the manufacture, distribution, application and overall
quality of enzymes in poultry feeds. Pages 175-184 in Joint Proc. 2nd Int. Roundtable on
Anim. Feed Biotechnol-Probiotics, and Workshop on Anim. Feed Enzymes, Ottawa,
Canada.
Dairy One. 2013. Dairy One Forage Laboratory Service. Accessed November 12, 2013.
http://www.dairyone.com/Forage/
de Vries, R. P. and J. Visser. 2001. Aspergillus enzymes involved in degradation of plant cell
wall polysaccharides. Microbiol. Mol. Biol. R. 65(4):497-522.
180
Dean, D. B., A. T. Adesogan, N. Krueger, and R. C. Littell. 2005. Effect of fibrolytic enzymes
on the fermentation characteristics, aerobic stability, and digestibility of bermudagrass
silage. J. Dairy Sci. 88:994-1003.
Dean, D. B., A. T. Adesogan, N. A. Krueger and R. C. Littell. 2008. Effects of treatment with
ammonia or fibrolytic enzymes on chemical composition and ruminal degradability of
hays produced from tropical grasses. Anim. Feed Sci. Technol. 145(1-4): 68-83.
Dean, D. B., C. R. Staples, R. C. Littell, S. C. Kim, and A. T. Adesogan. 2013. Effect of method
of adding a fibrolytic enzyme to dairy cow diets on feed intake digestibility, milk
production, ruminal fermentation, and blood metabolites. Anim. Nutr. Feed Techn.
13:287-302.
Demeyer, D. I. 1991.Quantitative aspects of microbial metabolism in the rumen and hindgut.
Pages 217-237 in Rumen microbial metabolism and ruminant digestion. J. P. Jouany, ed.
INRA Editions, Paris, France.
Dien, B. S., H.-J. G. Jung, K. P. Vogel, M. D. Casler, J. F. S. Lamb, L. Iten, R. B. Mitchell and
G. Sarath. 2006. Chemical composition and response to dilute-acid pretreatment and
enzymatic saccharification of alfalfa, reed canarygrass, and switchgrass. Biomass
Bioenerg. 30(10):880-891.
DMI. 2004. Dairy Management Inc. U.S. Dairy: The economic impact of the dairy industry.
Dairy Management, Inc. Rosemont, IL. Accessed November 11, 2013.
http://future.aae.wisc.edu/publications/US_Outlook_May_2004.pdf.
DuBois, M., K. A. Gilles, J. K. Hamilton, P. A. Rebers, and F. Smith. 1956. Colorimetric method
for determination of sugars and related substances. Anal. Chem. 28:350-356.
Dunnett, C.W. 1955. A multiple comparison procedure for comparing several treatments with a
control. J. Amer. Statist Assoc. 50:1096-1121.
Durand, M., and R. Kawashima. 1980. Influence of minerals in rumen microbial digestion. Pages
375-408 in Digestive Physiology and Metabolism in Ruminants. Y. Ruckebusch and P.
Thivend, eds. Springer Netherlands.
Eriksson, K. E. L., R. A. Blanchette, and P. Ander. 1990. Microbial and enzymatic degradation
of wood and wood components. Springer-Verlag, New York, N.Y.
Eun, J. S., and K. A. Beauchemin. 2005. Effects of proteolytic feed enzyme on intake, digestion,
ruminal fermentation, and milk production. J. Dairy Sci. 88(6):2140-2153.
Eun, J. S., and K. A. Beauchemin. 2007. Enhancing in vitro degradation of alfalfa hay and corn
silage using feed enzymes. J. Dairy Sci. 90(6):2839-2851.
Eun, J. S., K. A. Beauchemin, and H. Schulze. 2007. Use of exogenous fibrolytic enzymes to
enhance in vitro fermentation of alfalfa hay and corn silage. J. Dairy Sci. 90(3):1440-
1451.
181
Eun, J. S., and K. A. Beauchemin. 2008. Relationship between enzymic activities and in vitro
degradation of alfalfa hay and corn silage. Anim. Feed Sci. Technol. 145:53-67.
Evert, R. F. 2006. Esau’s plant anatomy, meristems, cells and tissues of the plant body: Their
structure, function, and development. 3rd ed. John Wiley & Sons, Inc. Hoboken, N. J.
ExplorEnz. 2013. The Enzyme Database. Accessed November 10, 2013. http://www.enzyme-
database.org/index.php.
Faulds, C. B., and G. Williamson. 1994. Purification and characterization of a ferulic acid
esterase (FAE-III) from Aspergillus niger: specificity for the phenolic moiety and binding
to microcrystalline cellulose. Microbiology+ 140(4): 779-787.
Faulds, C. B. 2010. What can feruloyl esterases do for us? Phytochem. Rev. 9:121-132
FAWN. 2013. Florida Automated Weather Network. Accessed September 28, 2013.
http://fawn.ifas.ufl.edu.
Feng, P., W. Hunt, G. T. Pritchard, and W. E. Julien. 1996. Effect of enzyme preparations on in
situ and in vitro degradation and in vivo digestive characteristics of mature cool-season
grass forage in beef steers. J. Anim. Sci. 74:1349-1357.
Fontes, C. M. G. A., J. Hall, B. H. Hirst, G. P. Hazlewood, and H. J. Gilbert. 1995. The
resistance of cellulases and xylanases to proteolytic inactivation. Appl. Microbiol.
Biotechnol. 43:52-57.
France, J. and j. Dijkstra. 2005. Volatile fatty acid production. Pages 157-176 in Quantitative
aspects of ruminant digestion and metabolism, 2nd ed. J. Dijkstra, J. M. Forbes and J.
France, eds. CABI Publishing. Oxfordshire, UK.
Gado, H. M., A. Z. M. Salem, P. H. Robinson, and M. Hassan. 2009. Influence of exogenous
enzymes on nutrient digestibility, extent of ruminal fermentation as well as milk
production and composition in dairy cows. Anim. Feed Sci. Technol. 154:36-46.
Galyean, M. L. 2010. Laboratory procedures in animal nutrition research. Texas Tech
University, Lubbock, Department of Animal and Food Sciences. Accessed November 10,
2013. http://www.depts.ttu.edu/afs/home/mgalyean/lab_man.pdf.
Gashe, B. A. 1992. Cellulase production and activity by Trichoderma sp. A-001. J. Appl.
Bacteriol. 73:79-82.
Gevens, A., and R.L. Nicholson. 2000. In This Issue Cutin Composition: A Subtle Role For
Fungal Cutinase? Physiol. Mol. Plant Pathol. 57:43-45.
Giraldo, L. A., M. L. Tejido, M. J. Ranilla, and M. D. Carro. 2008. Effects of exogenous
fibrolytic enzymes on in vitro ruminal fermentation of substrates with different forage:
concentrate ratios. Anim. Feed Sci. Technol. 141:306-325.
