The Influence of Beta-Mannan and Xylose on Energy Metabolism in the Pig Nichole F

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2018 The influence of beta-mannan and xylose on energy metabolism in the pig Nichole F. Huntley Iowa State University

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The influence of -mannan and xylose on energy metabolism in the pig

Nichole F. Huntley

A dissertation submitted to the graduate faculty

in partial fulfillment of the requirements for the degree of

DOCTOR OF PHILOSOPHY

Major: Animal Science

Program of Study Committee: John F. Patience, Major Professor Cheryl L. Morris Brian J. Kerr Howard D. Tyler Anna K. Johnson

The student author, whose presentation of the scholarship herein was approved by the program of study committee, is solely responsible for the content of this dissertation. The Graduate College will ensure this dissertation is globally accessible and will not permit alterations after a degree is conferred.

Iowa State University

Ames, Iowa

2018

TABLE OF CONTENTS

Page

LIST OF FIGURES ...... iv

LIST OF TABLES ...... v

ACKNOWLEDGMENTS ...... vii

ABSTRACT………………………………...... ix

CHAPTER 1 LITERATURE REVIEW: THE INFLUENCE OF CARBOHYDRATES ON ENERGY METABOLISM IN THE PIG...... 1

Introduction ...... 1 Arabinoxylans and -mannans – fiber in swine diets ...... 2 Carbohydrases in swine diets ...... 7 Impact of mannose and xylose on energy metabolism in the pig ...... 13 Summary and conclusions ...... 39 Literature cited ...... 40

CHAPTER 2 LIPOPOLYSACCHARIDE IMMUNE STIMULATION BUT NOT -MANNANASE SUPPLEMENTATION AFFECTS MAINTENANCE ENERGY REQUIREMENTS IN YOUNG WEANED PIGS ...... 61

Abstract ...... 61 Introduction ...... 63 Materials and methods ...... 65 Results ...... 74 Discussion ...... 77 Conclusions ...... 87 Literature cited ...... 88

CHAPTER 3 BETA-MANNANASE SUPPLEMENTATION EFFECTS ON NURSERY PIG GROWTH PERFORMANCE AND SERUM ACUTE PHASE PROTEIN CONCENTRATIONS ...... 106

Abstract ...... 106 Introduction ...... 107 Materials and methods ...... 109 Results ...... 111 Discussion ...... 112

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Conclusions ...... 116 Literature cited ...... 117

CHAPTER 4 XYLOSE METABOLISM IN THE PIG ...... 128

Abstract ...... 128 Introduction ...... 129 Materials and methods ...... 130 Results ...... 138 Discussion ...... 143 Conclusions ...... 152 Literature cited ...... 153

CHAPTER 5 INTEGRATIVE SUMMARY ...... 168

Literature cited ...... 172

LIST OF FIGURES

Page

Figure 1 -1,4-D-arabinoxylan...... 57

Figure 2 General structures of plant β-mannan ...... 58

Figure 3 Arabinoxylan degrading enzymes...... 59

Figure 4 Mannose binding lectin (MBL) structure ...... 59

Figure 5 D-xylose ...... 60

Figure 6 D-xylonic acid ...... 60

Figure 7 D-xylitol ...... 60

Figure 8 D-threitol ...... 60

Figure 1 Effect of treatment on young weaned pig (n = 10 per treatment) rectal temperature (C) ...... 103

Figure 2 Effect of treatment on complete blood count before (d 8) and after (d 10) challenge ...... 104

Figure 3 Effect of treatment on serum cytokine concentrations before (d 8) and after (d 10) challenge ...... 105

Figure 1 Effects of dietary mannan level and -mannanase supplementation on nursery pig serum haptoglobin concentration (휇g/ml) at baseline (d 0) and after 28 d on the experimental diet ...... 126

Figure 2 Effects of dietary mannan level and -mannanase supplementation on nursery pig serum C-reactive protein concentration (휇g/ml) at baseline (d 0) and after 28 d on the experimental diet ...... 127

Figure 1 Two-way heat map visualization and hierarchical clustering of urine metabolite distribution ...... 165

Figure 2 Proposed xylose metabolic pathway ...... 166

Figure 3 Allocation of urine gross energy (GE) from xylose, metabolites, and nitrogen (N)-containing compounds ...... 167

LIST OF TABLES

Page

Table 1 Arabinoxylan concentration in feedstuffs ...... 56

Table 2 β-mannan concentration in typical swine diet ingredients ...... 56

Table 1 Summary of experimental treatments ...... 95

Table 2 Experimental diet ingredient and analyzed nutrient composition (as- fed basis) ...... 96

Table 3 Complete blood count values in young weaned pigs fed a diet with or without -mannanase...... 97

Table 4 Effect of -mannanase on pig serum glucose, insulin, acute phase protein, and cytokine concentrations ...... 98

Table 5 Apparent total tract digestibility in pigs on the control, enzyme, or immune system stimulation treatment ...... 99

Table 6 Growth performance and nitrogen balance in pigs on control, enzyme, or immune system stimulation treatment ...... 100

Table 7 Effect of treatment on energy balance, respiratory quotient, maintenance energy requirements, and nutrient deposition ...... 101

Table 8 Dietary energy values and efficiency in pigs on control, enzyme, or immune system stimulation treatment ...... 102

Table 1 Ingredient composition (as-fed basis) of the low mannan diets for phases 1 - 3 ...... 120

Table 2 Ingredient composition (as-fed basis) of the high mannan diets for phases 1 - 3 ...... 121

Table 3 Phase 1 diet formulated and analyzed energy and nutrient composition (as-fed basis) ...... 122

Table 4 Phase 2 diet formulated and analyzed energy and nutrient composition (as-fed basis) ...... 123

Table 5 Phase 3 diet formulated and analyzed energy and nutrient composition (as-fed basis) ...... 124

Table 6 Effects of dietary mannan level and -mannanase supplementation on nursery pig BW, growth rate, feed intake, and efficiency ...... 125

Table 1 Diet ingredient composition, as-fed ...... 158

Table 2 Formulated and analyzed diet energy and nutrient composition (as-fed) ...... 159

Table 3 Effect of dietary xylose concentration on pig growth performance ...... 159

Table 4 Effect of dietary xylose concentration and collection period on water balance ...... 160

Table 5 Effect of dietary xylose concentration and collection period on diet digestibility and energy value and on nitrogen balance ...... 161

Table 6 Effect of dietary xylose inclusion on cecum and colon digesta pH and SCFA concentration and molar proportions ...... 162

Table 7 Effect of dietary xylose concentration and collection period on urine xylose and metabolite excretion ...... 163

Table 8 Effect of dietary xylose concentration and collection period on urine energy concentration and urine sugar concentration and excretion ...... 164

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ACKNOWLEDGMENTS

I would first like to acknowledge my major professor, Dr. John Patience, for taking a chance on a comparative nutritionist with no swine experience. Thank you for the opportunities to challenge myself and grow as a scientist. Your mentorship, leadership, and encouragement have been invaluable, and I sincerely appreciate your endless support throughout my Ph.D. program.

I would like to thank the members of my committee, Dr. Cheryl Morris, Dr. Brian

Kerr, Dr. Anna Johnson, and Dr. Howard Tyler, for their guidance and support. I appreciate and benefited from your diverse expertise. My thanks go to Dr. Morris for initially introducing me to Dr. Patience and initiating not only my wonderful experience at Iowa State University but also my interest and career in comparative nutrition. I have no idea where I would be if you hadn’t taken the time and interest in introducing me to the world of zoo nutrition as a high school zoo academy student! I would also like to extend my gratitude to Dr. Steven Lonergan, Dr. Elisabeth Huff-Lonergan, Matt Schulte, and Dr. James Koltes for their collaboration and training on analyses novel to the

Patience lab.

Appreciation is expressed to Elanco Animal Health and the Iowa Pork Producers

Association for financial support for this research and my graduate program.

Appreciation is also expressed to Ajinomoto Heartland, DSM, and Danisco for their in- kind contributions.

Next, I would like to thank the past and present members of the applied swine nutrition lab group, our exceptional undergraduate students, other graduate student colleagues, and the animal science department faculty and staff for making my time at

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Iowa State University a wonderful experience. I look forward to working with each of you throughout our careers. I especially want to offer my appreciation to Stacie Gould, without your organization, hard work, and assistance during these experiments, this publication would be impossible.

Finally, thanks to my family and friends for their encouragement and constant support. Mom and Dad, thank you for instilling in me the knowledge that I really can do anything I set my mind to. Jenny, thank you for always being there for me. Most importantly, my deepest appreciation and love goes to my husband, Matt Huntley. Thank you for being my partner in life and supporting me to achieve all of my ambitions. You always know how to make me smile and life is so fun with you by my side.

ABSTRACT

Feed ingredient economics are pushing pork producers to increase the use of higher-fiber coproducts while maintaining high production and efficiency goals. To do this, nutritionists utilize -mannanase and xylanase enzymes. It is important to evaluate the metabolic impacts of -mannan and xylose to accurately estimate enzyme effects on dietary energy availability. Therefore, the overall objective of the work presented in this dissertation was to evaluate the influence of -mannan and xylose on energy metabolism in the pig. -Mannanase is added to swine diets with the objective of inhibiting an energetically-expensive -mannan-induced immune response; however, this rationale was not supported by the experiments presented in Chapters 2 and 3. Research presented in Chapter 2 showed that -mannanase did not affect immune status, nutrient digestibility, growth performance, energy balance, or maintenance energy requirements

(MEm) of young pigs. These conclusions were further supported by a nursery growth trial that tested the interactions of dietary -mannan concentration and -mannanase inclusion, presented in Chapter 3. In Chapter 2 it was also found that a lipopolysaccharide-induced innate immune challenge elevated pigs’ MEm by 23.3% which lipid deposition by 30.2% and lead to an 18.3% decrease in ADG during the immune challenge. These novel data directly related decreased ADG to increased MEm independent of changes in feed intake in immune challenged pigs. Xylose metabolism in the pig was also evaluated and presented in Chapter 4. An improved understanding of xylose metabolism in the pig and its energetic value is essential for effective xylanase utilization. Our data showed that the pig can utilize xylose but does so less efficiently as more xylose is consumed; and only 40 – 60% of xylose was retained. Furthermore, pigs

x can adapt over time to improve xylose utilization. By applying metabolomic approaches, urinary metabolites of xylose were identified and quantified. This information facilitated construction of a comprehensive pathway for xylose metabolism in the pig. Overall, research presented in this dissertation improved our understanding of how -mannanase and xylanase impact pig energy metabolism. This research also furthered the understanding of how immune activation repartitions energy and increases maintenance energy requirements.

CHAPTER I

LITERATURE REVIEW: THE INFLUENCE OF CARBOHYDRATES ON

ENERGY METABOLISM IN THE PIG

Introduction

Plant carbohydrates represent the largest energy source in commercial pig diets, comprising 60 – 75% of total energy intake. A portion of the carbohydrates (dietary fiber) cannot be digested by endogenous enzymes in the small intestine and are partially fermented in the hindgut. Dietary fiber is a chemically complicated classification of carbohydrates possessing a wide range of physiological properties. These properties profoundly influence digestive physiology and overall energetics and must be considered to fully understand the value of high-fiber ingredients in pig diets. As industry economics continues to favor increasing inclusion of low-cost, high-fiber ingredients, the complexity of dietary carbohydrate components increases and the swine nutritionists’ role of assigning accurate energy values to these ingredients is a challenge.

Swine nutritionists are beginning to understand the influence of different fiber types on diet digestibility and intestinal physiology and have developed and implemented technologies such as exogenous enzymes to improve the utilization of high-fiber ingredients. However, the systemic effects of fiber components on energy balance are still poorly understood. Xylose and mannose are two of the primary monosaccharide building blocks of grain and legume polysaccharides and their influences on swine energy metabolism are poorly understood. Therefore, the objectives of this review are to provide an overview of mannose and xylose relevance in current

swine diets, their specific metabolic impacts as well as more comprehensive influences on overall physiology, and to outline the need to understand how dietary xylose and mannose impact energy metabolism in pigs.

Arabinoxylans and -mannans - fiber in swine diets

Dietary fiber is the fraction of carbohydrates not digested by endogenous enzymes secreted into or present in the small intestine. These includes non-starch polysaccharides

(NSP), resistant starch, oligosaccharides, and the non-carbohydrate polyphenolic lignin

[1]. Non-starch polysaccharides are the largest component of dietary fiber in swine diets and so for the purposes of this review, discussion will focus on dietary NSP and its components. The main NSP in cereals are arabinoxylans, mixed linkage -(1,3 and 1,4)- glucans (-glucans), and cellulose. Cellulose consists of long, highly structured chains of

-1,4-linked glucose molecules. Unlike cellulose, hemicellulose is chemically complex.

Hemicellulose is comprised of D-xylose, D-mannose, D-galactose, D-glucose, L- arabinose, 4-O-methylglucuonic acid, D-galacturonic acid, and glucuronic acid [2]. These sugars are linked together in a wide variety of combinations, branch types, and degrees of polymerization to form hemicellulose.

Arabinoxylans

Arabinoxylans are the main NSP component in cereal grains (Table 1.1), followed by cellulose and -glucan [1], and are composed of a linear backbone of -1,4-linked-D- xylopyranosyl residues mainly substituted with -L-arabinofuranosyl residues to varying degrees at the O-2 position, the O-3 position, or both [3,4] (Fig. 1).

-Mannans

β-Mannans are components of hemicellulose in the plant cell wall and are linear polysaccharides comprised of repeating β-1,4-linked D-mannose units with galactose and/or glucose substitutions [5]. Plant mannan polysaccharides are divided into three structural subgroups; the glucomannans, galactomannans, and galactoglucomannans (Fig.

2). Galactomannans are the most common type found in swine and poultry diets. In galactomannans, single α-1,6-linked D-galactose residues are attached to some of the mannose residues within the β-1,4-mannan backbone [6].

Galactomannans are mainly found in the endosperm of legume plant seeds such as soybeans [5] where they function in water retention and are the main storage carbohydrate [6,7]. In North American swine diets, -mannans are primarily found as galactomannans in soybean meal; but around the world, other sources include palm kernel meal, copra meal, and guar meal (Table 2).

Fiber effects on diet digestibility and pig growth performance

The pig industry is known for its ability to evolve and improve throughout ever- changing economic and social environments. Currently, feed ingredient economics are pushing producers to increase the use of higher-fiber coproducts while maintaining high production and efficiency goals. To do this, nutritionists must have a comprehensive understanding of fiber utilization in the pig as well as how fiber inclusion may affect digestion and absorption of other nutrients. Unlike starch and amino acids, dietary fiber is indigestible by mammalian endogenous enzymes and its contents are highly variable among sources. Fiber’s digestive fate and impact on gut physiology depend on

physicochemical properties such as solubility, viscosity, physical structure, and fermentability [8,9].

In general, increased fiber in diets decreases diet energy density and digestibility, resulting in impaired feed efficiency and potentially growth performance if pigs are not able to increase feed intake to maintain energy intake. Increasing dietary fiber decreases dry matter (DM) and energy digestibility and potentially amino acid and lipid digestibility, with varying magnitude according to the level and type, or source, of fiber

[9,10]. Cellulose, lignin, and insoluble arabinoxylans, such as that from corn and its coproducts, have limited fermentability resulting in increased fecal bulk and decreased

DM digestibility due to the trapping of starch and protein within the fiber matrix [11,12].

Soluble dietary fiber, such as in soybean hulls and oats, increases small intestine digesta viscosity which delays gastric emptying and reduces mixing of dietary components with endogenous digestive enzymes [13]. Both soluble and insoluble fiber sources may reduce intestinal transit time [14,15] and increase epithelial cell proliferation rate [16].