182
Glass, N. L., M. Schmoll, J. H. D. Cate, and S. Coradetti. 2013. Plant cell wall deconstruction by
Ascomycete fungi. Annu. Rev. Microbiol. 67:477-498.
Glusker, J. P. 2011. Encyclopedia of inorganic and bioinorganic chemistry. John Wiley & Sons,
Inc. New York, NY.
Gochman, N., and J. M. Schmitz. 1972. Application of a new peroxide indicator reaction to the
specific, automated determination of glucose with glucose oxidase. Clin. Chem. 18:943-
952.
Goering, H. K., and P. J. Van Soest. 1970. Forage fiber analysis (apparatus, reagents, procedures,
and some applications). Agric. Handbook No. 379. ARS USDA, Washington, DC.
Gorka, P., Z. M. Kowalski, P. Pietrzak, A. Kotunia, R. Kiljanczyk, J. Flaga, J. J. Holst, P.
Guilloteau, and R. Zabielski. 2009. Effect of sodium butyrate supplementation in milk
replacer and starter diet on rumen development in calves. J. of Physiol. Pharmacol.
60:47-53.
Gourlay, K., J. Hu, V. Arantes, M. Andberg, M. Saloheimo, M. Penttilä, and J. Saddler. 2013.
Swollenin aids in the amorphogenesis step during the enzymatic hydrolysis of pretreated
biomass. Bioresour. Technol. 142:498-503.
Grabber, J. H. 2005. How do lignin composition, structure, and cross-linking affect
degradability? A review of cell wall model studies. Crop Sci. 45:820-831.
Grabber, J. H., J. Ralph, and R. D. Hatfield. 1998a. Severe inhibition of maize wall degradation
by synthetic lignins formed with coniferaldehyde. J. Sci. Food Agric. 78:81-87.
Grabber, J. H., J. Ralph, and R. D. Hatfield. 1998b. Ferulate cross-links limit the enzymatic
degradation of synthetically lignified primary walls of maize. J. Agric. Food Chem.
46:2609-2614.
Grant, R. 2009. Fiber rumen function, dairy cow health and productivity. Pages 80-111 in Proc.
Dairy Cow Academy. Gainesville, FL. Prince Agri., Quincy, IL.
Gwayumba, W., and D. A. Christensen. 1997. The effect of fibrolytic enzymes on protein and
carbohydrate degradation fractions in forages. Can. J. Anim. Sci. 77:541–542. (Abstr.)
Hacker, J. B., and D. J. Minson. 1981. The digestibility of plant parts. Herb. Abstr. 51:459-482.
Hanna W. W., and L. E. Sollenberger.2007 Tropical and subtropical grasses. Pages 245-256 in
Forages: The Science of Grassland Agriculture. R. F. Barnes, C. J. Nelson, K. J. Moore
and M. Collins, eds. Blackwell Publishing. Ames, IA.
183
Harris, P. V., D. Welner, K. C. McFarland, E. Re, J.-C. Navarro Poulsen, K. Brown, R. Salbo, H.
Ding, E. Vlasenko, S. Merino, F. Xu, J. Cherry, S. Larsen, and L. L leggio. 2010.
Stimulation of lignocellulosic biomass hydrolysis by proteins of glycoside hydrolase
family 61: Structure and function of a large, enigmatic family. Biochemistry-US
49(15):3305-3316.
Harrison, G. A., K. A. Dawson, and R. W. Hemken. 1992. Effects of high iron and sulfate ion
concentrations on dry matter digestion and volatile fatty acid production by ruminal
microorganisms. J. Anim. Sci. 70(4):1188-1194.
Hartley, R. D. 1972. p-Coumaric and ferulic acid components of cell walls of ryegrass and their
relationships with lignin and digestibility. J. Sci. Food Agric. 23:347-54.
Hartley, R. D., and D. E. Akin. 1989. Effect of forage cell wall phenolic acids and derivatives on
rumen microflora. J. Sci. Food Agr. 49(4):405-411.
Hatfield, R. D. 1993. Cell wall polysaccharide interactions and degradability. Pages 285-313 in
Forage Cell Wall Structure and Digestibility. H. G. Jung, D. R. Buxton, R. D. Hatfield,
and J. Ralph, ed. ASA, CSSA and SSSA. Madison, WI.
Hatfield, R. D., J. Ralph, and J. H. Grabber. 1999a. Cell wall foundations: molecular basis for
improving forage digestibilities. Crop Sci. 39: 27-37.
Hatfield, R. D., J. Ralph and J. H. Grabber. 1999b. Review: Cell wall cross linking by ferulates
and diferulates in grasses. J. Sci. Food Agric. 79:403-407.
Hatfield, R. D., H. J. G. Jung, G. Broderick, and T. C. Jenkins. 2007. Nutritional chemistry of
forages. Pages 467-485 in Forages: The Science of Grassland Agriculture. R.F. Barnes,
C.J. Nelson, K.J. Moore and M. Collins, ed. Blackwell Publishing. Ames, IA.
Henrissat, B. 1991. A classification of glycosyl hydrolases based on amino acid sequence
similarities. Biochem. J. 280: 309-316.
Holtshausen, L., Y.-H. Chung, H. Gerardo-Cuervo, M. Oba, and K. A. Beauchemin. 2011.
Improved milk production efficiency in early lactation dairy cattle with dietary addition
of a developmental fibrolytic enzyme additive. J. Dairy. Sci. 94(2):899-907.
Hristov, A. N., L. M. Rode, K. A. Beauchemin, and R. L. Wuerfel. 1996. Effect of a commercial
enzyme preparation on barley silage in vitro and in sacco dry matter degradability. Pages
282–284 in Proc. Western Section Meeting, Am. Soc. Anim. Sci. New Mexico State
Univ. New Mexico.
Hristov, A. N., T. A. McAllister, and K. J. Cheng. 1998. Effect of dietary or abomasal
supplementation of exogenous polysaccharide-degrading enzymes on rumen fermentation
and nutrient digestibility. J. Anim. Sci. 76(12):3146-3156.
184
Hristov, A. N., T. A. McAllister, and K. J. Cheng. 2000. Intraruminal supplementation with
increasing levels of exogenous polysaccharide-degrading enzymes: effects on nutrient
digestion in cattle fed and barley grain diet. J. Anim. Sci. 78(2):477-487.
Hu G., J. Koh, M.J. Yoo, K. Grupp, S. Chen, and J.F. Wendel. 2013. Proteomic profiling of
developing cotton fibers from wild and domesticated Gossypium barbadense. New
Phytol. 200(2):570-582.
Hu, J., V. Arantes, and J. Saddler. 2011. The enhancement of enzymatic hydrolysis of
lignocellulosic substrates by the addition of accessory enzymes such as xylanase: is it an
additive or synergistic effect? Biotechnol. Biofuels. 4:1-13.
Huang, X. L., and M. H. Penner. 1991. Apparent substrate-inhibition of the Trichoderma reesei
cellulase system. J. Agr. Food Chem. 39(11):2096-2100.
Humberstone, F. J., and D. E. Briggs. 2000. Extraction and assay of acetic acid esterase from
Malted Barley. J. I. Brewing 106(1):31-38.
Igarashi, K., T. Uchihashi, A. Koivula, M. Wada, S. Kimura, T. Okamoto, M. Penttilä, T. Ando,
and M. Samejima. 2011. Traffic jams reduce hydrolytic efficiency of cellulose on
cellulose surface. Science 333 (6047):1279-1282.
Jalak, J., M. Kurašin, H. Teugjas, and P. Väljamäe. 2012. Endo-exo Synergism in Cellulose
Hydrolysis Revisited. J. Biol. Chem. 287:28802-28815.
John, M., and J. Schmidt. 1988. Xylanases and β-xylosidase of Trichoderma lignorum. Method.
Enzymol. 160:662-671.
Jung, H. J. G. 2003. Maize stem tissues: Ferulate deposition in developing internode cell walls.
Phytochemistry 63:543-549.
Jung, H. J. G., and M. D. Casler. 1991. Relationship of lignin and esterified phenolics to
fermentation of smooth bromegrass fibre. Anim. Feed Sci. Technol. 32:63-68.
Jung H. J. G., and D. A. Deetz. 1993. Cell wall lignification and degradability. Pages 315-346 in
Forage Cell Wall Structure and Digestibility. H. G. Jung, D. R. Buxton, R. D. Hatfield
and J. Ralph, eds. ASA, CSSA and SSSA. Madison, WI.
Jung, H. J. G., and M. S. Allen. 1995. Characteristics of plant cell walls affecting intake and
digestibility of forages by ruminants. J. Anim. Sci. 73:2774-2790.
Jung, H. J. G., W. Ni, C. C. S. Chapple, and K. Meyer. 1999. Impact of lignin composition on
cell wall degradability in an Arabidopsis mutant. J. Sci. Food Agric. 79:922-928.
Jung, H. J. G., and F. M. Engels. 2002. Alfalfa stem tissues: cell wall deposition, composition,
and degradability. Crop Sci. 42: 524-534.
185
Kanauchi, M., S. Watanabe, T. Tsukada, K. Atta, T. Kakuta, and T. Koizumi. 2008. Purification
and characteristics of feruloyl esterase from Aspergillus awamori G-2 strain. J. Food Sci.
73(6):458-463.
Kennedy, P. M., and P. Doyle. 1993. Particle-size reduction by ruminant-effects of cell wall
composition. Pages 499-534 in Forage Cell Wall Structure and Digestibility. H. G. Jung,
D. R. Buxton, R. D. Hatfield and J. Ralph, ed. ASA, CSSA and SSSA. Madison, WI.
Klyosov, A. 1990.Trends in biochemistry and enzymology of cellulose degradation.
Biochemistry-US 29:10577-10585.
Knowlton, K. F., B. P. Glenn, and R. A. Erdman. 1998. Performance, ruminal fermentation, and
site of starch digestion in early lactation cows fed corn grain harvested and processed
differently. J. Dairy Sci. 81(7):1972-1984.
Knowlton, K. F., J. M. McKinney, and C. Cobb. 2002. Effect of a direct-fed fibrolytic enzyme
formulation on nutrient intake, partitioning, and excretion in early and late lactation
holstein cows. J. Dairy Sci. 85:3328-3335.
Koga, J., Y. Baba, A. Shimonaka, T. Nishimura, S. Hanamura, and T. Kono. 2008. Purification
and Characterization of a New Family 45 Endoglucanase, STCE1, from Staphylotrichum
coccosporum and Its Overproduction in Humicola insolens. Appl. Environ. Microb.
74(13):4210-4217.
Kononoff, P. J., and A. J. Heinrichs. 2003. The effect of corn silage particle size and cottonseed
hulls on cows in early lactation. J. Dairy Sci. 86(7):2438-2451.
Krause, M., K. A. Beauchemin, L. M. Rode, B. I. Farr, and P. Nørgaard. 1998. Fibrolytic
enzyme treatment of barley grain and source of forage in high-grain diets fed to growing
cattle. J. Anim. Sci. 96: 1010–1015.
Krueger, N. A. 2007. Effect of fibrolytic enzymes on the nutritive value of tropical forages and
performance of beef steers. Ph.D. dissertation. Univ. of Florida, Gainesville.
Krueger, N. A., A. T. Adesogan, C. R. Staples, W. K. Krueger, S. C. Kim, R. C. Littell, and L. E.
Sollenberger. 2008. Effect of method of applying fibrolytic enzymes or ammonia to
bermudagrass hay on feed intake, digestion, and growth of beef steers. J. Anim. Sci.
86:882-889.
Kung, L. Jr., R. J. Treacher, G. A. Nauman, A. M. Smagala, K. M. Endres, and M. A. Cohen.
2000. The effect of treating forages with fibrolytic enzymes on its nutritive value and
lactation performance of dairy cows. J Dairy Sci. 83:115-122
Kung, L., Jr., M. A. Cohen, L. M. Rode, and R. J. Treacher. 2002. The effect of fibrolytic
enzymes sprayed onto forages and fed in a total mixed ration to lactating dairy cow. J.
Dairy Sci. 85:2396-2402
186
Lai, K. K., G. L. Lorca, and C. F. Gonzalez. 2009. Biochemical properties of two cinnamoyl
esterases purified from Lactobacillus johnsonii strain isolated from stool samples of
diabetes-resistant rats. Appl. Environ. Microb. 75:5018-5024.
Laidler, K. J., and B. F. Peterman. 2009. Temperature effects in enzyme kinetics. Pages 178-196
in Contemporary Enzyme Kinetics and Mechanism, 3rd ed. D. L. Purich, ed. Elsevier.
Oxford, UK.
Lam, T. B. T., K. Iiayama, and B. Stone. 1991. Quantitative determination of lignin-
polysaccharide associations in wheat internode cell walls: Ferulic acid bridges. Pages 29-
33 in Proc. 6th Int. Symp. of Wood and Paper Chemistry, Melbourne, Australia. 29 Apr. -
3 May 1991. Australian Pulp and Paper Ind. Tech. Assoc. (APPITA), Parkville, Victoria.
Lam, T. B. T., K. Iiyama, and B. Stone. 1992. Cinnamic acid bridges between cell wall polymers
in wheat and phalaris internodes. Phtytochemistry 31:1179-1183.
Levasseur, A., M. Saloheimo, D. Navarro, M. Andberg, F. Monot, T. Nakari-Setälä, M. Asther,
and E. Record. 2006. Production of a chimeric enzyme tool associating the Trichoderma
reesei swollenin with the Aspergillus niger feruloyl esterase A for release of ferulic acid.
Appl. Microbiol. Biot. 73(4):872-880.