Fiber inclusion in swine diets dilutes dietary energy concentration; thus, greater intake is required to achieve similar growth rates and feed efficiency is decreased [17,18].

Based on analysis of 78 diets with varied fiber inclusion, Bach Knudsen et al. [19] concluded fiber has the largest impact on organic matter digestibility and the smallest impact on lipid and starch digestibility throughout the small and large intestines. Energy digestibility declines linearly with fiber content and the decline extent is due to the specific fiber source fermentability [18,20].

Fiber fermentation is substantially more variable than starch digestion, crude protein, and fat (generally greater than 80%) [19]. In growing pigs, fiber digestibility

ranges from 15% for wheat straw, 44% for wheat bran, 47% for corn distillers’ dried grains with solubles (DDGS), to 79% for soybean hulls [21,22] and depends on viscosity and solubility properties [11,14]. Furthermore, fiber digestibility may be influenced by other nutrient concentrations, such as lipids. Gutierrez et al. [23] reported an interaction between soybean oil and reduced-oil DDGS inclusion on apparent ileal and total tract digestibility of neutral detergent fiber (NDF). It was hypothesized that increased dietary ether extract (EE) concentration increased digesta retention time [23], which may increase fiber fermentability due to longer exposure of substrates to the intestinal microbiota at the terminal ileum or in the hindgut [20,22,24]. Future research may provide insight into similar interactions with other nutrients.

Fiber decreases dietary energy concentration regardless of the physiological stage of the pig; however, the pig’s ability to utilize energy from fiber improves with increased physiological maturity [25]. Fiber digestibility increased with physiological maturity and the digestible energy (DE) value of fiber was on average 0.14 Mcal/kg DM higher in adult sows than in growing pigs [18]. This difference between sows and growing pigs is not constant but increases with fiber content (0.8 kcal for each gram of dietary NDF increase) due to larger volume hindguts and longer retention times [18,25]. The notion of increasing DE and net energy (NE) value of fibrous ingredients when fed to higher bodyweight pigs and sows is supported by numerous studies [26–30].

Crude protein (CP) digestibility is also negatively associated with fiber inclusion

[19,23,31]. This effect is more pronounced with soluble fiber compared to insoluble fiber

[32]. Fiber may increase endogenous nitrogen losses depending on concentration [33], type [34], and physicochemical properties such as water-holding capacity [35]. High

digesta viscosity may increase epithelial cell proliferation leading to lower apparent ileal amino acid digestibility [20,35]. Much of fiber’s negative effect on total tract CP digestibility derives from increased hindgut fermentation and nitrogen sequestration in microbial protein [36,37]. The inclusion of fermentable fiber shifts nitrogen excretions from urea in urine to bacterial protein in feces without affecting total nitrogen excretion or retention [24,37].

A similar effect on microbial lipid synthesis may influence observed fiber effects on lipid digestibility [30,38]. Increased dietary fiber concentration has been associated with decreased fecal digestibility of EE [24,39]; however, in an analysis of 78 diets [19] it was concluded that fiber had a small impact on lipid digestibility throughout the small and large intestine. Gutiérrez et al. [23] found that at low dietary EE levels (2% soybean oil) increased DDGS inclusion deceased EE apparent ileal and total tract digestibility, but the same effect was not observed at higher dietary EE levels (6% soybean oil). Fiber’s negative influence on lipid digestibility may be related to solubility characteristics. When insoluble fiber is provided in diets, little to no effect on ether extract digestibility is observed [40,41].

While much variation exists in dietary fiber literature, it can be concluded that increasing fiber concentrations reduces diet energy density and DM digestibility resulting in impaired feed efficiency and potentially growth performance. Continued research will provide a more comprehensive understanding of fiber utilization in the pig, its interactions with other nutrients, and improvement in feeding strategies and technologies to achieve pork production goals.

Carbohydrases in swine diets

Because animals do not possess the endogenous enzymatic capability to digest - linked polysaccharides (dietary fiber), exogenous carbohydrase enzymes, such as xylanase and -mannanase, have been developed and employed to mitigate the negative effects of dietary fiber on feed efficiency.

The most commonly discussed benefits of NSP hydrolysis due to carbohydrase supplementation are the release of cell wall-bound nutrients for absorption and a reduced ability of the polysaccharides to form viscous solutions which inhibit nutrient diffusion and absorption [42–44]. An often overlooked benefit is the potential to utilize enzyme hydrolysis products, the oligosaccharides and monosaccharides [45]. While some studies in growing pigs supplemented with NSP-hydrolases show improved nutrient digestibility, this is not always reflected in enhanced growth performance. Responses depend on the nutrient composition of the diet [46], age of the pig [47], health status, and other factors.

Although the use of carbohydrase enzymes has not yet been fully refined or applied in a targeted manner, the potential value of this technology is substantial.

Carbohydrases have the potential to improve feed efficiency and nutrient digestibility, allow greater flexibility in types of ingredients available to feed, decrease the negative impact of swine production on the environment thus, improve the profitability and sustainability of swine operations by decreasing excretion of undigested nitrogen and phosphorus in manure, and to improve gut health through production of prebiotic oligosaccharides [8,44,48,49].

-Mannanase

Endo-β-mannanases are hydrolase enzymes that cleave internal glycosidic bonds of the mannan backbone, producing β-1,4-manno-oligosaccharides [6,50]. β-Mannanase is supplemented into swine diets that have high inclusion of β-mannan containing ingredients such as soybean meal, palm kernel meal, or copra meal. This enzyme was developed based on the hypothesis that dietary -mannans resemble carbohydrate structures found on the surface of pathogenic bacteria and therefore might induce an innate immune response.

This “feed-induced immune response” hypothesis will be discussed in detail in a later section of this review; for now, it is sufficient to understand that stimulating the immune system is an energetically expensive process. It is hypothesized that after - mannanase supplementation, the hydrolyzed manno-oligosaccharides no longer retain the degree of polymerization necessary to stimulate and cross-link multiple mannose receptors, thus reducing innate immune stimulation. However, this phenomenon has yet to be demonstrated in pigs.

Xylanase

Xylanase catalyzes the hydrolysis of β-1,4-D-xylosidic linkages in the indigestible arabinoxylan constituents of hemicellulose. Xylanase is supplemented into swine diets containing wheat and barley which have more soluble and viscous arabinoxylans [51]. Xylanase is also used in diets including higher concentrations of grain coproducts, such as wheat middlings and corn DDGS, which contain the residual seed coats or bran of the parent grain and therefore have 2 to 3 times greater NSP concentrations [11,52].

Due to the heterogeneity of arabinoxylan, its complete hydrolysis requires the action of a complex enzyme system including the interaction of main chain- and side chain-cleaving enzymes (Fig. 3) [2]. The enzymes which cleave the main chain, or xylan backbone, are endo--1,4-xylanases (GH10 and 11 families, EC 3.2.1.8; what is referred to by “xylanase”) and -1,4-xylosidases (GH3 family, EC 3.2.1.37) [53]. The primary side chain-cleaving enzymes include -L-arabinofuranosidases and -D-glucuronidases

[2]. Endoxylanases release a range of products after hydrolysis of xylan such as xylose, xylobiose, xylotriose, and further polymerized xylo-oligosaccharides [2,54,55]. Beta-1,4- xylosidases cleave chain-ending xylose monomers preferentially from the non-reducing end of the substrates, with some xylosidases able to hydrolyze xylobiose [2].

Carbohydrase mechanisms of action

Viscosity

In the intestinal lumen, soluble NSP can create viscous solutions by aggregating into large networks. So, if exogenous carbohydrases degrade the highly polymerized structures into shorter oligosaccharides, they can no longer associate in large entanglements and the viscosity of the digesta is decreased [56]. Viscosity affects nutrient digestibility in poultry; however, few data are available to support this claim for swine.

Few studies have reported results of enzyme supplementation on intestinal viscosity. Of those that have, some reported reduced intestinal viscosity with xylanase supplementation

[57,58] while others reported increased viscosity [59,60]; yet, all resulted in improved nutrient digestibility. Other studies found no effect of enzyme supplementation on intestinal viscosity [60,61]. From the available data, intestinal viscosity is not likely to be a significant mode of action of carbohydrases in pigs.

Entrapped nutrient release

Plant cell wall structures (i.e. fiber) are a matrix of NSP, lignin, and other soluble nutrients such as starch and amino acids. The idea behind this mode of action is that these soluble nutrients are “trapped” within the fiber matrix which impedes digestive enzyme access and limits their availability for absorption. As carbohydrases hydrolyze and depolymerize the NSP structures, the entrapped nutrients are released and become available for absorption. This is a plausible mode of action supported by data showing increased apparent ileal digestibility (AID) of nutrients other than NSP [62–67].

However, improvements in AID are not consistent and do not always translate into improved growth performance [59,63,68].

Impact on intestinal microbial populations

As carbohydrases hydrolyze the highly polymerized NSP, oligosaccharides are produced which may have a prebiotic effect in the distal ileum, cecum, and colon and thus influence hindgut microbial populations [53,69,70]. Carbohydrase supplementation alters hindgut microbial population abundance and composition with increases in

Lactobacillus sp. and decreases in Escherichia coli [71]. Often, the implication of these changes are difficult to interpret or no effect of enzyme supplementation is reported

[72,73]. Performance benefits may derive from the provision of any of the manno-, gluco-, or xylo-oligosaccharides, but consistency of the response may be more reliant on delivery of the correct quantity of a specific oligosaccharide rather than the variable delivery of multiple oligosaccharides [74].

In the same way, carbohydrase supplementation likely changes the composition of substrates reaching the hindgut which may influence hindgut environment. In addition to

oligosaccharide production, carbohydrases may be increasing fermentability of the fiber; thus, increasing the amount of energy obtained from short chain fatty acids (SCFA). This hypothesis is supported by improvements in total tract fiber digestibility, with increases cecal and colonic SCFA concentrations and decreases in luminal pH [43,60]. However, this response is variable and depends on the enzyme and fiber composition, solubility, and fermentability [60,69,75,76].

Together, carbohydrases’ influences on microbial populations, decreases in luminal pH, and increases in butyrate production may improve gut health. This idea is primarily based on the theoretical understanding that decreased luminal pH may inhibit pathogen colonization and that butyrate may improve colonocyte health [77,78]. Few data are available which directly evaluate this carbohydrase mode of action. Preliminary data in our laboratory showed that supplementation of an enzyme blend decreased intestinal permeability in nursery pigs, demonstrated by a decrease in the ratio of lactulose: mannitol absorption and increases in mRNA abundance of tight junction proteins [76].

Energy sparing effect

As previously discussed in the -mannanase section, this enzyme was developed to break down dietary -mannans which are hypothesized to stimulate the immune system. The energy once necessary for induction and maintenance of the immune response can then theoretically be repartitioned toward other biological functions such as growth. There is evidence in poultry to suggest that diets high in -mannan content may induce immune stress [79]. Poultry fed high -mannan diets had elevated serum acute phase protein concentrations and this effect was ameliorated by -mannanase

supplementation [79]. However, in pigs, growth and efficiency responses due to - mannanase supplementation are much more variable than they are in poultry and data regarding its effects on immune stimulation in pigs are scarce. In North American swine diets, -mannans are primarily found in soybean meal. Due to the ubiquitous inclusion of soybean meal in the diets for all stages of pig production, it is imperative to understand - mannans’ impact on the immune system and energy metabolism in pigs and how - mannanase affects these responses.

Monosaccharide release

As NSPs are hydrolyzed by exogenous carbohydrases, the degree of polymerization is decreased. The types of products released from NSP hydrolysis depend on the polysaccharide’s chemical composition and structure (i.e. branching and degree of polymerization) and the type of exogenous enzyme(s) supplemented [53,54,80]. Initial arabinoxylan hydrolysis by endo-xylanase enzymes releases arabino- and xylo- oligosaccharides, with continued hydrolysis and additional enzyme activity required for the release of monosaccharides [56]. There is a paucity of data on the quantification of xylose release from arabinoxylans with xylanase supplementation. In vitro data suggest that it is possible, but not highly effective, with the combined action of endo-xylanases and xylosidases [80]. Therefore, as carbohydrase efficacy improves, small but impactful quantities of xylose, arabinose, and glucose monosaccharides will be released and absorbed in the small intestine.

Glucose and other hexose sugars contribute energy to the pig with much greater efficiency when absorbed in the small intestine than through fermentation in the large intestine, but this may not be the case for pentose sugars such as xylose and arabinose. It

is commonly assumed that these sugars are poorly metabolized in the pig and contribute little metabolizable energy (ME) if absorbed in the small intestine [81,82]. As interest in xylanase supplementation continues to increase, an improved understanding of xylose metabolism in the pig and its energetic value is essential for effective xylanase utilization.

Therefore, an objective of this dissertation research was to evaluate how xylose, if enzymatically released from arabinoxylan in the small intestine, would impact pig energy metabolism. A review of previously available literature on xylose metabolism will be presented later in this review.

Impact of mannose and xylose on energy metabolism in the pig

The energy value of dietary carbohydrates depends on the location of digestion in the gastrointestinal tract. In the small intestine, starch is efficiently digested by endogenous pancreatic and brush border enzymes and is absorbed into the portal vein for utilization throughout the body. Carbohydrates not digested and absorbed in the small intestine enter the cecum and large intestine where they can be fermented by the microbial populations and SCFA are produced. The SCFA are absorbed and can be utilized for energy by the intestinal epithelium, liver, and other tissues. Compared to enzymatic digestion in the small intestine, a smaller portion of the carbohydrate’s potential energy is available to the pig as SCFA due to losses of efficiency through the fermentation process [83].

It is important to remember that fiber fermentation does not occur solely in the hindgut. Forty to ninety-five percent of the non-digestible oligosaccharides and 20 to

25% of the NSP are degraded before the end of the small intestine [84]. In addition,

microbes in the small intestine also modify the size and structure of NSP [85–87].

Through this initial modification in the small intestine, the NSP are then provided to the cecum and large intestine in a form that is more readily available for fermentation [84].

With arabinoxylans, the section of the intestinal tract in which they are fermented depends upon their solubility and degree of substitution. The greater the degree of arabinose substitution of the xylan backbone and the lower the solubility of the polymer, the slower the rate of fermentation. The same NSP could have significant and varied effects throughout the intestinal tract depending on where it is fermented. This means that exogenous carbohydrase supplementation likely plays a similar role, shifting the site of the majority of NSP degradation more proximally into the small intestine, perhaps liberating some monosaccharides to be available for absorption before they are fermented. Therefore, to accurately determine the NE value of mannose and xylose to the pig, knowing the site of absorption and the form in which it provides energy

(monosaccharide vs. SCFA) is key.

Energy efficiency of NSP fermentation

The energy available to the pig due to fermentation is not inconsequential.

Depending on the diet and age of the animal, SCFA arising from fiber fermentation can account for 7 to 25% of the pig’s available NE for maintenance [88–91]. Through fermentation, only about 60 – 75% of the potential energy of NSP is captured as SCFA with the remaining energy lost as the heat of fermentation, microbial mass, methane, and

CO2 [88,92,93].

Short chain fatty acids are efficiently absorbed across the intestinal epithelium

[88,94] and the absorption mechanism depends on the SCFA ionization form. Generally,

SCFA are thought to be absorbed through passive diffusion across the epithelium, but this only occurs for the undissociated form of SCFA (ex. acetic acid). With a pKa of about

4.8 and a luminal pH of approximately 6, only a very small proportion of the SCFA exist in the undissociated form [94]. So, it is more probable that active transport of the ionized

SCFA (ex. acetate) is the primary absorption mechanism [95]. Short chain fatty acid anion transport can be coupled to bicarbonate secretion into the lumen or can occur through members of the monocarboxylate transporter family [94].