Lewis, G. E., C. W. Hunt, W. K. Sanchez, R. Treacher, G. T. Pritchard, and P. Feng. 1996.
Effect of direct-fed fibrolytic enzymes on the digestive characteristics of a forage-based
diet fed to beef steers. J. Anim. Sci. 74:3020-3028.
Lewis, G. E., W. K. Sanchez, C. W. Hunt, M. A. Guy, G. T. Pritchard, B. I. Swanson, and R. J.
Treacher. 1999. Effect of direct-fed fibrolytic enzymes on the lactational performance of
dairy cows. J. Dairy Sci. 82:611-617.
Liu, Q., C. Wang, W. Z. Yang, B. Zhang, X. M. Yang, D. C. He, P. Zhang, K.H. Dong, and Y.
X. Huang. 2009. Effects of isobutyrate on rumen fermentation, lactation performance and
plasma characteristics in dairy cows. Anim. Feed Sci. Technol. 154:58-67.
Littell, R. C., Milliken, G. A., Stroup, W. W., Wolfinger, R. D., and O. S. Schabenberger. 2006.
Analysis of repeated measures. Pages 159-204 in SAS for Mixed Models. SAS Institute
Inc. US.
Lopez, S. 2005. In vitro and in situ techniques for estimating digestibility. Pages 87-122 in
Quantitative Aspects of Ruminant Digestion and Metabolism. CABI Publishing.
Wallingford, UK
Lynd, L. R., P. J. Weimer, W.H. van Zyl, and I. S. Pretorius. 2002. Microbial cellulose
utilization: Fundamentals and biotechnology. Microbiol. Mol. Biol. R. 66:506-577.
Mandebvu, J. W., J. W. West, G. M. Hill, R. N. Gates, R. D. Hatfield, B. G. Mullinix, A. H.
Parks and A. B. Caudle. 1999. Comparison of tifton 85 and coastal bermudagrass for
yield, nutrients traits, intake, and digestion by growing beef steers. J. Anim. Sci. 77:1572-
1586.
187
Marsh, W. H., B. Fingerhut, and H. Miller. 1965. Automated and manual direct methods for the
determination of blood urea. Clin. Chem.. 11:624-627.
Martinez, A., and D. C. Church. 1970. Effect of Various Mineral Elements on In Vitro Rumen
Cellulose Digestion. J. Anim. Sci. 31(5):982-990.
Mathieu, Y., E. Gelhaye, S. Dumarçay, P. Gérardin, L. Harvengt, and M. Buée. 2013. Selection
and validation of enzymatic activities as functional markers in wood biotechnology and
fungal ecology. J. Microbiol. Meth. 92(2):157-163.
Matsui, M., I. Toyosawa, and M. Fukuda. 1995. Purification and characterization of a glycine-
rich protein from the aleurone layer of soybean seeds. Biosci. Biotech. Bioch. 59:2231-
2234.
McAllister, T. A., H. D. Bae, G. A. Jones, and K. J. Cheng. 1994. Microbial attachment and feed
digestion in the rumen. J. Anim. Sci. 72:3004–3018.
McAllister, T. A., A. N. Hristov, K. A. Beauchemin, L. M. Rode, and K. J. Cheng,. 2001.
Enzymes in ruminant diets. Pages 273-298 in Enzymes in Farm Animal Nutrition. M.
Bedford and G. Partridge, ed. CABI Publishing, Oxford, UK.
McCrae, S. I., K. M. Leith, A. H. Gordon, and T. M. Wood. 1994. Xylan-degrading enzyme
system produced by the fungus Aspergillus awamori: isolation and characterization of a
feruloyl esterase and a p-coumaroyl esterase. Enzyme Microb. Technol. 16(10):826-834.
Merchen, N. R. 1988. Digestion, absorption and excretion in ruminants. Pages 172-201 in The
Ruminant Animal. D. C. Church, ed. Waveland Press, Inc. Long Grove, IL.
Mertens, D. R. 1977. Dietary fiber components: relationship to the rate and extent of ruminal
digestion. Fed. Proc. 1977:87-92.
Mertens, D. R. 1997. Creating a system for meeting the fiber requirements of dairy cows. J.
Dairy Sci. 80(7):1463-1481.
Mertens, D. R. 2003. Challenges in measuring insoluble dietary fiber. J. Anim. Sci. 81(12):3233-
3249.
Mertens, D. R. 2007. Digestibility and Intake. Pages 487-508 in Forages: The Science of
Grassland Agriculture. R. F. Barnes, C. J. Nelson, K. J. Moore and M. Collins, eds.
Blackwell Publishing. Ames, IA.
Mohammed, R., S. M. McGinn, and K. A. Beauchemin. 2011. Prediction of enteric methane
output from milk fatty acid concentrations and rumen fermentation parameters in dairy
cows fed sunflower, flax, or canola seeds. J. Dairy. Sci. 94:6057-6068.
Moore, K. J., and R. D. Hatfield. 1994. Carbohydrates and forage quality. Pages 229-280 in
Forage quality, evaluation and utilization. G. C. Fahey, ed. ASA, CSSA and SSSA.
Madison, WI.
188
Moore, K. J., K. J. Boote, and M. A. Sanderson. 2004. Physiology and developmental
morphology. Pages 179-216 in Warm-Season (C4) grasses. L. E. Moser, B. L. Burson and
L. E. Sollenberger, ed. ASA, CSSA and SSSA. Madison, WI.
Morgavi, D. P., C. J. Newbold, D. E. Beever, and R. J. Wallace. 2000a. Stability and
stabilization of potential feed additive enzymes in rumen fluid. Enzyme Microb. Technol.
26:171–177.
Morgavi, D. P., K. A. Beauchemin, V. L. Nsereko, L. M. Rode, A. D. Iwaasa, W. Z. Yang, T. A.
McAllister, and Y. Wang. 2000b. Synergy between ruminal fibrolytic enzymes and
enzymes from Trichoderma Longibrachiatum. J. Dairy Sci. 83:1310–1321.
Morgavi, D. P., K. A. Beauchemin, V. L. Nsereko, L. M. Rode, T. A. McAllister, A. D. Iwaasa,
Y. Wang, and W. Z. Yang. 2001. Resistance of feed enzymes to proteolytic inactivation
by rumen microorganisms and gastrointestinal proteases. J. Anim. Sci. 79:1621–1630.
Morgavi, D. P., K. A. Beauchemin, V. L. Nsereko, L. M. Rode, T. A. McAllister, and Y. Wang.
2004. Trichoderma enzymes promote Fibrobacter succinogenes S85 adhesion to, and
degradation of, complex substrates but not to pure cellulose. J. Sci. Food Agric. 84:1083-
1090.
Moser, L. E. and C. J. Nelson. 2003. Structure and Morphology of grasses. Pages 25-50 in
Forages an introduction to grassland agriculture. 6th ed. R. F. Barnes, C. J. Nelson, M
Collins and K. J. Moore, ed. Blackwell Publishing, Ames, IA.