Short chain fatty acids are partially metabolized by epithelial cells during the process of absorption and transportation to the bloodstream. Little butyrate ever makes it to portal vein circulation as it is a preferred energy substrate for colonic epithelial cells

[88,95]. Propionate is largely metabolized by the liver, entering the TCA cycle through oxaloacetate and acetate is mostly metabolized in peripheral tissues for fatty acid synthesis [88]. Once absorbed, the energy utilization efficiency of SCFA for maintenance or gain ranges between 0.60 – 0.70, which is about 5 – 10% lower than energy supplied as starch digested and absorbed in the small intestine [18,88].

Taking into account efficiencies of digestion, conversion to SCFA, and SCFA metabolic utilization, the overall ME utilization efficiency of NSP averages 0.50 in growing pigs – 0.60 in sows [18]. In general, utilization of fermented NSP energy is equivalent to about 0.70 of enzymatically digested carbohydrates because of the additional energy costs for ingestion, digestion, fermentation, and metabolism of the intestines which are greater in high fiber diets [18]. It is important to consider these energetic efficiencies when determining NSP energy values.

Impact of mannose and -mannan on energy metabolism

If released as a monosaccharide in the small intestine, mannose (a 6-carbon sugar) can be absorbed through similar mechanisms as glucose and efficiently metabolized [96].

Within the cell, mannose is phosphorylated by hexokinase and can then be efficiently converted by phosphomannose isomerase to glucose for catabolism. A small proportion of mannose is used for N-glycosylation [96].

More commonly, -mannan remains mostly intact throughout the gastrointestinal tract but still has the potential to indirectly impact energy metabolism. As previously discussed, intact β-mannans are available to bind carbohydrate recognition domains

(CRD) of pattern recognition receptors (PRR) on innate immune cells surveying the intestinal epithelium for potential pathogens [97–99]. In this way, β-mannan is theoretically capable of stimulating innate immune cells resulting in a nonproductive, energy-draining, inflammatory immune response [79,100,101]. This effect has been termed a feed-induced immune response (FIIR) and is one of the cited reasons for supplementing swine diets with -mannanase.

The initial FIIR descriptions were in poultry and reported increased macrophage and lymphocyte proliferation [102] and plasma acute phase protein concentrations [79] due to increasing dietary soybean meal inclusion. This response was then attenuated by

-mannanase addition and was supported by improvements in feed efficiency [79,102].

This research has been interpreted to support the hypothesis that improvements in energetic efficiency due to β-mannanase supplementation are due, in part, to amelioration of a β-mannan-induced innate immune response. It was hypothesized that the hydrolyzed manno-oligosaccharides no longer retain the degree of polymerization necessary to

stimulate and cross-link multiple mannose receptors, thus reducing innate immune stimulation. However, there is a paucity of data regarding immune responses of pigs to dietary -mannan and -mannanase supplementation; and a -mannan-derived FIIR has yet to be demonstrated.

Potential mechanisms for -mannan stimulation of innate immunity

Dietary -mannans are proposed to stimulate the innate immune system through direct interactions with the carbohydrate-binding domains of mannose recognition receptors such as the membrane-bound mannose receptor and secreted mannose binding lectin. C-type lectin receptors (CLR) are a type of PRR with a central role in innate immunity through recognition of highly conserved pathogen-associated molecular patterns. This type of receptor, specifically the mannose receptor, could be responsible for inducing innate immune responses to dietary β-mannans. Mannose is frequently found on pathogen surfaces but is not commonly found in terminal positions of endogenous mammalian oligosaccharides [103].

The mannose receptor mediates responses including pathogen binding, phagocytosis, and cytokine production [98]. Downstream signaling due to mannose binding has been described [99] and seems to require coordination with other PRR molecules such as TLR2 and 4. In vitro studies have demonstrated mannan stimulation of macrophages and dendritic cells resulting in increased pro-inflammatory cytokine production [100,101,104].

Another potential mannose-detecting receptor is called the mannose binding lectin

(MBL, also called mannose binding protein; Fig. 4) which is a soluble CLR synthesized in the liver as part of the acute phase response [105,106]. Unlike typical acute phase

proteins that increase 10-1000 fold in serum concentration in response to inflammation,

MBL levels fluctuate only 2-3 fold [105,106]. Pathogens recognized by MBP include gram-negative bacteria such as Escherichia coli and fungi in the Candida and

Cyptococcus groups [103].

Mannose ligand binding by several types of C-type lectins, including MBL, is dependent upon formation of multimer complexes [107]. Individually, CRD demonstrate low carbohydrate affinity but confer broad specificity when they are complexed [103].

Thus, repeating sugar arrays decorating pathogen surfaces, or located within dietary plant-based -mannan polymers, provide potential targets for MBL binding [106]. As part of the lectin complement system of innate immunity, mannose-bound MBL can induce the complement protease cascade resulting in pathogen lysis, stimulation of phagocytosis through direct opsonization, and inflammation induction.

Impact of -mannans and -mannanase in pigs

Although β-mannan’s influence on the immune system is of great interest, commonly, only growth and energy efficiency responses to β-mannan intake are reported in swine nutrition literature. Reduced feed efficiency [108–111] and decreased growth performance [111,112] are frequently reported due to increasing dietary β-mannan concentrations. Negative growth performance effects may be explained by several factors: increased intestinal digesta viscosity [111,113,114]; exclusion of endogenous digestive enzymes from dietary substrates [6], or potentially, the FIIR [79]. For the purposes of this review, we will highlight the scarce research regarding β-mannan or - mannanase effects on immune parameters.

Dietary supplementation with galactomannan oligosaccharides in piglets was reported to increase interleukin (IL)-1 mRNA expression in peripheral blood mononuclear cells, the jejunal mucosa, and mesenteric lymph nodes [115]. Furthermore, serum IL-1 and IL-6 concentrations were increased with galactomannan oligosaccharide supplementation [115]. A prebiotic of -galactomannan was developed from carob bean and was reported to inhibit Salmonella association with intestinal epithelial cells due to attachment of salmonellae to the surface of the -galactomannan oligosaccharide [116].

In this study, -galactomannan did not induce pro-inflammatory effects; rather, the data suggested anti-inflammatory effects due to prevention of pathogen adhesion and invasion

[116].

Contrary to the immune stimulation hypothesis, dietary -mannans have also been reported to be beneficial during infection. In a Salmonella enterica in vitro infection model, β-galactomannan actually inhibited Salmonella-induced proinflammatory cytokine and chemokine expression in porcine intestinal epithelial and dendritic cell cultures [117]. Similarly, in vivo E. coli challenges have demonstrated an inhibitory effect of dietary β-mannan [118]. The authors hypothesized that the pathogens bind to the mannose residues on the dietary polysaccharides instead of onto epithelial cells, thus reducing pathogen internalization and infection [118].

Clearly, a more direct evaluation of dietary β-mannan in swine diets and its effects on immune parameters is required to truly address the FIIR hypothesis in pigs. A better understanding of β-mannan effects on the innate immune system and maintenance energy requirements of pigs will allow more appropriate and efficient use of β- mannanase to improve feed utilization in growing pigs. In addition to evaluating if, and

how, dietary -mannans impact the immune system in pigs, it is important to understand the energetic cost of inducing and maintaining an innate immune response.

The energetic cost of an immune response

Significant interplay exists between an animal’s nutritional status and immune system function. While much research has been conducted on the impact of certain nutrients or feeding strategies on an animal’s ability to respond to a pathogenic challenge, in this review we will focus on energy partitioning during an immune challenge. In immune-challenged animals, energy is diverted away from growth and toward initiating and maintaining an appropriate and efficient response to clear infection or the perceived pathogen from the body [119–121].

During the innate immune response, proinflammatory cytokines orchestrate a response resulting in fever, acute phase protein production, and neutrophil and lymphocyte proliferation, each of which increases amino acid and energy requirements

[122]. For example, IL-6 infusions in healthy human volunteers increased resting metabolic rates by 25% [123], and a core body temperature increase of 1°C has been reported to increase basal caloric requirements between 15 - 30% [124,125].

Furthermore, cytokines invoke other growth modulators such as glucocorticoids, prostaglandins, and catecholamines [121]. Together these responses generally suppress feed intake and accelerate protein degradation rates resulting in an economically costly decrease in feed efficiency [126].

Decreases in growth and feed efficiency have been well documented during disease challenges, yet data quantifying the energy requirements of an immune response are scarce. Few studies have evaluated the magnitude of change in energy balance during

an immune challenge to inform more effective dietary intervention strategies. Previous studies have evaluated increased caloric requirements with fever [127,128], but few have directly related a chronic immune challenge with increased maintenance energy requirements. In vitro studies with isolated mitochondria from rats stimulated with TNF or IL-1 showed up to 30% increases in respiration rate [129]. Demas et al. [128] showed that mice injected with a mild antigen caused limited immune activation resulting in significantly more O2 consumption that control mice injected with saline and a study in humans found that IL-6 infusions increased resting metabolic rates by 25% [123].

In pigs, however, the direct relationship between immune stimulation and increased energy requirements has not previously been demonstrated. Immune system stimulation did not impact rate or efficiency of gain, or energy balance measurements in studies by Moon et al. [130] and Williams et al. [131]. However, Moon et al. [130] reported fibroblast formation at the injection site which encapsulated the immunogen and prevented systemic delivery. Williams et al. [131] used the comparative slaughter technique and found no differences in the energetic costs of maintenance and protein and lipid deposition between pigs raised in environments encouraging high or low chronic immune activation.

Conversely, Labussière et al. [132] and Campos et al. [133] reported decreased heat production (HP) in pigs during inflammatory challenges. A single injection of complete Freund’s adjuvant in nursery pigs decreased resting and total HP in the residual feed intake (RFI) positive genetic lines, but not the RFI negative lines. However, it is important to note that the pigs were not measured for HP until the day after injection and only re-entered the calorimetry chamber after visual recovery which likely biased the

response [132]. Campos et al. [133] reported a 14% decrease in total HP in response to a chronic lipopolysaccharide (LPS) inflammatory challenge in growing pigs even though typical inflammatory-type and febrile response were observed. Decreased HP was mainly attributed to feed intake depression. In addition to evaluating the energetic costs of dietary -mannans and potential -mannanase energy sparing effects, it was clear that a better understanding of the energetic cost of pathogen innate immune stimulation was also needed.

Impact of xylose on energy metabolism

While studying the xylose literature to prepare for this dissertation work it was apparent that there was a glaring paucity of information and research available on xylose metabolism in all vertebrate species. No comprehensive review of the available literature had been published. Therefore, the following review is part of the manuscript published in the Journal of Animal Science and Biotechnology (2018, vol. 9(1): 4; doi:

10.1186/s40104-017-0226-9)

Xylose, as a major constituent of plant xylan polymers, is one of the most abundant carbohydrates on Earth, second only to glucose [134,135]. This abundant pentose sugar, along with arabinose, makes up a majority of the hemicellulose backbone as arabinoxylan in the cell walls of cereal grains fed to pigs [12]. In contrast to α-linked starch, which is hydrolyzed to glucose by endogenous enzymes, β-linkages in arabinoxylan and other NSP must be degraded by microbial enzymes. Thus, pigs cannot derive energy directly from arabinoxylans. The large proportion of arabinoxylan in the hemicellulose fraction of cereal grains and coproducts makes xylanase supplementation of great interest in swine nutrition [136,137]. While greater progress is needed to achieve

consistent enzyme efficacy, it is yet to be determined how the products of xylanase hydrolysis will be utilized by the pig and to what extent these products may contribute to overall energy balance.

Xylanase hydrolysis products have not been well studied in vivo, but an understanding of enzyme function indicates that a mix of oligosaccharides, disaccharides, and monomeric pentose sugars, such as xylose (Fig. 5), may be released through the combined actions of xylanases with various activities [53,54,138]. Typical growing pig diets will contain 3 - 5% polymeric xylose that may significantly impact the nutrition of growing pigs if released as free xylose. Yet, the current understanding of xylose metabolism and potential contribution to dietary energy is vague at best. This review examines published, peer-reviewed research on xylose absorption and metabolism in the pig and identifies gaps in this knowledge that are essential to understanding the value of carbohydrase hydrolysis products for swine nutrition. Xylose research in pigs began with initial observations by Wise et al. [139] in 1954; however, little research hass been conducted since. Therefore, this review will also discuss relevant xylose absorption, metabolism, and fermentation research in other monogastric species, including humans.

Furthermore, xylose effects on glucose and energy balance in monogastric animals are considered.

Effects of xylose on animal performance

In both pigs and poultry, increasing the dietary inclusion of D-xylose generally results in a linear decrease in ADG, feed intake, feed efficiency, and excreta DM content

[45,139–142]. Linear decreases in the apparent total tract digestibility of DM, energy, and nitrogen were reported in broilers consuming dietary D-xylose concentrations of 0, 5, and

15% [142]. Exceptions exist in poultry; Longstaff et al. [143] reported no significant differences at 5 or 20% dietary D-xylose inclusion and in pigs where ADG was not reduced when a basal diet was supplemented with 10% D-xylose compared to 5% D- glucose [81]. However, xylose generally imparts negative performance responses that, in some cases, have been severe. Two-week-old pigs fed purified diets containing 18.7 or

37.4% D-xylose had drastically decreased feed intake and ADG compared to pigs fed

37.4% D-glucose [139]. The authors also noted decreased physical activity in xylose-fed pigs along with vomiting, diarrhea, cataract formation, and nephritis in pigs fed the highest xylose diet [139].

Importantly, these drastic xylose effects on animal performance are only observed when purified D-xylose is supplemented, not with xylanase supplementation [43,62,144].

This distinction could be explained if exogenous xylanase releases only small quantities of monomeric xylose in vivo, as compared to the levels fed in the above-mentioned studies. Based on the NSP composition of fibrous ingredients fed to growing pigs, most diets rarely contain more than 5% xylose [12,136,137,145], regardless of the degree of polymerization. Furthermore, only a small proportion of the polymeric xylose would likely be hydrolyzed to its monomeric components [57]. Even if 100% of the xylose molecules are liberated through exogenous enzyme supplementation, this level is unlikely to compromise pig health. However, the minimum level of xylose required to impact pig health has not yet been defined empirically.

Xylose absorption and disappearance

In pigs, xylose has a high disappearance rate in the small intestine, ranging from

96 – 99% when supplemented as purified D-xylose [81,82,140,146]. Similarly, xylose

disappearance in poultry is very high (93.5 – 97.8%) when supplemented at 5% of diet

DM; the rate of disappearance from the gut declined as the quantity of xylose in the diet increased: 91.9% at 20% and 87.3% at 40% dietary D-xylose [140,143]. Although it is possible that a portion of the xylose disappearance is due to microbial fermentation in the small intestine, there is empirical evidence that xylose is also readily absorbed in the duodenum and proximal jejunum in rats [147,148]. This is assumed to be similar in pigs and poultry based on disappearance prior to the cecum [82,146,149,150]. However, xylose mucosal transport throughout the entire digestive tract, including the colon, has been described [149]. Providing purified D-xylose in the diet at high levels (> 20%), also decreases ileal digestibility of DM, organic matter, gross energy and nitrogen in pigs

[81,82].

Mechanisms of xylose absorption have yet to be fully characterized in pigs and poultry. Passive diffusion was initially reported [151], but most research indicates the existence of a sodium-dependent active transport system similar to that described for glucose and amino acids [82,149,150,152]. D-Xylose in rat diets induced expression of sodium-linked glucose transporter mRNA [153] suggesting at least some xylose is carried by this transporter and that competitive binding with glucose may occur. [152]. Xylose did not affect mRNA abundance of either GLUT2 or GLUT5 [153]. An active transport mechanism is further supported by data from equine and rabbit jejunal tissue which accumulated xylose against a concentration gradient when incubated in a 1 mmol/L D- xylose solution [154]. However, when the tissues were incubated in a 5 mmol/L D-xylose solution, no accumulation was reported.

Freeman’s data in horses and rabbits [154] suggest that an active transport system may exist for xylose although it has a low affinity and is easily saturated. If this system does exist in swine, the proportion of xylose that is actively transported is likely very low. This hypothesis may relate to the considerably slower rate of xylose absorption compared to glucose [148]. Furthermore, because the transportation system becomes saturated at low xylose concentrations, absorption would be expected to continue to occur through diffusion given the high xylose disappearance rate in the small intestine [154]. In humans, however, xylose is absorbed by sodium-independent passive diffusion [155,156] and no inhibitory effects of glucose have been observed [156].

Although precise mechanisms of xylose absorption and retention have not been reported for pigs and poultry, based on data in other species, it is clear that xylose is readily absorbed but at a slower rate than galactose and glucose and faster than arabinose

[45,148,157]. Like most other water-soluble monosaccharides, xylose is then probably transported from the serosal side of the enterocyte to systemic circulation via the portal vein to the liver [158]. However, transporters potentially involved in this transfer have not been described. Once xylose is absorbed into the enterocyte it must be retained.

Unlike glucose, reports on xylose phosphorylation are variable and the retention mechanism is unclear. Mucosal homogenates from rats did not demonstrate the ability to phosphorylate xylose [159]. In contrast, 13 and 36% of the xylose accumulated in jejunal tissues during D-xylose incubation was phosphorylated in the equine and rabbit tissues, respectively [154]; xylose phosphorylation following active absorption has also been reported [160].

While absorption mechanisms of xylose may not be fully revealed, its ability to decrease glucose absorption has been more clearly demonstrated. Currently, xylose is of interest in human nutrition as a supplement to suppress postprandial glucose and insulin surges. Xylose has been reported to selectively inhibit sucrase activity in a non- competitive manner [161]. When consumed with a glucose solution or high carbohydrate meal, xylose decreased serum glucose levels up to 30 min post-consumption in one study in humans [162] and up to 120 min in another [163]. Additionally, xylose supplementation decreased insulin area under the curve for up to 90 min compared to a pure glucose solution or high carbohydrate meal. [162,163].

Suppression of the glycemic effect may occur in part through sucrase inhibition as well as through stimulation of glucagon like peptide-1 (GLP-1) secretion. In humans with type 2 diabetes consumption of a 50-g D-xylose solution 40 min prior to a high carbohydrate meal attenuated the postprandial glycemic and insulin responses associated with GLP-1 secretion before the meal [164]. The authors initially postulated that GLP-1 secretion was stimulated in response to SCFA produced through bacterial fermentation of xylose, confirmed by an increase in hydrogen production in breath samples [164].

However, GLP-1 concentrations began to increase within 20 min of xylose ingestion, suggesting that more direct metabolism of the pentose was also involved [164].

Some researchers have speculated that xylose supplementation would not exert similar effects in a high-fat meal due to the typically lower glycemic response [163], yet positive responses have been demonstrated. In obese mice fed high-fat diets, 5 and 10%

D-xylose supplementation attenuated fasting blood glucose concentration back to healthy control levels, compared to mice fed a high-fat diet without xylose [165]. However, in

poultry, no effect of xylose supplementation on serum glucose concentration was reported and the responses to increasing dietary xylose concentration on serum insulin were highly variable [142]. This discrepancy may be related to the tight regulation of circulating glucose concentrations in birds [166]. Xylose effects on glucose absorption and regulation must be considered when determining how xylanase hydrolyzed monomers impact overall energy balance in pigs.

Xylose fermentation

Results of most experiments with monomeric xylose report almost complete disappearance prior to the terminal ileum; furthermore, increasing dietary xylose inclusion has been reported to increase ileal SCFA flow [82], cecal SCFA concentration

[167], and cecal weight [45], indicating some degree of microbial fermentation of xylose in the small intestine. In humans, consumption of a 50-g D-xylose solution was reported to increase breath hydrogen production, a measure of bacterial fermentation, within 40 min of consumption, and continued increasing through at least 280 min post consumption

(the last time point sampled) compared to a placebo control [164].

The importance of the microbiota to D-xylose disappearance is verified by studies with germ-free or antibiotic-treated mice and rats [168–171]. Conventional rats had

87.8% total tract D-xylose disappearance compared to 74.3% in germ-free rats, with all differences occurring in the cecum and colon [169]. Studies with wild-caught rock mice

(Aethomys namaquensis) who consume xylose containing nectar showed a 43% reduction in xylose metabolism following a heavy antibiotic regimen to drastically reduce gut microbial populations [170,171]. The xylose-metabolizing gut bacteria were identified as an inducible population with higher numbers present when xylose is present in the diet

[170]. If a xylose-metabolizing hindgut population is inducible in pigs, it is possible that the extent to which xylose is fermented may increase as pigs are adapted to diets with higher free xylose concentrations.

Xylose oligomers hydrolyzed from arabinoxylans in the small intestine by xylanase can be readily fermented by microbial populations in the distal ileum, cecum, and colon [82,167,172]. Xylose utilization was demonstrated by pig cecal cultures which fermented xylose at the same rate and curve of gas production as glucose fermentation

[167]. Xylose fermentation generally results in greater acetate and butyrate proportions compared to glucose fermentation [167,172,173]. These data indicate that the ability of pig microbiota to utilize xylose is not limited.

Xylose metabolism

Distinguishing the mechanism by which xylose disappears from the small intestine, either by absorption of the intact sugar or by microbial fermentation, is essential for determining its value to the pig in terms of energy metabolism. Very little is known about xylose metabolic utilization in vertebrates but a large majority of dietary supplemented D-xylose consistently appears in the urine [82,139,147] and significantly decreases the ME available from the diet [81,143,146,157]. For most carbohydrates, fermentation is a less efficient means of releasing dietary energy as compared with direct absorption and oxidation; however, in the case of xylose, it can be argued that fermentation may be more efficient.

In all species, urinary xylose excretion increases as dietary D-xylose inclusion increases [45,81,82,157,174]. Dietary D-xylose supplementation at 0.2% of DM resulted in 35% being excreted in the urine [146], while 10% D-xylose inclusion resulted in 37 –

45% urinary excretion [81,82], and at 20% dietary D-xylose, 52.6% was excreted in urine

[82]. The finding that the proportion of xylose excreted in the urine increases with higher intake suggests there may be some type of threshold for xylose metabolic capacity and that the energy potentially derived from xylose metabolism may change with the amount absorbed.

The detrimental effect of D-xylose on diet ME may be due to more than increased urinary xylose excretion. Verstegen et al. [81] compared ME values in pigs fed diets with

10% D-xylose or 5% D-glucose. Metabolizable energy was significantly lower in xylose diets (673.58 vs. 698.85 kcal/d) and the difference was solely due to greater energy excreted in the urine (73.63 kcal/d vs 21.03 kcal/d, for xylose and glucose, respectively).

Surprisingly, the extra urinary energy from the xylose diet was more than could be accounted for by the presence of xylose alone. Non-xylose energy in urine was double that of control pigs [81]. Similarly, differences in urinary excretion of xylose between pigs fed either 10 or 20% D-xylose were not completely reflected in urinary excretion of energy [82]. Urinary xylose metabolites, such as threitol, were not measured in these experiments but may have contributed to the energetic differences.

Heat production data determined through indirect calorimetry helps to clarify when and how xylose may be altering energy metabolism in the pig. Pigs dosed with 500 g of D-xylose had increased heat production and the majority of xylose urine excretion was on the day of dosing, with only 2 - 6 g excreted on the following day [146]. Yet, the increase in heat production persisted until the third day after xylose dosing [146]. The authors proposed that this reflected either delayed metabolism of xylose or an increase in fermentation. These data provide further evidence of the pig’s capacity to convert xylose

into energy, although the efficiency may be limited. The relative contribution of fermentation to the energy balance remains unclear.

Clearly, xylose does not contribute to ME with the same efficiency as glucose when provided as a purified monomer at high dietary concentrations (5 - 40%). It has yet to be determined if this effect holds for xylose monomers hydrolyzed in vivo from arabinoxylans resulting from inclusion of xylanase and supporting enzymes in the diet.

The concentration of xylose released would depend on dietary NSP composition and carbohydrase efficiency, which is not frequently reported. Other than quantifying its contribution to ME, experiments in pigs have not examined further metabolic effects of increased xylose absorption.

Few data are available regarding xylose metabolism in pigs, but relevant studies in other monogastric species help to clarify the potential metabolic impact of xylose.

Dietary xylose supplementation in rats increased blood glucose levels over time, but not liver glycogen [175,176]. Intravenous infusions of D-xylose solutions (10 - 20 g over 10 min) in humans also increased blood glucose concentrations and resulted in a drop in serum inorganic phosphate concentration [174]. This research seemed to indicate xylose conversion to glucose, a phenomenon not supported by current research. However, if xylose can be oxidized to CO2 for energy production or metabolized through the pentose phosphate pathway, it is possible that glucose may be spared.

Xylose oxidation to CO2 has been demonstrated in humans; infused D-xylose-1-

14 14 C appeared as expired CO2 in humans within 15 min [174]. Expired CO2 was maximally labeled 45 min following the end of D-xylose-1-14C infusion; the label was detectable for 6 h, and cumulatively represented 15.5% of the administered label [174].

This study was the first to demonstrate metabolism of xylose carbon separate from microbial fermentation.

Similarly, guinea pigs have been shown to oxidize xylose to CO2 [177]. Expired

14 CO2 was maximally labeled 75 min after intraperitoneal injection of D-xylose-1- C, with total 14C recovery representing 11.3% of the injected dose. About 50% of the xylose was excreted in the urine with the vast majority appearing within 5 h after injection [177].

Results from this thorough experiment indicate the ability of multiple tissues to absorb xylose. The distribution of 14C retained in tissues was greatest in the muscle followed by spleen, pancreas, liver, heart, and was lowest in the kidney. However, when slices of the

14 14 tissues were measured for the oxidation of D-xylose-1- C to CO2, the kidney was the most active followed by the liver, pancreas, spleen, heart, diaphragm, and lastly muscle

[177]. Kidney tissue oxidized 164 times more D-xylose-1-14C than muscle.

Furthermore, D-xylose-1-14C metabolism was compared between control, intact animals and nephrectomized guinea pigs [177]. Control animals oxidized 10.8% of the

14 injected dose to CO2 in 4 h with 41.3% excreted in urine and 47.9% unrecovered.

14 Nephrectomy increased CO2 excretion 2 to 3-fold with about 75% of the injected dose unrecovered [177]. However, the capacity may have been even greater considering that the kidneys significantly contributed to xylose oxidation; therefore, their removal may have reduced the overall capacity to oxidize xylose. In sum, these results provide evidence for xylose oxidation to CO2 in multiple tissues, with the greatest capacity in the kidney. Some absorbed D-xylose may be retained in tissues throughout the body for longer than 6 h, and the capacity for greater xylose oxidation to CO2 exists if no other means of excretion are available [177].

These data provide convincing evidence that multiple species are capable of metabolizing at least some absorbed xylose to CO2 [174,177]. Evidence thus far seems to point to critical roles of the liver and kidneys. Severe liver dysfunction had little impact in human xylose absorption tests, indicating that hepatic tissue is not significantly involved [147]. Furthermore, xylose supplementation to fasted rats did not increase liver glycogen concentration compared to fasted controls [178], indicating that the liver cannot metabolize xylose to glucose for storage as glycogen.

Current understanding of metabolic pathways in pigs and other monogastric animals support two possible routes for D-xylose metabolism. The first is oxidation by D- xylose dehydrogenase (EC 1.1.1.175) to D-xylonic acid (Fig. 6) via a D-xylonolactone intermediate. D-xylose dehydrogenase has been identified and purified in pig liver [179] as well as in monkey kidney, dog liver, and rabbit lens [179,180]. In vitro data from guinea pigs supports the presence of pentose dehydrogenase activity in liver extracts which is separate and distinct from dehydrogenases for glucose [177]. In liver extracts,

D-xylose-1-14C was oxidized to D-xylonic acid-1-14C and D-xylonic acid-1-14C injected

14 in vivo was also oxidized to CO2 [177]. These data support that xylose oxidation to CO2 can involve conversion to xylonic acid and subsequent decarboxylation [177,179,180].

A second potential pathway is the reduction of D-xylose by aldose reductase (EC

1.1.1.21) to D-xylitol (Fig. 7) which is then converted to D-xylulose and can be metabolized through the pentose phosphate pathway. Aldose reductase exhibits broad substrate specificity, including D-xylose and a variety of sugars [181]; expression of aldose reductase genes has been measured in pig lens [182], spleen, lung, ovary, adrenal, endometrium, kidney, and liver tissues [182,183]. Xylose metabolism through either of

these two potential pathways is possible; however, efficiency of the reactions depends on the distribution, abundance, and activity of D-xylose dehydrogenase and aldose reductase in pig tissues, something which has not yet been fully described.

Xylose can also be metabolized to D-threitol, a urinary metabolite (Fig. 8). It has been measured in urine following D-xylose consumption by humans [184] and pigs [81]; human data seem to indicate a role for the liver. In healthy patients dosed with D-xylose,

15% of the xylose excreted in the urine was actually recovered as D-threitol within 5 h post-dosing; this proportion increased when the collection time was extended to 24 h

[184]. Delayed excretion is not observed with xylose measured in the urine and seems to indicate a delayed conversion to and excretion of threitol. Threitol excretion was decreased in patients with cirrhotic liver disease which indicates a substantial portion of the xylose to threitol conversion may occur in the liver [184]. Furthermore, similar amounts of urinary threitol were noted after both intravenous and oral administration of xylose, supporting the hypothesis that the conversion is occurring in the liver as opposed to the intestines [184].

Although specific pathways of xylose metabolism have not yet been clarified, results from more recent studies provide information on xylose supplementation effects on metabolism of other nutrients and overall physiology. Hepatic expression of enzymes and transcription factors involved in glucose and lipid metabolism was measured in broiler chickens fed diets containing 0, 5, or 15% D-xylose [142]. The expression of phosphoenol pyruvate carboxykinase (PEPCK), a key enzyme in gluconeogenesis, did not appear to be affected by xylose supplementation at any time point up to 300 min post ingestion [142]. However, in poultry, the kidney is the major organ for gluconeogenesis

[166] and treatment differences may have been observed if kidney samples had been analyzed for this enzyme instead. In the fasted state, hepatic pyruvate carboxylase was linearly increased by xylose [142]. Because pyruvate carboxylase is a rate limiting enzyme for hepatic gluconeogenesis which is active when glycogen stores are depleted, such as occurs in starvation, this result corroborates reports that xylose depletes hepatic glycogen stores in poultry [141,157].

Understanding the effect of xylose on glucose metabolism is integral to understanding its contribution to overall energy metabolism. Potential effects must be gleaned from studies on other monogastric species as no data are available in pigs. Rats have been used as a model to evaluate potential xylose anti-diabetic effects; in one study, they were fed meals containing 5 or 10% D-xylose for 14 d [185]. In agreement with the attenuated glycemic effects discussed earlier, xylose supplementation decreased fasting blood glucose in a dose-dependent manner and decreased hepatic glycogen concentrations [185]. In contrast to the data in poultry, hepatic PEPCK protein levels were reduced in xylose supplemented rats indicating lower rates of gluconeogenesis. In vitro, xylose also increased insulin secretion from pancreatic -cells [185]. These data indicate xylose is unlikely to contribute substantially to overall energy balance in pigs; in high enough concentrations, xylose may affect postprandial glucose metabolism

[142,185].