Moser, L. E. and J.A. Jennings. 2007. Grass and legume structure and morphology. Pages 15-36
in Forages: The Science of Grassland Agriculture. R.F. Barnes, C.J. Nelson, K.J. Moore
and M. Collins, ed. Blackwell Publishing. Ames, IA.
Mouriño, F., R. Akkarawongsa, and P. J. Weimer. 2001. Initial pH as a determinant of cellulose
digestion rate by mixed ruminal microorganisms in vitro. J. Dairy Sci. 84(4):848-859.
Muck, R. E., and J. T. Dickerson. 1988. Storage temperature effects on proteolysis in alfalfa
silage. Trans. ASAE 31: 1005–1009.
Nakagawa, Y., V. G. H. Eijsink, K. Totani, and G. Vaaje-Koolstad. 2013. Conversion of α-
Chitin substrates with varying particle size and crystallinity reveals substrate preferences
of the chitinases and lytic polysaccharide monooxygenase of Serratia marcescens. J. Agr.
Food Chem. In Press.
Nelson, D. L., and M. M. Cox. 2008. Lehninger Principles of Biochemestry.5th ed. Freeman and
Company. New York, NY.
Newman, Y. 2007. Costal vs. Tifton 85 Bermudagrass for Horses.Facts & Myths.University of
Florida, IFAS Extension. Accessed November 10, 2013.
http://cflag.ifas.ufl.edu/documents/2007EquineInstit/CoastalvsT-85.pdf.
189
Noel, R. J., and L. G. Hambleton. 1976. Collaborative study of a semi automated method for
determination of crude protein in animal feeds. J. Assoc. Offic. Anal. Chem. 59: 134-140.
NRC. 2000. Nutrient requirements of beef cattle. 7th ed. National Academy Press. Washington,
DC.
NRC. 2001. Nutrient requirements of dairy cattle. 7th ed. National Academy Press. Washington,
DC.
Nsereko, V. L., D. P. Morgavi, and L. M. Rode. 2000. Effect of fungal enzyme preparations on
hydrolysis and subsequent degradation of alfalfa hay fiber by mixed rumen
microorganisms in vitro. Anim. Feed Sci. Technol. 88:153-170
Nsereko, V. L., K. A. Beauchemin, D. P. Morgavi, L. M. Rode, A. F. Furtado, T. A. McAllister,
D. Iwaasa, W. Z. Yang, and Y. Wang. 2002. Effect of a fibrolytic enzyme preparation
from Trichoderma longibrachiatum on the rumen microbial population of dairy cows.
Can. J. Microbiol. 48:14–20.
Nutt, A. 2006. Hydrolytic and oxidative mechanisms involved in cellulose degradation. Doctoral
Thesis, Uppsala University. Acta Universitatis Upsaliensis.
O’Neill, M. A. and W. S. York. 2003. The composition and structure of plant primary cell walls.
Pages 1- 54 in The Plant Cell Wall. J. K. C. Rose, ed. Annual Plant Reviews, Vol. 8.
Balckwell Publishing, CRC Press. Oxford, UK.
O’Sullivan, A. C. 1997. Cellulose: The structure slowly unravels. Cellulose 4(3):173-207.
Oba, M., and M. S. Allen. 1999. Evaluation of the importance of the digestibility of neutral
detergent fiber from forage: Effects on dry matter intake and milk yield of dairy cows. J.
Dairy Sci. 82:589-596.
Owens, F. N., and A. L. Goetsch. 1988. Ruminal Fermentation. Pages 145-171 in The Ruminant
Animal: Digestive Physiology and Nutrition. D. C. Church, ed. Waveland Press, Inc.
Long Grove, IL.
Pahlow, G., R. E. Muck, F. Driehuis, S. J. W. H. Oude Elferink and S. F. Spoelstra. 2003.
Microbiology of ensiling. Pages 31-93 in Silage Science and Technology. D. R. Buxton,
R. E. Muck and J. H. Harrison, eds. ASA, CSSA and SSSA. Madison, WI.
Paloheimo, M., J. Pironen, and J. Vehamaanperä. 2010. Xylanases and cellulases as feed
aditives. Pages 12-53 in Enzymes in Farm Animal Nutrition, 2nd ed. M Bedford and G.
Partridge, eds. CAB International, Oxfordshire, UK.
Phillips, C. M., I. William T. Beeson, J. H. Cate, and M. A. Marletta. 2011. Cellobiose
Dehydrogenase and a Copper-Dependent Polysaccharide Monooxygenase Potentiate
Cellulose Degradation by Neurospora crassa. ACS Chem. Biol. 6(12):1399-1406.
190
Pitman, W. D. 2007. Forage systems for humid areas. Pages 303-311 in Forages: The Science of
Grassland Agriculture. R. F. Barnes, C. J. Nelson, K. J. Moore and M. Collins, eds.
Blackwell Publishing. Ames, IA.
Poutanen, K. 1988. An α-L-arabinofuranosidase of Trichoderma reseei. J. Biotechnol. 7(4):271-
281.
Purich, D. L. 2011. Enzyme kinetics. Catalysis & control: a reference of theory and best-practice
methods. Elsevier, London, UK.
Qing, Q., B. Yang, and C. E. Wyman. 2010. Xylooligomers are strong inhibitors of cellulose
hydrolysis by enzymes. Bioresour. Technol. 101(24):9624-9630.
Queiroz, O. C. M., A. T. Adesogan, J. L. P. Daniel, J. J. Romero, J. H. Shin, C. R. Staples, and J.
E. P. Santos. 2011. Effects of adding fibrolytic enzymes to diets containing bermudagrass
silage harvested at two maturity stages on the performance of lactating Holstein cattle.
ADSA-ASAS Joint Annual Meeting, New Orleans, Louisiana, USA. J. Anim. Sci. 89(E-
Suppl. 1):378. (Abstr).
Quinlan, R. J., M. D. Sweeney, L. L. Leggio, H. Otten, J.-C. N. Poulsen, K. S. Johansen, K. B.
R. M. Krogh, C. I. Jørgensen, M. Tovborg, A. Anthonsen, T. Tryfona, C. P. Walter, P.
Dupree, F. Xu, G. J. Davies, and P. H. Walton. 2011. Insights into the oxidative
degradation of cellulose by a copper metalloenzyme that exploits biomass components. P.
Natl. Acad. Sci. USA 108(37):15079-15084.
Ralph, J. 1996. Cell wall cross-linking in grasses: The importance of understanding plant
chemistry and biochemistry. Informational Conference with Dairy and Forage Industries.
US Dairy Forage Center, Madison, WI.
Ralph, J., and R. Helm. 1993. Lignin/hydroxycinnamic acid/polysaccharide complexes: synthetic
models for regiochemical characterization. Pages 201-246 in Forage Cell Wall Structure
and Digestibility. H. G. Jung, D. R. Buxton, R. D. Hatfield and J. Ralph, ed. ASA, CSSA
and SSSA. Madison, WI.
Ralph, J., J. H. Grabber, and R. D. Hatfield. 1995. Lignin-ferulate crosslinks in grasses: Active
incorporation of ferulate polysaccharide esters into ryegrass lignins. Carbohyd. Res.
275:167-178.
Rinne, M., S. Jaakkola, and P. Huhtanen. 1997. Grass maturity effects on cattle fed silage-based
diets. 1. Organic matter digestion, rumen fermentation and nitrogen utilization. Anim.
Feed Sci. Technol. 67:1-17.
Riou, C., J. M. Salmon, M. J. Vallier, Z. Günata, and P. Barre. 1998. Purification,
Characterization, and Substrate Specificity of a Novel Highly Glucose-Tolerant β-
Glucosidase from Aspergillus oryzae. Appl. Environ. Microb. 64(10):3607-3614.
Rode, L. M., W. Z. Yang, and K. A. Beauchemin. 1999. Fibrolytic enzyme supplements for dairy
cows in early lactation. J. Dairy Sci. 82(10):2121-2126.
191
Rode, L. M., T. A. McAllister, and K. A. Beauchemin, D.P. Morgavi, V.L. Nsereko, W.Z.
Yang.2001.Enzymes as direct-fed additives for ruminants. Pages 301-332 in
Biotechnology in Animal Husbandry. Springer Netherlands eds.
Romero, J. J., M. A. Zarate, O. C. M. Queiroz, J. H. Han, J. H. Shin, C. R. Staples, W. F. Brown
and A. T. Adesogan. 2013. Fibrolytic enzyme and ammonia application effects on the
nutritive value, intake, and digestion kinetics of bermudagrass hay in beef cattle. J. Anim.
Sci. 91: 4345-4356.
Rosen, G. D. 2010. Holo-analysis of the efficacy of exogenous enzyme performance in farm
animal nutrition. Pages 273-303 in in Enzymes in Farm Animal Nutrition, 2nd ed. M. R.
Bedford and G. G. Partridge, eds. CAB International. Oxforshire, UK.
Rudall, P. 2007. Anatomy of flowering plants. 3rd ed. Cambridge University Press, NY.
Russell, J. B. 2002. Rumen microbiology and its role in ruminant nutrition. ARS-USDA. Ithaca,
NY.
Saad, R. R., and E. M. Fawzi. 2004. Purification and some properties of an extracellular 1,4-β-
endoglucanase from Trichoderma longibrachiatum. New Egypt. J. Microbiol. 8:1-9.
Saha, B. C. 2003. Hemicellulose bioconversion. J. Ind. Microbiol. Biot. 30(5):279-291.
Saloheimo, M., M. Paloheimo, S. Hakola, J. Pere, B. Swanson, E. Nyyssönen, A. Bhatia, M.
Ward, and M. Penttilä. 2002. Swollenin, a Trichoderma reesei protein with sequence
similarity to the plant expansins, exhibits disruption activity on cellulosic materials. Eur.
J. Biochem. 269(17):4202-4211.
Sanchez, W. K., C. W. Hunt, M. A. Guy, G. T. Pritchard, B. I. Swanson, T. B. Warner, and J. M
Higgins. 1996. Effect of fibrolytic enzymes on lactational performance of dairy cows. J.
Dairy Sci. 79 (Suppl. 1):183. (Abstr).
Sang, S. L., G. Li, X. P. Hu, and Y. H. Liu. 2011. Molecular Cloning, Overexpression and
Characterization of a Novel Feruloyl Esterase from a Soil Metagenomic Library. J. Mol.
Microb. Biotech. 20(4):196-203.
Sarkanen, K. V., and C. H. Ludwig. 1971. Lignins: Occurrence, formation, structure and
reactions. Wiley-Interscience, New York.
Schingoethe, D. J., G. A. Stegeman, and R. J Treacher. 1999. Response of lactating dairy cows to
a cellulase and xylanase enzyme mixture applied to forages at the time of feeding. J.
Dairy Sci. 82: 996-1003
Schwarz, W. H. 2001. The cellulosome and cellulose degradation by anaerobic bacteria. Appl.
Microbiol. Biotechnol. 56:634–649.
Segel, I. H. 1976. Biochemical calculations. John Wiley and Sons, New York, NY.
192
Selig, M. J., E. P. Knoshaug, W. S. Adney, M. E. Himmel, and S. R. Decker. 2008. Synergistic
enhancement of cellobiohydrolase performance on pretreated corn stover by addition of
xylanase and esterase activity. Bioresource Technol. 99(11):4997-5005.
Shallom, D. and Y. Shoham. 2003. Microbial hemicellulases. Curr. Opin. Microbiol. 6(3):219-
228.
Silva-Sanchez, C., S. Chen, N. Zhu, Q. B.Li, and P. S. Chourey. 2013. Proteomic comparison of
basal endosperm in maize miniature1 mutant and its wild-time Mn1. Front. Plant Sci.
4:211.
Singh, A., A. K. Agrawal, A. B. Abidi, and N. S. Darmwal. 1990. Properties of exoglucanase
from Aspergillus niger. J. Gen. Appl. Microbiol. 36(4):245-253.
Soberon, M. A., J. H. Cherney, R. H. Liu, D. A. Ross, and D. J. R. Cherney. 2012. Free ferulic
acid uptake in lactating cows. J. Dairy Sci. 95(11):6563-6570.
Southeast Milk. 2011. Accessed November 10,
2013.https://secure.southeastmilk.org/v2/links/#news.
Staples, C. R. 2003. Bermudagrass: Growing, Storing, and Feeding for Dairy Animals. Vol.
2013. IFAS Extension, Gainesville. Accessed November 10, 2013.
http://edis.ifas.ufl.edu/pdffiles/DS/DS13100.pdf.
Sun, J. X., X. F. Sun, R. C. Sun, and Y. Q. Su. 2004. Fractional extraction and structural
characterization of sugarcane bagasse hemicelluloses. Carbohyd. Polym. 56(2):195-2004.
Sun, R., J. M. Lawther, and W. B. Banks. 1996. Fractional and structural characterization of
wheat straw hemicelluloses. Carbohyd. Polym. 29(4):325-331.
Sutton, J. D., R. H. Phipps, D. E. Beever, D. J. Humphries, G. F. Hartnell, J. L. Vicini, and D. L.
Hard. 2003. Effect of method of application of a fibrolytic enzyme product on digestive
processes and milk production in Holstein-Friesian cows. J. Dairy Sci. 86:546-556.
Svihus, B. 2010.Effect of digestive tract conditions, feed processing and ingredients on response
to NSP enzymes. Pages 129-159 in in Enzymes in Farm Animal Nutrition. M. R. Bedford
and G. G. Partridge, eds. CABI Publications. Wallingford, UK.