Xylose supplementation may also affect lipid metabolism. D-Xylose supplementation at 5 and 10% of a high-fat diet reduced weight gain, improved serum lipid profiles and reduced hepatic lipid accumulation in obese mice [165]. These effects appeared to be influenced by changes in the expression of genes that mediate adipocyte

differentiation, lipogenesis, and -oxidation of fatty acids in adipose tissues. Xylose supplementation was associated with lower mRNA levels of sterol regulatory element- binding protein 1C, fatty acid synthase, adipocyte marker, and CCAAT/enhancer-binding protein- in visceral adipose tissue of mice [165]. In the liver, xylose supplementation decreased protein levels of fatty acid synthase and peroxisome proliferator-activated receptor- by at least 40% compared to the high-fat diet control group [165]. These results indicate that xylose supplementation may suppress lipogenesis and may give insight into mechanisms through which dietary xylose levels greater than 5% reduce

ADG in pigs and poultry.

In the muscle, xylose may not be metabolized directly or efficiently for energy

[177], but it does appear to at least be transported into the muscle cell [177]; and this transportation is increased in the presence of insulin [186]. Additionally, xylose may influence myocyte glucose transport. High levels of xylose were found to upregulate the glucose transport system [187] and generally increase glucose uptake by mouse myotubes

[185]. Gruzman et al. [187] reported that xylose did not utilize glucose transporters to enter myotubes but suggested the presence of a highly specific pentose transport system in skeletal muscles. These data corroborate earlier reports of xylose retention in muscle

[177], yet the transport mechanism and eventual metabolic fate remain unclear.

While the ambiguity surrounding xylose metabolism is frustrating from a nutritional perspective, it is not surprising from an evolutionary perspective. Monogastric animals do not have endogenous enzymes capable of degrading xylan to release monomeric xylose for absorption in the small intestine; consequently, there was little need to develop efficient mechanisms for xylose metabolism. Furthermore, xylose is a

rare carbohydrate in mammalian cells and so far is only found as a link between the protein and glycosaminoglycan chains of some proteoglycans and in the Notch receptor

[134]. The Notch receptor is part of a highly conserved core signaling pathway required for various cell fate decisions at multiple stages of development [188]. However, UDP- xylose, the activated precursor for xylose involvement in these structures, is synthesized from UDP-glucose and is not derived from dietary xylose sources [134]. As a result, the absence of an efficient or well-conserved pathway for xylose metabolism comes as no surprise because there has been no need. In practical terms, it also means that biochemists are not likely to find consistent mechanisms of xylose metabolism across species. Nor would nutritionists be highly motivated to evaluate the specific effects of xylose supplementation or xylose release from enzyme hydrolysis. One consistency regarding xylose metabolism is the ability of intestinal microbiota to ferment xylose and arabinoxylan. Given the inefficiency of xylose metabolism when absorbed intact by enterocytes, a critical comparison with the energy derived from xylose fermentation is necessary to determine the most energetically efficient use of dietary xylose.

Xylose conclusions

Xylose appears to be poorly utilized by monogastric animals, but xylose metabolism to CO2 is possible. Increased dietary concentrations of D-xylose linearly decreases ADG, feed intake, and feed efficiency in pigs. Even though xylose disappears almost completely from the small intestine, a high proportion of what is absorbed is excreted in the urine (35 – 50%), either as xylose or as a metabolite. Of the portion retained in the body, only a small percentage is fully oxidized to CO2. This indicates that xylose is unlikely to contribute in any significant manner to energy balance in the pig

through oxidative pathways. Its energetic contribution through fermentation may be less efficient, but quantitatively more important. However, this hypothesis has yet to be tested. More specifically, xylose inhibited sucrase activity leading to decreased postprandial blood glucose and insulin levels; further, xylose decreases expression of genes involved in lipogenesis. Combined, these effects are beneficial to humans, especially for those with diabetes, but are negative in growing pigs.

It is important to consider that only one study has been reported using dietary concentrations of D-xylose below 5% in pigs [146]. It has yet to be determined if dietary concentrations more relevant to swine nutrition (i.e. 1 - 4% xylose) would exert similar effects. It must also be determined if pigs develop increased capacity for xylose metabolism as adaptation time to diets supplemented with xylose or xylanase increases.

Data regarding xylose absorption support the hypothesis that mechanisms exist to handle low concentrations of xylose, but those mechanisms may be easily overwhelmed as luminal concentrations increase. It is possible that these mechanisms may be up-regulated as the pig adapts to higher luminal xylose concentrations. Furthermore, xylose effects on nitrogen balance and muscle metabolism have yet to be evaluated. Lastly, there is a need for a more comprehensive understanding of the action and hydrolysis products of exogenous xylanase in vivo, an understanding of how pig tissues metabolize hydrolyzed xylose monomers, and clarity on the contribution of microbial xylose fermentation to dietary xylose absorption and the energy value of xylanase supplemented diets.

Summary and conclusions

Mannose and xylose are important components of the NSP fraction of swine diets, yet as both intact polysaccharides as individual sugars, their effects on energy metabolism in the pig are poorly understood. A more direct evaluation of dietary β- mannan in swine diets and its effects on immune parameters is required to truly address the FIIR hypothesis in pigs. A better understanding of β-mannan effects on the innate immune system and maintenance energy requirements of pigs will allow more appropriate and efficient β-mannanase application to improve feed utilization in growing pigs. In addition to evaluating if, and how, dietary -mannans impact the immune system in pigs, it is also important to understand the energetic cost of inducing and maintaining an innate immune response.

Additionally, with improved xylanase efficacy, an increasing proportion of xylose will be absorbed in the small intestine. Currently, xylose metabolism is poorly understood but understanding its effects in pigs and the energetic value of xylose is essential for effective xylanase utilization. As the use of -mannanase and xylanase enzymes continues to increase, it is important to evaluate their metabolic impact in order to accurately estimate enzyme effects on dietary energy availability to the pig. This information will allow nutritionist to improve the accuracy of diet formulations when high fiber ingredients and carbohydrase enzymes are utilized.

Therefore, the overall objective of this dissertation work was to evaluate the impact of -mannan and xylose on energy metabolism in the pig. More specifically, the objective of Experiment 1 (Chapter 2) was to evaluate the -mannan-induced immune response hypothesis in pigs by testing the impacts of -mannanase supplementation and

true immune stimulation on maintenance energy requirements. As a follow up, the objective of Experiment 2 (Chapter 3) was to evaluate the effects of dietary mannan level, -mannanase supplementation, and their interaction on immune parameters, growth performance, and feed efficiency in nursery pigs. Finally, the objectives of

Experiment 3 (Chapter 4) were to evaluate the impact of dietary xylose concentration and adaptation time on xylose metabolism and its energy value to the pig.

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Table 1. Arabinoxylan concentration in feedstuffsa Feed ingredient Arabinoxylan concentration, g/kg Corn bran 221 Corn DDGs-reduced oil 143 Soybean hulls 124 Barley 84 Wheat 73 Corn 47 Soybean meal 43 Table 1.1. Arabinoxylan concentration in feedstuffsa aAdapted from [12,52,137]

Table 2. β-mannan concentration in typical swine diet ingredientsa Feed ingredient β-mannan concentration, g/kg Palm kernel meal 367 Copra meal 250 Guar meal 87 Hulled soybean meal 21 Dehulled soybean meal 13 Rye 6.1 Wheat 0.9 Corn 0.8 aAdapted from [6]

Figure 1. 훽-1,4-D-arabinoxylan. 훼-1,3 L-Arabinose residues are linked on position O3, or O2 and O3 if di-substituted, of a D-xylose unit.

A) Mannan

B) Galactomannan

C) Glucomannan

D) Galactoglucomannan

Figure 2. General structures of plant β-mannan. A) A typical mannan polymer of repeating β-(1,4)-linked D-mannose units; B) Galactomannan structure consisting of a β- (1,4)-mannan backbone with α-(1,6)-linked D-galactose residues attached to some mannose residues. C) Glucomannan structure consisting of β-(1,4)-linked D-glucose substituted within the β-(1,4)-mannan backbone. D) Galactoglucomannan structure consisting of the β-(1,4)-linked glucomannan chain with α-(1,6)-linked D-galactose residues attached to some mannose residues. Figure adapted from [7].

Figure 3. Arabinoxylan degrading enzymes. Arrows indicate hydrolysis sites of xylan degrading enzymes.

Figure 4. Mannose binding lectin (MBL) structure. The MBL consists of a monomeric polypeptide chain with a cysteine-rich N-terminal region, a collagen region, “neck region”, and a carbohydrate recognition domain (oval) at the C-terminal. MBL monomers then form trimers the trimers form multimers of trimers resulting in a “bouquet-like” conformation. Figure adapted from [189].

Figure 5 D-xylose

Figure 6 D-xylonic acid

Figure 7 D-xylitol

Figure 8 D-threitol

CHAPTER II

LIPOPOLYSACCHARIDE IMMUNE STIMULATION BUT NOT -

MANNANASE SUPPLEMENTATION AFFECTS MAINTENANCE ENERGY

REQUIREMENTS IN YOUNG WEANED PIGS

Modified from a paper published in 2018 by the Journal of Animal Science and

Biotechnology (In Press) doi: 10.1186/s40104-018-0264-y

Nichole F. Huntley1, C. Martin Nyachoti2, John F. Patience1*

1Department of Animal Science, Iowa State University, Ames, IA 50011; and

2Department of Animal Science, University of Manitoba, Winnipeg, MB R3T 2N2,

Canada

Abstract

Pathogen or diet-induced immune activation can partition energy and nutrients away from growth, but clear relationships between immune responses and the direction and magnitude of energy partitioning responses have yet to be elucidated. The objectives were to determine how β-mannanase supplementation and lipopolysaccharide (LPS) immune stimulation affect maintenance energy requirements (MEm) and to characterize immune parameters, digestibility, growth performance, and energy balance. In a randomized complete block design, 30 young weaned pigs were assigned to either the control treatment (CON; basal corn, soybean meal and soybean hulls diet), the enzyme

treatment (ENZ; basal diet + 0.056% -mannanase), or the immune system stimulation treatment (ISS; basal diet + 0.056% -mannanase, challenged with repeated increasing doses of Escherichia coli LPS). The experiment consisted of a 10-d adaptation period, 5- d digestibility and nitrogen balance measurement, 22 h of heat production (HP) measurements, and 12 h of fasting HP measurements in indirect calorimetry chambers.

The immune challenge consisted of 4 injections of either LPS (ISS) or sterile saline

(CON and ENZ), one every 48 h beginning on d 10. Blood was collected pre- and post- challenge for complete blood counts with differential, haptoglobin and mannan binding lectin, 12 cytokines, and glucose and insulin concentrations. Beta-mannanase supplementation did not affect immune status, nutrient digestibility, growth performance, energy balance, or MEm. The ISS treatment induced fever, elevated proinflammatory cytokines and decreased leukocyte concentrations (P < 0.05). The ISS treatment did not impact nitrogen balance or nutrient digestibility (P > 0.10) but increased total HP (21%) and MEm (23%), resulting in decreased lipid deposition (-30%) and average daily gain (-

18%) (P < 0.05). This experiment provides novel data on -mannanase supplementation effects on immune parameters and energy balance in pigs and is the first to directly relate decreased ADG to increased MEm independent of changes in feed intake in immune challenged pigs. Immune stimulation increased energy partitioning to the immune system by 23% which limited lipid deposition and weight gain. Understanding energy and nutrient partitioning in immune-stressed pigs may provide insight into more effective feeding and management strategies.

Introduction

The negative influence of an immune challenge on animal growth is well established. Pro-inflammatory cytokines orchestrate an immune response resulting in fever, acute phase protein (APP) production, and leukocyte proliferation, each of which requires additional energy and amino acids (AA). Therefore, a perceived immune challenge can theoretically partition energy and nutrients away from productive processes such as muscle growth and negatively impact the efficiency and cost of meat production

[1]. Innate immune activation occurs when pathogen-associated molecular patterns are detected such as the lipid-A component of lipopolysaccharide (LPS) from gram-negative bacteria [2]. However, certain dietary components, such as -mannan in soybean, copra, and palm kernel meals, mimic carbohydrate structures on pathogen surfaces [3] and have previously been shown to activate the innate immune system [4,5], termed a feed-induced immune response (FIIR).

To inhibit a -mannan derived FIIR, interest in β-mannanase enzyme supplementation has increased. It is hypothesized that the hydrolyzed manno- oligosaccharides can no longer crosslink and stimulate multiple mannose receptors, thus reducing immune stimulation and associated energy costs. Research in poultry demonstrated that β-mannanase decreased plasma APP concentration and improved growth performance and feed efficiency leading to the conclusion that β-mannanase supplementation spared energy through prevention of the FIIR [6,7]. In pigs, performance responses to β-mannanase are less consistent than in poultry and reports on immune responses are limited and effects on energy partitioning have yet to be evaluated.

Nutrient partitioning in pigs during a pathogen challenge has received more attention, often utilizing a LPS challenge model [8]. Physiological responses to an LPS challenge in pigs have been well characterized. Similar to disease challenges, LPS induces anorexia, fever, and nutrient repartitioning leading to decreased growth and efficiency [1,2,9]. Fever is an energetically expensive process and its effects on sheep and human maintenance energy requirements have been estimated [10]. Immune system activation also significantly shifts glucose metabolism and glucose requirements during an LPS challenge in pigs have been estimated to be approximately 1.1 g/kg BW0.75/h

[11]. Yet few studies have addressed comprehensive changes in energy partitioning during an immune response and clear relationships between measured immune responses and the direction and magnitude of changes in energy partitioning have yet to be elucidated.

Therefore, the objectives of this experiment were to determine how β-mannanase supplementation and innate immune stimulation each affect maintenance energy requirements and to characterize changes in immune parameters, nutrient digestibility, growth performance, and energy balance. We hypothesized that innate immune stimulation would increase maintenance energy requirements by initiating a cytokine- driven febrile response and inflammatory state, and that β-mannanase supplementation would decrease maintenance energy requirements through an energy sparing effect of

FIIR prevention.

Materials and methods

All experimental procedures adhered to guidelines for the ethical and humane use of animals for research and were reviewed and approved by the University of Manitoba

Animal Care Committee.

Animals and experimental design

Thirty growing barrows ([Yorkshire × Landrace] × Duroc) were acquired from the Glenlea Swine Research Unit, University of Manitoba at an average body weight

(BW) of 9.60 ± 2.00 kg. The experiment was conducted using a randomized complete block design. Pigs were blocked by weight and randomly assigned to one of three treatments (Table 1). A staggered time course was utilized to accommodate the limited number of calorimetry chambers available, whereby 10 blocks of three pigs each (one pig per treatment) began the experiment 4 d after the previous block. Day one BW was similar among treatments (10.27 ± 0.08 kg).

Experimental diets, treatments and procedures

All diets were formulated on the ratio of standardized ileal digestible lysine to metabolizable energy (ME) and met or exceeded all specified nutrient requirements of growing pigs from 11 to 25 kg [12]. Pigs were fed at 2.5 times their maintenance ME requirements [12], once daily at 0800 h and had free access to water at all times. Pigs were fed a common pre-trial diet that was corn-soybean meal-based. The experimental basal diet (Table 2) was formulated with high soybean meal and soybean hull inclusion levels to increase dietary β-mannan concentration.