Tejirian, A., and F. Xu. 2010. Inhibition of Cellulase-Catalyzed Lignocellulosic Hydrolysis by
Iron and Oxidative Metal Ions and Complexes. Appl. Environ. Microb. 76(23):7673-
7682.
Terashima, N., K. Fukushima, L-F. He, and K. Takabe. 1993. Comprehensive model of the
lignified cell wall. Pages 247-270 in Forage Cell Wall Structure and Digestibility. H. G.
Jung, D. R. Buxton, R. D. Hatfield, and J. Ralph, eds. ASA, CSSA and SSSA. Madison,
WI.
193
Thakur, B. R., R. K. Singh, A. K. Handa and M. A. Rao. 1997. Chemistry and uses of pectin – A
review. CRC Cr. Rev. Food Sci. 37(1):47-73.
Theander, O. and E. Westerlund. 1993. Quantitative analysis of cell wall components. Pages 83-
104 in Forage Cell Wall Structure and Digestibility. H. G. Jung, D. R. Buxton, R. D.
Hatfield and J. Ralph, eds. ASA, CSSA and SSSA. Madison, WI.
Theander, O., P. Udén and P. Aman. 1981. Acetyl and phenolic acid substituents in timothy of
different maturity and after digestion with rumen microorganisms or a commercial
cellulose. Agr. Environ. 6:127-133.
Theander, O. 1995. Total dietary fiber determined as neutral sugar residues, uronic acid residues,
and Klason lignin (the Uppsala method): collaborative study. J. AOAC Int. 78(4):1030-
1044.
Tipton, K. F., A. G. McDonald, and H. B. F. Dixon. 2009. Effects of pH on enzymes. Pages 123-
177 in Contemporary enzyme kinetics and mechanisms, 3rd ed. D. L. Purich, ed.
Academic Press, London, UK.
Tricarico, J. M., J. D. Johnston, and K. A. Dawson. 2008. Dietary supplementation of ruminant
diets with an Aspergillus oryzae alpha-amylase. Anim. Feed Sci. Technol. 145(1-4):136-
150.
USDA. 2013. Florida Fluid Milk Report. Florida Marketing Area. Atlanta, GA. Accessed:
November 18, 2013. http://www.fmmatlanta.com/pdfstorage/curbulletinfl.pdf.
Vahjen, W., and O. Simon. 1999. Biochemical characteristics of non starch polysaccharide
hydrolyzing enzyme preparations designed as feed additives for poultry and piglet
nutrition. Arch. Tierernahr. 52(1):1-14.
Väljamäe, P., K. Kipper, G. Pettersson and G. Johansson. 2003. Synergistic cellulose hydrolysis
can be described in terms of fractal-like kinetics. Biotechnol. Bioeng. 84(2): 254-257.
Van de Vyver, W. F. J., K. A. Dawson, N. H. Casey, and J. M. Tricarico. 2004. Effect of
glycosylation on the stability of fungal xylanase exposed to proteases or rumen fluid in
vitro. Anim. Food Sci. Technol. 116: 259-269.
Van Soest, P.J. 1987. Effect of environment and quality of fibre on the nutritive value of crop
residues. International Livestock Centre for Africa, Addis Adaba, Ethiopia.
Van Soest, P. J. 1994. Nutritional Ecology of the Ruminant, 2nd ed. Cornell. University Press,
Ithaca, NY.
Van Soest, P. J., J. B. Robertson, and B. A. Lewis. 1991. Methods for dietary fiber, neutral
detergent fiber and non-starch polysaccharides in relation to animal nutrition. J. Dairy
Sci. 74:3568-3597.
194
VandeHaar, M. J. 2005. Dairy cattle. Pages 413-438 in Basic animal nutrition and feeding, 5th
ed. W. G. Pond, D. C. Church, K. R. Pond, P. A. Schoknecht, ed. John Wiley & Sons,
Inc.
Vanholme, R., B. Demedts, K. Morreel, J. Ralph, and W. Boerjan. 2010. Lignin biosynthesis and
structure. Plant Physiology 153(3):895-905.
Vicini, J. L., H. G. Bateman, M. K. Bhat, J. H. Clark, R. A. Erdman, R. H. Phipps, M. E. van
Amburgh, G. F. Hartnell, R. L. Hintz, and D. L. Hard. 2003. Effect of feeding
supplemental fibrolytic enzymes or soluble sugars with malic acid on milk production. J.
Dairy Sci. 86:576-585.
Vlasenko, E., M. Shulein, J. Cherry, and F Xu. 2010. Substrate specificity of family 5, 6, 7, 9,
12, and 45 endoglucanases. Bioresour. Technol. 101(7):2405-2411.
Voet, D., J. G. Voet and C. W. Pratt. 2010. Fundamentals of Biochemistry. Upgrade edition.
John Wiley & Sons, Inc. New York, NY.
Vogel, J. 2008. Unique aspects of the grass cell wall. Curr. Opin. Plant Biol. 11(3):301-307.
Wallace, R. J., S. J. A. Wallace, N. McKain, V. L. Nsereko, and G. F. Hartnell. 2001. Influence
of supplementary fibrolytic enzymes on the fermentation of corn and grass silages by
mixed ruminal microorganisms in vitro. J. Anim. Sci. 79:1905-1916.
Wang, Y., T. A. McAllister, L. M. Rode, K. A. Beauchemin, D. P. Morgavi, V. L. Nsereko, A.
D. Iwaasa, and W. Yang. 2001. Effects of an exogenous enzyme preparation on microbial
protein synthesis, enzyme activity and attachment to feed in the rumen simulation
technique. Brit. J. of Nutr. 2001. 85:325-332
Wang, Y. and T. A. McAllister. 2002. Rumen microbes, enzymes and feed digestion – a review.
Asian-Aust. J. Anim. Sci. 15:1659-1676.
Wang, Z. and H. Feng. 2010. Fractal kinetic analysis of the enzymatic saccharification of
cellulose under different conditions. Bioresour. Technol. 101:7995-8000.
Wang, Y., J. E. Ramirez-Bribiesca, L. J. Yanke, A. Tsang and T. A. McAllister. 2012. Effect of
exogenous fibrolytic enzyme application on the microbial attachment and digestion of
barley straw in vitro. Asian-Aust. J. Anim. Sci. 25(1):66-74.
Weimer, P. J., J. M. Hackney, H. J. G. Jung, and R. D. Hatfield. 2000. Fermentation of a
bacterial cellulose/xylan composite by mixed ruminal microflora: Implications for the
role of polysaccharide matrix interactions in plant cell wall biodegradability. J. Agric.
Food Chem. 48: 727-1733.
White, D., J. Drummond, and C. Fuqua. 2012. The physiology and Biochemistry of Prokaryotes,
4th ed. Oxford University Press. New York, NY.
195
Wildman, E. E., G. M. Jones, P. R. Wagner, and R. L. Boman. 1982. A dairy cow body condition
scoring system and its relation to selected production characteristics. J. Dairy Sci.