Due to the availability of three indirect calorimetry chambers, three experimental treatments were evaluated (Table 1). The control treatment (CON) received the basal

diet, while the enzyme treatment (ENZ) received CON supplemented with 0.056% β- mannanase (HemicellTM HT-D, Elanco Animal Health, Guelph, ON, Canada; endo-1,4-β- mannanase (160 × 106 units/kg) from Paenibacillus alvei). The third treatment was challenged with repeated LPS immune system stimulation (ISS) and received the same diet as ENZ. This treatment design was determined based on the hypothesis, supported by previous research, that β-mannanase would inhibit a FIIR if it occurred in CON [6,7]. In this way, the effect of innate immune stimulation by LPS could be evaluated independent of a FIIR.

Upon arrival and during the pre-trial period, pigs were housed individually in pens (1.83 × 1.22 m) with plastic-covered expanded metal flooring in a temperature- controlled room (26 ± 2°C). Daily feed allotment during the pre-trial period was adjusted based on BW measured every 4 d. Pigs were maintained on the pre-trial diet for at least 4 d until initiation of the experiment for their respective block, at which time pigs received their assigned treatment diets. The experiment consisted of a 10-d adaptation phase, a 5-d total feces and urine collection phase, and 34 h of heat production (HP) measurements.

At trial initiation (d 1), pigs were individually housed in adjustable metabolism crates (1.80 × 0.60 m) with smooth transparent plastic sides and plastic-covered expanded metal flooring in a temperature-controlled room (26 ± 2°C). Body weight was measured on d 1, 5, 10, 16, daily feed allotment was adjusted accordingly, and pigs were trained to consume the entire meal within 1 h of feeding at 0800 h. Orts, if any, were measured to determine average daily feed intake (ADFI).

Immune challenge

A low-dose, repeated LPS challenge, following the modified procedures described by Rakhshandeh and de Lange [8], was chosen to induce an inflammatory response representative of sustained immune system stimulation in the ISS treatment. The challenge consisted of four repeated low-dose injections of Escherichia coli LPS serotype

O55:B5 (Sigma–Aldrich, St. Louis, MO, USA) for pigs on treatment ISS, or a control injection of sterile saline for pigs in treatments CON and ENZ. The LPS was dissolved in sterile PBS so that an injection of 0.1 mL/kg of BW achieved the desired dosage [13].

A pilot study with 12 pigs was conducted prior to experiment initiation to discern the lowest appropriate initial LPS dose and the subsequent dose increase regimen required to limit LPS tolerance development. Results of the pilot study (not reported herein) indicated that an initial dose of 20 g LPS/kg of BW with subsequent dose increases of 20, 30, and 40% was the regimen that maintained a febrile response (rectal temperature ≥ 40°C) at all four challenges while minimizing anorexia and vomiting.

Pigs were injected intramuscularly at 1000 h on d 10, 12, 14, and 16 with either sterile saline or LPS, following the previously described dosing regimen determined from the pilot study. Baseline rectal temperature was measured on d 5 and 8 at 1400 h and at 4 h post-challenge (1400 h) on d 10, 12, and 14. Blood samples were then collected on d 8

(pre-challenge) and d 10 (post-challenge) via jugular venipuncture into two 10 mL tubes for EDTA-whole blood and serum. Whole blood samples were placed on ice pending transportation to the laboratory for complete blood count (CBC) analysis with white blood cell (WBC) manual differential. Serum was separated by centrifugation (2000 × g

for 15 min at 4°C), collected and divided into three subsamples, and stored at -80°C until analyzed.

Digestibility

On d 10, pigs received 5 g of ferric oxide as an indigestible marker mixed with

100 g of feed; the remaining allotted feed was offered after the marked feed was consumed. Fecal collection commenced when the marker first appeared in the feces. On d

15, pigs were offered 100 g of marked feed as previously described, and fecal collection terminated when the marker appeared in the feces. Feces were weighed and stored at -

20°C until further processing. Total urine collection commenced at 0800 h on d 11 and terminated at 0800 h on d 16. Urine was collected once daily into jugs containing 10 mL of 6 mol/L HCl. Urine was weighed, thoroughly mixed and subsampled (10% of urine weight), strained through glass wool, and stored at -20°C. Urine subsamples were pooled per pig throughout the collection period.

Heat production

On d 16, within 30 min of consuming their daily feed allotment, pigs were transferred to open-circuit indirect calorimetry chambers (1.22 × 0.61 × 0.91 m metallic box with a glass door on the front side, plastic-covered expanded metal sheet flooring, and a valve at the bottom to collect urine; Columbus Instruments, Columbus, OH, USA) for 34 h of calorimetric measurements. Pigs were randomly assigned to chambers to reduce the possibility of a chamber bias. The first 2 h of HP (0800 – 1000 h), measured prior to pigs receiving the fourth and final challenge of either LPS or saline, were designated as acclimation and not included in HP calculations. Total oxygen consumption

(VO2) and carbon dioxide production (VCO2) were measured every 12 min

corresponding to 170 values over 34 h. The first 24 h following feeding was designated as the fed state and the last 12 h as the fasting state [14,15]. Urine voided during the fed and fasting periods was collected separately and processed as previously described.

Personnel movement within the room was minimized during HP measurement to avoid any disturbance of the pigs. The system was validated using the alcohol combustion method described by Aulick et al. [17] and the O2 and CO2 sensors were calibrated prior to each block of the experiment. The chambers were air-conditioned to maintain a constant temperature (23 ± 1°C). Pig BW on d 16 was similar across all treatments (14.1

± 0.3 kg). Heat production was measured in only seven of the ten experimental blocks because of equipment failure during three blocks; therefore, for HP data, n = 7.

Analytical methods

All diet, orts, and fecal samples were dried at 60°C to a constant weight and were ground to a particle size of 1 mm. Urine samples were thawed, sieved through cotton gauze, and filtered with glass wool. Urine, diet, and fecal samples were analyzed in duplicate for nitrogen (N; method 990.03 [17]; TruMac®; LECO Corp., St. Joseph, MI,

USA). An EDTA sample (9.56% N) was used as the standard for calibration and was determined to be 9.55 ± 0.01% N. Crude protein (CP) was calculated as N × 6.25. Diets were analyzed for β-mannanase concentration (colorimetric determination, Elanco

Animal Health, Gaithersburg, MD).

Diet and fecal samples were analyzed in duplicate for dry matter (DM; method

930.15), acid hydrolyzed ether extract (EE; method 2003.06), and starch (Total Starch

Assay Kit, Megazyme International, Wicklow, Ireland, method 996.11) using standard methods [17]; and in triplicate for neutral and acid detergent fiber components (NDF [18]

and ADF [19], respectively). Hemicellulose was calculated as the difference between

NDF and ADF concentrations. Gross energy (GE) was determined using a bomb calorimeter (model 6200; Parr Instrument Co., Moline, IL). Benzoic acid (6,318 kcal

GE/kg; Parr Instrument Co.) was used as the standard for calibration and was determined to contain 6,325 ± 6.9 kcal GE/kg. Urine GE was calculated as 192 plus 31 times the concentration of urinary N [20] and multiplied by a factor of 0.239 to convert the unit to kcal.

Pre- and post-challenge whole blood samples and blood smears were analyzed for

CBC performed by Manitoba Veterinary Diagnostic Services (Winnipeg, MB, Canada) using Advia 2120i (Siemens Healthcare Diagnostics, Tarrytown, NY, USA) with a manual differential. The main response variables of interest were total cell, RBC, and

WBC (mature and immature neutrophils, eosinophils, basophils, lymphocytes, and monocytes) concentrations.

Serum was divided into three subsamples. One set was analyzed for glucose and insulin concentrations by Animal Health Laboratory (University of Guelph, ON, Canada).

Glucose concentration was determined on an automated Roche Cobas C501 analyzer

(GLUC3 application, Roche Diagnostics, Indianapolis, IN, USA) and insulin concentration was quantified using commercial RIA kits (PI-12K, EMD Millipore,

Billerica, MA, USA). The second serum subset was analyzed for cytokine concentrations

(granulocyte macrophage colony-stimulating factor (GM-CSF), tumor necrosis factor alpha (TNFα), Interleukin (IL)-one-alpha (IL-1α), IL-1β, IL-one-receptor antagonist

(IL-1ra), IL-2, IL-4, IL-6, IL-8, IL-10, IL-12, and IL-18) by a commercial multiplex assay using laser bead technology (Eve Technologies, Calgary, AB, Canada). The third

serum set was analyzed for concentration of APPs haptoglobin and mannose-binding lectin A (MBL) using porcine-specific commercial ELISA kits (Immunology Consultants

Laboratory, Inc., Portland, OR, USA; MyBioSource, Inc., San Diego, CA, USA, respectively).

Calculations

Dry matter, GE, CP, EE, starch, hemicellulose, NDF, and ADF apparent total tract digestibility (ATTD; %) were calculated on a DM basis as [(nutrient intake – nutrient output in feces)/nutrient intake] × 100. Digestible energy (DE) content of the diet was calculated as GE × GE ATTD. Dietary ME was calculated according to the equation of

Noblet et al. [21]: ME = DE – [urine GE + (0.4% of DE intake)]. Nitrogen retention (NR) was calculated by the difference between N intake and N excreted in the feces and urine, and protein deposition (PD) was determined as NR × 6.25.

Heat production was calculated from respiratory gas exchanges and urinary N production according to the equation of Brouwer (1965): HP (kcal) = 3.87 × VO2 consumed (L) + 1.20 × VCO2 produced (L) – 1.43 × urinary N production (g). Methane production was not accounted for, but has been estimated to be very low in growing pigs

(< 1%, [23]). All HP parameters were normalized to a period of 24 h, expressed as kcal of heat produced per kg of BW0.60 [24] and per kg of DM intake (DMI) in order to remove known effects of variations in BW and DMI [25,26].

Total heat production (HPtotal) was the average HP during the 22 h of post- challenge, fed state measurement. Total fasting heat production (FHPtotal) was the average HP over the 12 h fasted-state. Because the system was not equipped to quantify and separate HP due to physical activity, fasting heat production (FHP) was derived from

the 10 lowest HP values over the fasted-state, reflecting energy metabolism due to basal metabolic rate and not associated with feed consumption, digestion, or physical activity

[27,28]. The respiratory quotient (RQ) was calculated as VCO2 divided by VO2 during the fed (RQfed) and fasting (RQfast) states.

To best estimate components of HP not attributed to the basal metabolic rate, HP values for physical activity and the thermic effect of feeding (TEF) were calculated.

Activity heat production (AHP) was estimated utilizing fed-state HP data measured over

10 h post-challenge to represent normal post-feeding daytime behavior. The difference between the average HP over this 10 h period (HP10) and the average of the 10 lowest HP values over the same time (HPlow; representative of sedentary, resting behavior; [27,28]) was designated as AHP. The TEF was calculated as the difference between HP10 and the sum of AHP and FHP, so that any HP in excess of basal metabolism and activity was partitioned toward TEF. Heat increment (HI) was then calculated as the sum of AHP and

TEF. Using these data, the efficiency of utilizing ME for maintenance and growth (kmg,

%) was calculated as (1 – HI / ME intake) × 100 [25,29]. To address the primary research question of how an immune challenge and β-mannanase supplementation impact maintenance energy requirements, metabolizable energy used for maintenance (MEm) was then calculated as FHP × 100/ kmg [25,29].

Together, dietary energy, N balance, and HP values were utilized to characterize energy use and balance in the pig. Retained energy (RE) was calculated by the difference of ME intake and the total HP during the 24 h fed-state (both pre- and post-challenge) to account for all energy not available for tissue accretion [29]. Energy retained as protein

(REp) was calculated from N balance assuming a PD (g) energy value of 5.64 kcal/g [30].

Energy retained as lipid (REl) was calculated as the difference between RE and REp [30].

Lipid deposition (LD) was then determined from REl assuming an energy content of 9.49 kcal/g of deposited lipid [22]. Dietary net energy (NE; kcal/kg) was calculated as the sum of RE and FHP divided by DMI [21].

Statistical analyses

Data were analyzed as a randomized complete block design with pig as the experimental unit. The UNIVARIATE procedure of SAS (Version 9.4, SAS Inst., Cary,

NC) was used to verify normality and homogeneity of variances. Statistical outliers (> 3

SD away from the mean) were removed; therefore, one pig from the ISS treatment was removed from HP data because of poor feed intake (45% decrease) on the day of HP measurement. Immature neutrophil, eosinophil, and basophil CBC data were log transformed to achieve a normal distribution.

The main effects of dietary treatment and block were analyzed using the MIXED procedure of SAS. The staggered block experimental design resulted in variations in time and body weight among blocks. This variation was expected and resulted in statistical detection of block as a significant main effect in most response variables. Therefore, block remained in the statistical model, but block P-values are not reported herein.

Differences among treatments were determined using ANOVA and means were separated using the least square means statement and the PDIFF option. Immune and rectal temperature data were analyzed as repeated measures and covariance structures resulting in the lowest AIC values for each variable were applied. To further evaluate β- mannanase effects on immune parameters pre-challenge, contrasts comparing CON versus ENZ and ISS values were generated using the contrast statement of the MIXED

procedure. Differences were considered significant if P was ≤ 0.05 and a trend if P was >

0.05 and ≤ 0.10.

Results

Immune response parameters

Immune system stimulation effects

Pigs on the ISS treatment exhibited minimal vomiting, diarrhea and signs of lethargy and hyperventilation after the first and to a lesser extent, the second LPS injection. After the third and fourth challenges, ISS pigs continued to demonstrate signs of lethargy and hyperventilation, but vomiting and diarrhea were not observed. No pigs died after any injection. The immune stimulation model successfully induced a sustained febrile response (rectal temperature ≥ 40°C) in ISS pigs on d 10, 12, and 14 (treatment by day interaction P < 0.0001; Fig. 1). Pigs in CON and ENZ treatments maintained normal rectal temperatures (38.85°C ± 0.15) throughout the experiment.

There was a significant interaction between the effects of time (pre- or post- challenge) and treatment on WBC, mature neutrophil, lymphocyte, and monocyte counts and a trend for an interaction on RBC count. In all four variables, treatments had similar cell counts at the pre-challenge time point (P ≥ 0.10). Post-challenge, immune stimulation by LPS decreased WBC, mature neutrophil, lymphocyte, and monocyte counts compared to CON and ENZ (P ≤ 0.05; Fig. 2). There were no differences among treatments or time periods for total cell, immature neutrophil, eosinophil, or basophil counts (P > 0.10).

Glucose, insulin, haptoglobin, and MBL serum concentrations were not affected by the interaction of time and treatment (P > 0.10), but concentrations were higher pre- challenge compared to post-challenge for glucose, haptoglobin, and MBL.

Lipopolysaccharide challenge increased IL-1β, IL-1ra, IL-6, IL-8, and TNFα concentrations post-challenge compared to ISS pre-challenge and both pre- and post- challenge concentrations in CON and ISS (Fig. 3). All other cytokines were not significantly impacted by the interaction or main effects of time and treatment (P > 0.10).

Interferon-gamma was not detected in any of the samples. Serum GM-CSF concentrations were not different (P > 0.10) among CON and ENZ pre-and post- challenge and ISS pre-challenge, while ISS post-challenge GM-CSF concentration was increased compared to the ISS pre-challenge value and CON and ENZ post-challenge values (P ≤ 0.015; Fig. 3).

β-mannanase effects

Contrasts comparing immune cell dynamics of pigs fed either the control or β- mannanase diet prior to the first challenge on d 10 detected no differences in CBC values

(P ≥ 0.10; Table 3). Similarly, serum glucose, insulin, MBL, and cytokine concentrations

(except IL-1α) did not differ with β-mannanase supplementation (P ≥ 0.230; Table 4).

Serum haptoglobin and IL-1α concentrations were decreased in diets supplemented with

β-mannanase (P ≤ 0.05; Table 4)

Pig growth performance, nitrogen balance, and diet digestibility

Average initial BW was 10.27 ± 0.15 kg, d 16 average BW was 15.12 ± 0.27 kg, and BW did not differ among treatments at either time point (P ≥ 0.471). Average daily gain (ADG) over the entire 16-d experiment was not different among treatments (P =

0.13; Table 6), but ISS ADG during the immune challenge (d 10-16) was less than CON and ENZ gain (P = 0.010; Table 7).

Immune system simulation numerically decreased ADFI and thus N intake compared to CON and ENZ (P = 0.021), resulting in decreased fecal N excretion on a g per d basis (P = 0.007). Urine N excretion during the challenge period was similar among treatments (P = 0.045) but retained N in the ISS treatment was less than that of CON, with ENZ being intermediate (P = 0.045; Table 6). Partitioning of excreted N to either the feces or urine was not different among treatment (P = 0.78). When N excretion was expressed as a percent of N intake, the previously observed significant treatment effect on fecal N excretion was no longer evident (Table 6). There were no differences among treatments in ATTD of any analyzed nutrient (P ≥ 0.120; Table 5) and all ATTD coefficients were within normal ranges for 10 to 15 kg pigs

Heat production, maintenance energy requirements, and energy retention

Day 16 ME intake was similar among treatments (759.4 ± 37.7 kcal/kg BW0.60/kg

DMI/d; P = 0.92). Immune system stimulation increased fed state HPtotal compared to

CON and ENZ (P = 0.040; Table 7). In the fasting state, neither immune stimulation nor

β-mannanase supplementation affected FHPtotal or FHP compared to control (P ≥ 0.135).

Treatment did not affect RQ in the fed and fasting states (P ≥ 0.23; Table 7).

0.60 Immune system stimulation increased MEm (kcal/kg BW /kg DMI/d) compared to CON or ENZ pigs (P = 0.045), but kmg among treatments did not differ (P = 0.13;

Table 7). When MEm was expressed as kcal/d, the significant treatment effect was no longer detected (P = 0.90). Beta-mannanase supplementation did not change MEm

relative to CON whether expressed as kcal/kg BW0.60/kg DMI/d (P = 0.98), kcal/kg

BW/d (P = 0.72), or kcal/d (P = 0.77).

Absorbed energy not lost via urine, gases, heat increment, activity and TEF, or maintenance, is retained as either protein or lipid. Immune system stimulation decreased

REl compared to CON and ENZ (P = 0.046) but REp and total RE were not different among treatments (P > 0.32) when expressed as kcal/kg BW0.60/kg DMI/d (Table 7).

When RE was expressed as a proportion of ME intake, similar treatment effects were observed for REl and REp, but a significant decrease in total RE was detected due to ISS

(P = 0.033; Table 7). As less energy was retained as lipid, LD was decreased in the ISS treatment compared to CON and ENZ (P = 0.047) while no differences were observed in

PD (P = 0.15; Table 7).

Dietary energy values and efficiency

The ENZ and ISS treatments tended to decrease diet DE and ME values relative to CON (P ≤ 0.052; Table 8). Neither ISS nor β-mannanase supplementation (ENZ treatment) affected dietary NE value (P = 0.75) or ME and NE efficiency (P ≥ 0.46).

Discussion

During an immune challenge, pro-inflammatory cytokines initiate a shift in nutrient partitioning away from tissue growth to support activation and maintenance of an immune response [1,11,31]. The results of this experiment clearly demonstrate a systemic inflammatory response to LPS occurred, verified by increased concentrations of pro- inflammatory cytokines and elevated body temperature. To our knowledge, these data are the first to directly relate decreased ADG to increased MEm independent of changes in

feed intake during an immune response. Additionally, this experiment provides novel data on -mannanase supplementation effects on immune parameters and energy balance in pigs.

-Mannanase

As a constituent of hemicellulose, -mannan is not digested by mammalian endogenous enzymes [32]. Thus, intact -mannans are available to bind carbohydrate recognition domains of pattern recognition receptors on innate immune cells surveying the intestinal epithelium for potential pathogens [3,33]. In this way, -mannans are hypothesized to be capable of stimulating innate immune cells resulting in a nonproductive, energy draining immune response [4–6].

Typically, only growth and feed efficiency responses have been measured from - mannanase supplementation and reported in the animal nutrition literature. Reduced feed efficiency and ADG have been reported with increasing dietary -mannan concentrations

[34]. Therefore, there is interest in -mannanase supplementation to alleviate these negative effects by enzymatic hydrolysis of -mannan polysaccharides. The FIIR was alleviated through -mannanase supplementation in poultry [6,35]; however, - mannanase supplementation responses in swine have been inconsistent. This experiment demonstrated no -mannanase effect on the ATTD of DM, GE, CP, EE, or hemicellulose.

Growth performance responses are similarly inconsistent with positive results in some studies [36–38], but no -mannanase effect in others [39–42]. In this experiment, ENZ did not improve ADG, protein, or lipid deposition.

Dietary -mannans are proposed to stimulate the innate immune system through direct interactions with the carbohydrate binding domains of mannose recognition receptors such as the membrane bound mannose receptor and secreted MBL. Therefore, serum MBL concentrations were measured to determine if -mannanase supplementation decreased circulating MBL, theoretically by removing the substrate for activation and synthesis. Serum MBL concentrations were not affected by -mannanase. This may indicate that the -mannan concentration in the intestinal lumen was not high enough to either interact with MBL, MBL-dietary -mannan interaction was not affected by - mannanase supplementation, or this interaction is not a mechanism through which - mannans are sensed by the innate immune system.

Two significant differences in serum parameters were detected when contrasts were applied to compare pre-challenge values between control pigs (no -mannanase,

CON treatment) and -mannanase supplemented pigs (ENZ and ISS treatments). Beta- mannanase supplementation decreased serum haptoglobin and IL-1 concentrations. In poultry, decreased haptoglobin has been proposed as evidence of immune stress alleviation due to -mannanase supplementation [6]. However, this response occurred in conjunction with growth performance and feed efficiency improvements which were not observed in this study. Beta-mannanase effects on IL-1 concentrations have not been previously reported. Interleukin-1-alpha can be involved in inflammation initiation, but the relationship between serum concentration and magnitude of immune challenge is not as clear as the implication of its counterpart, IL-1 on systemic inflammation [43].

Interleukin-1-beta concentrations were not affected by -mannanase supplementation in this study.

Collectively, decreased serum IL-1 and haptoglobin concentrations are not strong enough evidence of an alleviated systemic FIIR when taken in context with the lack of all other measured inflammatory-type variables. Importantly, no differences were observed in HP, MEm, and growth performance. It is possible that a localized response may have occurred at the intestinal level yet went undetected systemically. However, if this occurred, whole body nutrient and energy partitioning were still unaffected. The hypothesis that -mannanase supplementation would decrease MEm was not supported.

Pigs fed diets supplemented with -mannanase had similar WBC counts, cytokine concentrations, nutrient digestibility, ADG, N and energy balance, PD, LD, and MEm compared to CON pigs.

Immune stimulation

Innate immune stimulation was successfully induced in pigs using sequential, increasing doses of E. coli LPS. Elevated rectal temperature, increased pro-inflammatory cytokine concentrations, and altered nutrient and energy partitioning are all hallmarks of a chronic immune challenge [1] and were observed in ISS pigs in this study. One limitation of this study was the number of calorimetry chambers available which limited the experiment to a total of three treatments. Due to this limitation, we were unable to evaluate the interaction of -mannanase supplementation with LPS immune stimulation.

Thus, interpretation of ISS effects has been made in comparison to the ENZ treatment.

However, as discussed above, there were no differences between the CON and ENZ treatments in nutrient digestibility, ADG, or N and energy balance. The major finding of

this research indicates that the innate immune challenge increased young pig maintenance energy requirements by 23.3% which translated into a 18.3% decrease in ADG.

Unique to this study, decreased ADG could be attributed primarily to increased

MEm in ISS pigs as opposed to decreased feed intake or effects on nutrient digestibility.

Anorexia is a well-established response to systemic immune stimulation [2,9,44] induced by pro-inflammatory cytokine actions (especially IL-1) in the brain and modulation of metabolism and hormone release [45]. In this study, a numerical but not statistically significant decrease in ADFI was observed in ISS pigs during the challenge period even though IL-1 increased. It is likely that a stronger ADFI decrease was not observed as a consequence of challenging the pigs 2 h post-feeding and limit feeding to 2.5 times maintenance energy requirements [12]. This feeding level was designed to achieve similar ADFI for pigs on all treatments because of the known effect of previous feeding level on HP [25]. To further ensure HP results were separated from feed intake and BW effects, all energy balance calculations were conducted on a kcal/BW0.60/DMI/d basis.

Just as feed intake did not influence the observed decrease in ADG of ISS pigs, nutrient digestibility was not different across treatments. This is in agreement with other studies reporting ATTD during a chronic LPS challenge [46,47].

Febrile response

Before the challenge period, rectal temperatures and blood immune parameters in

ISS pigs were not different from those on the CON and ENZ treatments. This confirmed that prior to the challenge all pigs were in good health and of similar immune status.

Therefore, any subsequent differences during the challenge were attributed to LPS

immune stimulation. Elevated rectal temperatures (> 40°C) post-challenge on d 10, 12, and 14 indicated a febrile response in ISS pigs.

Fever is energetically expensive with increased caloric requirement estimates ranging from 7-15% for each 1°C increase in body temperature [48]. Utilizing the average rectal temperature of CON and ENZ pigs and the post-challenge temperature of

ISS pigs on day 14, an increase of 1.2°C resulted in a 23.3% increase in MEm caloric requirements. This value is higher than the previously described range and may indicate that the majority, but not all of the increase in maintenance caloric requirement is to support the febrile response. The remainder may be partially explained by an increase in immune cell glucose requirements [11].

Cytokines

Key pro-inflammatory cytokines include TNF, IL-6, and IL-1 [49] and ISS pigs had increased serum concentrations of all three after the first LPS challenge. Pro- inflammatory cytokines shift metabolism away from anabolic processes toward a more catabolic state to generate AAs and energy necessary to support fever, increase immune cell proliferation, and APP synthesis [50,51]. In this study, the pro-inflammatory cytokine profile of ISS pigs clearly shifted metabolism toward a lipolytic state and this resulted in significantly less energy retained as lipid and decreased lipid deposition compared to non-immunologically challenged pigs.

Complete blood count

Immune stimulation decreased WBC counts, specifically neutrophils, lymphocytes, and monocytes. This is similar to other instances of leukopenia observed due to LPS administration [11,52]. However, WBC distribution drastically changed

following LPS administration and circulating concentrations are dependent upon the time of sampling relative to immune challenge [52,53]. Thus, variable responses in WBC counts have been reported due to LPS immune stimulation. Rakhshandeh and de Lang observed 1.6 times greater WBC in one study [8], but in a second, WBC count decreased by 9% [54]. At the time of sampling in this study, leukocyte extravasation into the LPS injection site and into immunologically important tissues likely explains the observed leukopenia.

Acute phase proteins

In addition to increased pro-inflammatory cytokine production and leukocyte migration that occur during infection, the acute phase response typically includes increased APP synthesis by the liver. However, in this study, ISS APP concentrations did not differ compared to CON. This was an unexpected result because LPS has been demonstrated to increase APPs such as haptoglobin [8,46] and C-reactive protein [55] in pigs. A less responsive APP, MBL has been demonstrated to attenuate LPS-induced pro- inflammatory cytokine production [56] and inhibit T-lymphocyte activation [57].

However, in this study it did not appear that LPS induced greater MBL or haptoglobin production.

Although a MBL response was not necessarily expected, a haptoglobin response was. Haptoglobin is a primary APP in pigs and is synthesized in the liver when activated by IL-6 and to a lesser extent IL-1 [58], both of which were significantly elevated in ISS pigs post-challenge. Similar to our results, Koopmans et al. [55] discussed unpublished data which showed no LPS effect on haptoglobin concentrations even though there were clear increases in plasma cortisol, TNF, IL-6, and C-reactive protein over a 24-h period

after LPS challenge. One possible explanation for a lack of haptoglobin response could be time related. Serum samples in this study were collected 4 h after the first challenge and haptoglobin may be a better indicator of chronic inflammation [59].

Nitrogen balance

Disease is associated with decreased growth performance and changes in nutrient partitioning. Often, N metabolism is affected because of increasing AA requirements for immune cell proliferation and APP synthesis [51]. In this study, only numerical decreases in protein deposition were measured in ISS pigs compared to CON and ENZ. If protein catabolism had increased to provide AAs for APPs, an increase in urinary N would have been expected because APPs have a distinctly different AA profile than skeletal muscle

[50,60]. However, due to the high dietary CP concentration, it is possible that these excessive dietary amino acids may have provided the additional amino acids required for

APP synthesis and prevented the typically observed increase in skeletal muscle protein catabolism.

Energy balance

Disease is well known to be detrimental to pig efficiency and productivity. A considerable amount of research has focused on products to mitigate the drop in performance [13,61] or prevent initial disease onset [62]. Yet few studies have evaluated the energetic cost of an immune challenge in order to generate more effective dietary interventions. In this study, total HP increased by 21.1% in ISS pigs compared to the

ENZ treatment.

Campos et al. [46] also evaluated HP components during an immune response and reported significant decreases in ADFI leading to decreased TEF compared to baseline

values. In this study DMI did not differ, potential feed intake effects on TEF were removed by interpreting the data after normalizing to a constant feed intake, and TEF values were not affected by ISS. Therefore, both experiments indicate that a chronic inflammatory response did not increase HP through increased TEF. This is supported by the lack of treatment differences in diet digestibility and further supports our supposition that the impact of immune stimulation on energy balance in this study is not through influences on diet digestion or nutrient uptake.

However, it is clear that energy partitioning between maintenance and growth was affected by ISS. A 23.3% increase in MEm was detected due to ISS. As caloric requirements for maintenance increased to support the immune system, less dietary energy was retained for growth. This manifested as less REl resulting in a 30.2% decrease in lipid deposition.

Previous studies across all species have related increased caloric requirements with fever [48,63], but few have directly related a chronic immune challenge with increased MEm. In vitro studies with isolated mitochondria from rats stimulated with

TNF or IL-1 showed up to 30% increases in respiration rate [64]. Demas et al. [63] reported that mice injected with a mild antigen had limited immune activation that resulted in significantly more O2 consumption than control mice injected with saline.

Interleukin-six infusions in humans increased resting metabolic rates by 25% [65].

In pigs, the direct relationship between immune stimulation and increased energy requirements has not previously been demonstrated. Some studies reported that immune system stimulation did not impact growth, efficiency, or energy balance measurements

[66,67] However, Moon et al. [66] reported fibroblast formation at the injection site

which encapsulated the immunogen and prevented systemic delivery. Williams et al. [67] used the comparative slaughter technique and reported no differences in the energetic costs of maintenance, PD, and LD between pigs raised in environments encouraging high or low chronic immune activation.

Conversely, Labussière et al. [68] and Campos et al. [46] reported decreased HP in pigs during inflammatory challenges. Labussière et al. [68] administered a single injection of complete Freund’s adjuvant to young weaned pigs but did not measured HP until the day after challenge and only re-entered the calorimetry chamber after visual recovery [68]; and this likely biased the response. Campos et al. [46] reported a 14% decrease in total HP (kcal/BW0.60/d) in response to a repeated LPS challenge in growing pigs even though typical inflammatory-type and febrile responses were observed.