65:495-501.
Willats, W. G. T., L. McCartney, W. Mackie and J. P. Knox. 2001. Pectin: cell biology and
prospects for functional analysis. Plant Mol. Biol. 47:9-27.
Wilman, D., and M. A. K. Altimimi. 1982. The digestibility and chemical composition of plant
parts in Italian and perennial ryegrass during primary growth. J. Sci. Food Agric. 33:595-
602.
Wilson, J. R. 1976. Variation of leaf characteristics with level of insertion on a grass tiller. II.
Anatomy. Aust. 1. Agric. Res. 27:355-364.
Wilson, J. R. 1990. Influence of plant anatomy on digestion and fibre breakdown. Pages 99-117
in Microbial and Plant Opportunities to Improve the Utilization of Lignocellulose by
Ruminants. D. E. Akins, R. B. Brussel, L. G. Ljungdhal, J. R. Wilson, and P. J. Harris ed.
Elsevier Sci. Pub. Co., New York.
Wilson, J. R. 1991. Plant structures, their digestive and physical breakdown. Pages 207-216 in
Recent Advances on the Nutrition of Herbivores. Y. W. Ho, H. K. Wong, N. Abdullah,
and Z. A. Tajuddin, ed. Malay.Soc. of Anim. Prod. Kuala Lumpur.
Wilson, J. R. 1993. Organization of forage plant tissues. Pages 1-32 in Forage Cell Wall
Structure and Digestibility. H. G. Jung, D. R. Buxton, R. D. Hatfield, and J. Ralph, ed.
ASA, CSSA and SSSA. Madison, WI.
Wilson, J. R., and R. D. Hatfield. 1997. Structural and chemical changes of cell wall types during
stem development: Consequences of fibre degradation by rumen microflora. Austr.
Agric. Re. 48:165-180.
Wilson, J.R., and P.M. Kennedy. 1996. Plant and animal constraints to voluntary feed intake
associated with fiber characteristics and particle breakdown and passage in ruminants.
Aust. J. Agric. Res. 47:199-225.
Withers, S., and Williams, S. 2013. Glycoside Hydrolases in CAZypedia.Accessed 01 October
2013. http://www.cazypedia.org/.
Wood, T. M. 1985. Properties of cellulolytic enzyme systems. Biochem. Soc. Trans. 13: 407-
410.
Wood, T. M., and S. I. McCrae. 1986. The cellulase of Penicillium pinophilum: synergism
between enzyme components in solubilizing cellulose with special reference to the
involvement of two immunologically distinct CBHs. Carbohyd. Res. 234:93-99.
Wood, T. M., and K. M. Bhat. 1988. Methods for measuring cellulase activities. Pages 87-112 in
Methods in Enzymology. W. A. Wood, and S. T. Kellogg, eds. Vol. 160. Academic
Press. London, U.K.
196
Ximenes, E., Y. Kim, N. Mosier, B. Dien, and M. Ladisch. 2010. Inhibition of cellulases by
phenols. Enzyme Microb. Technol. 46:170-176.
Yang, W. Z., K. A. Beauchemin, and L. M. Rode. 1999. Effects of an enzyme feed additive on
extent of digestion and milk production of lactating dairy cows. J Dairy Sci. 82:391-403.
Yang, W. Z., K. A. Beauchemin, and L. M. Rode. 2000. A comparison of methods of adding
fibrolytic enzymes to lactating cow diet. J Dairy Sci. 83:2512-2520.
Zebeli, Q., J. R. Aschenbach, M. Tafaj, J. Boguhn, B. N. Ametaj, and W. Drochner. 2012.
Invited review: Role of physically effective fiber and estimation of dietary fiber adequacy
in high-producing dairy cattle. J. Dairy Sci. 95(3):1041-1056.
Zhang, Y. H. P. and L. R. Lynd. 2004. Toward and aggregated understanding of enzymatic
hydrolysis of cellulose: Noncomplexed cellulose systems. Biotechnol. Bioeng. 88:797-
824.
Zhang, Y. H. P., M. E. Himmel, and J. R. Mielenz. 2006. Outlook for cellulose improvement:
Screening and selection strategies. Biotechnol. Adv. 24(5):452-481.
Zheng, W., D. J. Schingoethe, G. A. Stegeman, A. R. Hippen, and R. J. Treacher. 2000.
Determination of when during the lactation cycle to start feeding a cellulase and xylanase
enzyme mixture to dairy cows. J. Dairy Sci. 83:2319-2325
Zhou, Q., X. L., X. Zhang, X. Meng, G. Chen, W. Liu, S. 2013. Evaluation of swollenin from
Trichoderma pseudokoningii as a potential synergistic factor in the enzymatic hydrolysis
of cellulose with low cellulase loadings. World J. Microb. Biotechnol. 27(8):1905-1910.
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BIOGRAPHICAL SKETCH
Juan Jose Romero was born in Lima, Peru in 1984. He graduated from San Agustin High
School in 2001 and his favorite courses were biology, chemistry and history. Both of his parents
were Crop Science professors in the College of Agriculture at Universidad Nacional Agraria La
Molina, Peru. Therefore, he was exposed to the agricultural challenges in Peru at a very early age
as well as how research can be used to devise solutions to such challenges. He realized that there
was a big gap between the modern agriculture industry on the coast of Peru and the agriculture of
subsistence in the Andes and he promised himself that he would endeavor to improve agriculture
in the Andes in the future. Though his parents were Crop scientists, Juan preferred cows and
llamas, so he enrolled in the Animal Science program at Universidad Nacional Agraria La
Molina for his B.S. degree. During his studies, he came across a paper titled ‘Effect of alkali
pretreatment of wheat straw on the efficacy of exogenous fibrolytic enzymes’ and realized that
enzyme treatment may be an effective strategy to increase the digestibility and utilization of the
abundant supplies of straw in the Andes. He conducted his undergraduate thesis research on this
subject and won the 2007 Alltech Young Animal Scientist Contest for Latin America for an
essay on this subject and later graduated with a B.S. in 2007, finishing at the top of his Animal
Science class. Soon after his graduation, he was admitted to UF to pursue an MS degree in
Animal Science under the supervision of Dr. Bill Brown. The fact that he was going to work on
improving forage quality was particularly exciting to him. After Dr. Brown moved to the
University of Tennessee, he was supervised by Dr. Adesogan until he completed his M.S.
program. He continued his Ph. D. program under Dr. Adesogan’s guidance working on using
fibrolytic enzymes to improve the digestibility of bermudagrass and the performance of lactating
dairy cows fed bermudagrass-based rations. His time at UF has been productive and sometimes
hard, but the training and knowledge he acquired will serve as the foundation of his future career.
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He wants to become a well-rounded ruminant nutritionist. His ultimate goal is to go back to Peru
and help to develop the Andean community and he has the knowledge, the expertise, the skills
and the drive to achieve this goal.