Decreased HP was mainly attributed to lower TEF which reflected the effect of feed intake depression on HP. According to the relationship reported by Labussière et al. [25],

0.60 lower ADFI should have decreased MEm by 24 kcal/BW /d. Because this drop in MEm did not occur, the authors reasoned that the immune stimulation did in fact increase MEm relative to baseline [46]. This supports our experimental model of limit feeding to encourage similar feed intake and to evaluate energy balance on a kcal/BW0.60/DMI/d basis. Feed intake clearly influences and can bias HP results and interpretations.

Interpretation of our results in context with the previously discussed reports suggests that an inflammatory response does increase MEm relative to healthy control animals, but in some experiments this response may be masked by decreased HP related to decreased feed intake. This may mean that during an immune response the total caloric requirement may not drastically change because of decreased feed intake, but how those

calories are partitioned does change; and this results in growth and feed efficiency depressions commonly observed during disease challenges.

These results supported our hypothesis that energy partitioning shifts to allocate more energy for initiation and maintenance of immune functions and less toward nutrient deposition. Other research would support changes in N metabolism [46,67,69] whereas our data suggest that less energy was allocated for LD. Both result in decreased ADG and efficiency losses in pork production, yet these effects are generally given little consideration in commercial swine feeding practices.

Conclusions

This experiment provides novel data on -mannanase supplementation effects on immune parameters and energy balance in pigs. Beta-mannanase supplementation did not affect immune status, nutrient digestibility, growth performance, energy balance, or MEm in young pigs fed a corn, soybean meal, and soybean hulls-based diet. More research is needed to determine how -mannanase functions in pigs and in which environments and diets it might be effective. These novel data directly relate decreased ADG to increased

MEm independent of changes in feed intake in immune challenged pigs. An innate immune challenge increased proinflammatory cytokine concentrations which induced a febrile response and elevated HP and MEm by 23.3%. Increased energy partitioning toward the immune response limited LD by 30.2% leading to a 18.3% decrease in ADG during the immune challenge. These data expand upon the available literature to describe the magnitude of increase in MEm in immune challenged pigs relative to healthy control animals. Understanding the extent to which energy requirements and nutrient deposition

change in pigs experiencing sustained immune stress may help develop more effective feeding strategies for health challenged herds and encourage appreciation for the economic benefits of maintaining high health populations.

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Table 1. Summary of experimental treatments Experimental treatment

CON1 ENZ2 ISS3

Diet Control Control + β-mannanase Control + β-mannanase

β-mannanase inclusion No Yes Yes

Challenge treatment Saline Saline E. coli LPS 1Control treatment (CON) = pigs fed basal diet with no LPS (Escherichia coli serotype O55:B5) injection 2Enzyme treatment (ENZ) = pigs fed enzyme diet (0.056% β-mannanase) with no LPS injection 3Immune system stimulation treatment (ISS) = pigs fed enzyme diet (0.056% β-mannanase) with LPS injection

Table 2. Experimental diet ingredient and analyzed nutrient composition (as-fed basis) Item Control diet Enzyme diet Ingredient, % of diet Corn 47.33 47.27 Soybean meal, 38.40 38.40 (dehulled, solvent extracted) Soybean hulls 10.00 10.00 Soybean oil 1.85 1.85 Limestone 1.04 1.04 Monocalcium phosphate 0.60 0.60 Vitamin premix1 0.33 0.33 Trace mineral premix2 0.20 0.20 Salt 0.25 0.25 Hemicell HT-D3 0.00 0.06 Calculated composition, % of diet SID Lys 1.18 1.18 SID Met 0.32 0.32 SID Thr 0.75 0.75 SID Trp 0.26 0.26 SID Cys + Met 0.62 0.62 β-mannan5 1.33 1.33 Analyzed composition, % of diet DM 86.75 87.17 GE, Mcal/kg 4.03 4.00 CP 22.28 21.83 EE4 4.02 3.97 Starch 29.34 30.89 NDF 12.17 11.83 ADF 6.97 6.79 Below detectable Endo-1,4-β-mannanase6, U/kg 150,000 limit 1Provided per kilogram of complete diet: 6,614 IU of vitamin A; 827 IU of vitamin D; 26 IU of vitamin E; 2.6 mg of vitamin K; 29.8 mg of niacin; 16.5 mg of pantothenic acid; 5.0 mg of riboflavin; 0.023 mg of vitamin B12. 2 Provided per kilogram of complete diet: Zn, 165 mg as ZnSO4; Fe, 165 mg as FeSO4; Mn, 39 mg as

MnSO4; Cu, 17 mg as CuSO4; I, 0.3 mg as Ca(IO3)2; and Se, 0.3 mg as Na2SeO3. 3HemicellTM HT-D, Elanco Animal Health, Guelph, ON, Canada; endo-1,4-β-mannanase (160 × 106 units/kg) from Paenibacillus alvei 4Acid hydrolyzed ether extract 5β-mannan concentration was calculated using values reported in Shastak et al. [70] 6endo-1,4-β-mannanase activity. One U = the amount of enzyme which generates 0.72 micrograms of reducing sugars per minute from a mannose-containing substrate at pH 7.0 and temperature of 40°C.

Table 3. Complete blood count values in young weaned pigs fed a diet with or without - mannanase1,2

3 4 Control diet -mannanase diet Contrast Treatment Estimate SEM Estimate SEM P-value

Cell type count

(cells x109/L)5 Total cells 468.6 34.9 432.4 23.8 0.405 WBC 24.50 1.43 21.51 0.98 0.104 Neut 6.47 0.94 6.83 0.64 0.758 Bands 0.200 0.050 0.266 0.034 0.504 Eos 0.202 0.099 0.300 0.068 0.599 Baso 0.048 0.061 0.100 0.042 0.451 Lymph 15.35 1.46 12.46 1.00 0.124 Mono 0.727 0.181 0.709 0.123 0.934 RBC 7.44 0.14 7.55 0.09 0.547 1Blood was collected on d 8 of the experiment 6 h post-feeding, prior to the immune challenge beginning on d 10. 2n = 10 pigs per treatment 3Control diet was a corn, soybean mean, and soy hulls based diet containing 1.33% β-mannans and did not contain β-mannanase enzyme. Pigs on the control (CON) treatment were fed the control diet and estimates are representative of the CON treatment only. 4Enzyme diet was the control diet supplemented with 0.056% β-mannanase (HemicellTM HT-D, Elanco Animal Health, Guelph, ON, Canada; endo-1,4-β-mannanase (160 × 106 units/kg) from Paenibacillus alvei). Pigs on the enzyme (ENZ) and immune system stimulation (ISS) treatments were fed the enzyme diet. Estimates are representative of the ENZ and ISS treatments prior to immune stimulation. 5Basophils (Baso); eosinophils (Eos); immature neutrophils (Bands); lymphocytes (Lymph); mature neutrophils (Neut); monocytes (Mono); white blood cells (WBC)

Table 4. Effect of -mannanase on pig serum glucose, insulin, acute phase protein, and cytokine concentrations1,2

3 4 Control diet Enzyme diet Contrast Treatment Estimate SEM Estimate SEM P-value Glucose, mmol/L 7.70 0.46 7.41 0.25 0.585 Insulin, pmol/L 88.92 10.99 83.35 6.06 0.664 Insulin:Glucose 11.56 1.21 11.08 0.67 0.734 Acute phase protein, mg/mL Haptoglobin 1.57 0.22 1.02 0.14 0.050 MBL5 125.4 8.6 117.5 5.2 0.445 Cytokine, pg/mL6 GM-CSF 17.01 13.01 14.56 7.85 0.874 IL-1 35.01 6.81 7.50 4.27 0.004 IL-1 1064 598 1138 361 0.917 IL-1ra 428.6 257.5 673.7 161.5 0.435 IL-2 328.4 157.6 228.8 95.0 0.596 IL-4 995.5 545.6 711.6 329.0 0.662 IL-6 190.2 61.5 98.72 38.59 0.230 IL-8 263.1 95.4 402.2 57.5 0.230 IL-10 499.4 159.1 332.9 95.9 0.383 IL-12 1570 173 1730 104 0.439 IL-18 1994 546 1380 329 0.350 TNF 24.29 39.90 52.30 25.02 0.563 1Blood was collected on d 8 of the experiment 6 h post-feeding, prior to the immune challenge beginning on d 10. 2n = 10 pigs per treatment 3Control diet was a corn, soybean mean, and soy hulls based diet containing 1.33% β-mannans and did not contain β-mannanase enzyme. Pigs on the control (CON) treatment were fed the control diet and estimates are representative of the CON treatment only. 4Enzyme diet was the control diet supplemented with 0.056% β-mannanase (HemicellTM HT-D, Elanco Animal Health, Guelph, ON, Canada; endo-1,4-β-mannanase (160 × 106 units/kg) from Paenibacillus alvei). Pigs on the enzyme (ENZ) and immune system stimulation (ISS) treatments were fed the enzyme diet. Estimates are representative of the ENZ and ISS treatments prior to immune stimulation. 5Mannose binding lectin A (MBL) 6Granulocyte-macrophage colony-stimulating factor (GM-CSF); interleukin-1α (IL-1α); interleukin-1 (IL-1); interleukin-1 receptor antagonist (IL-1ra); interleukin-2 (IL-2); interleukin-4 (IL-4); interleukin-6 (IL-6); interleukin-8 (IL-8); interleukin-10 (IL-10); interleukin-12 (IL-12); interleukin-18 (IL-18); tumor necrosis factor alpha (TNFα)

Table 5. Apparent total tract digestibility (ATTD) in pigs on the control, enzyme, or immune system stimulation treatment1

Item CON2 ENZ3 ISS4 SEM Treatment P-value

ATTD5, %

DM 88.05 87.36 87.66 0.33 0.356 GE 87.35 86.73 86.86 0.33 0.418 CP 87.59 86.47 86.77 0.44 0.212 EE6 70.41 69.62 66.85 1.18 0.122 Starch 99.41 99.50 99.56 0.10 0.565 NDF 68.81 66.11 70.23 1.55 0.178 ADF 71.65 68.04 74.13 2.26 0.175 Hemicellulose7 65.00 63.51 64.99 0.92 0.429 1n = 10 pigs per treatment 2Control treatment (CON) = pigs fed basal diet (0.0% β-mannanase) with saline injection 3Enzyme treatment (ENZ) = pigs fed enzyme diet (0.056% β-mannanase) with saline injection 4Immune system stimulation treatment (ISS) = pigs fed enzyme diet (0.056% β-mannanase) with LPS (Escherichia coli serotype O55:B5) injection 5ATTD, % = [(nutrient intake (kg) – fecal nutrient output (kg)) / nutrient intake (kg)] × 100 6Acid hydrolyzed ether extract 7Hemicellulose = NDF - ADF

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Table 6. Growth performance and nitrogen balance in pigs on control, enzyme, or immune system stimulation treatment1

Item CON2 ENZ3 ISS4 SEM Treatment P-value Body Weight, kg

d 0 10.23 10.21 10.38 0.15 0.651 d 16 15.25 15.26 14.86 0.27 0.471 ADG d 1-16, g/d 313.9 316.0 279.7 13.6 0.128 Nitrogen (N) balance, g/d

Intake 21.02a 20.93a 17.38b 0.95 0.021 Excreted 6.21 7.08 6.24 0.45 0.318 in feces 2.62a 2.83a 2.27b 0.11 0.007 in urine 3.59 4.25 3.97 0.47 0.624 Retained 14.81a 13.85ab 11.14b 0.99 0.045 % of excreted N in feces 41.31 41.21 38.42 3.29 0.778 % of excreted N in urine 58.69 58.79 61.58 3.29 0.778 Nitrogen balance, % of intake Excreted in feces 12.41 13.54 13.23 0.44 0.212 Excreted in urine 17.38 21.40 24.56 4.14 0.495 a,bWithin a row, treatment means without a common superscript differ, P < 0.05. 1n = 10 pigs per treatment 2Control treatment (CON) = pigs fed basal diet (0.0% β-mannanase) with saline injection 3Enzyme treatment (ENZ) = pigs fed enzyme diet (0.056% β-mannanase) with saline injection 4Immune system stimulation treatment (ISS) = pigs fed enzyme diet (0.056% β-mannanase) with LPS (Escherichia coli serotype O55:B5) injection

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Table 7. Effect of treatment on energy balance, respiratory quotient, maintenance energy requirements, and nutrient deposition1 Treatment P-value Item CON2 ENZ3 ISS4 SEM d 16 BW, kg 14.37 14.19 13.77 0.26 0.313 d 16 DMI, kg 0.51 0.51 0.46 0.03 0.348 0.60 Energy, kcal/kg BW /kg DMI/d ME intake 771.3 755.9 751.1 37.7 0.924 5 Heat production b b a HPtotal 278.8 274.9 333.0 14.9 0.040

FHPtotal 287.8 276.0 324.3 17.1 0.178 FHP 207.8 206.6 243.3 12.9 0.135 Retained energy6

As protein 197.5 173.6 191.0 18.3 0.627 As lipid 291.4a 302.9a 227.7b 19.2 0.046 Total 488.9 476.5 418.7 32.0 0.318

kmg, % 87.07 86.44 83.01 1.34 0.130 b b a Estimated MEm 239.0 239.5 295.5 15.3 0.045 Retained energy, % of ME intake

As protein 25.68 23.15 24.99 1.27 0.354 As lipid 37.77a 40.07a 29.81b 2.02 0.013 Total 63.44a 63.22a 54.80b 2.18 0.033 Respiratory quotient

Fed state 0.92 0.90 0.88 0.01 0.225 Fasting state 0.74 0.73 0.73 0.01 0.381 Nutrient deposition7, g/d

As protein 87.74 78.55 69.80 5.86 0.150 As lipid 76.22a 79.43a 55.45b 6.21 0.047 ADG d 10-16, g/d 447.1a 404.8a 330.7b 21.3 0.010 a,bWithin a row, treatment means without a common superscript differ, P < 0.05. 1n = 7 pigs per treatment (CON and ENZ) and 6 pigs per treatment (ISS) 2Control treatment (CON) = pigs fed basal diet (0.0% β-mannanase) with saline injection 3Enzyme treatment (ENZ) = pigs fed enzyme diet (0.056% β-mannanase) with saline injection 4Immune system stimulation treatment (ISS) = pigs fed enzyme diet (0.056% β-mannanase) with LPS (Escherichia coli serotype O55:B5) injection 5 Heat production (HP) = (3.87 × O2 consumption (L) + 1.20 × CO2 production (L) – 1.43 × urinary 0.60 N)/BW (kg) [22]; Total HP (HPtotal)= avg. HP over 22 h fed state, post- challenge; Total fasting HP

(FHPtotal) = avg. HP over 12 h fasted state; Fasting HP (FHP) = avg. of 10 lowest HP values over the 12 h fasted state [27,28]; HP10 = avg. HP over 10 h post-challenge (1000 h – 2000 h), fed state; HPlow = avg. of 10 lowest HP values over 10 h post-challenge (1000 h – 2000 h), fed state; Activity HP (AHP) = HP10 -

HPlow; Thermic effect of feeding (TEF) = HP10 - AHP – FHP; Heat increment (HI) = AHP + TEF; ME efficiency for maintenance and growth (kmg) = (1 – HI) × 100 [25]; ME used for maintenance (MEm) =

FHP × 100/kmg [25] 6 Retained energy (RE) = ME intake – total fed-state HP, pre-and post-challenge [29]; RE as protein (REp) 0.60 = [PD (g) × 5.66 (kcal/g)]/ BW /DMI [30]; RE as lipid (REl) = RE - REp [30] 7 Protein deposition (PD) nitrogen retention (g) × 6.25; Lipid deposition = REl (kcal) / 9.49 (kcal/g) [30]

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Table 8. Dietary energy values and efficiency in pigs on control, enzyme, or immune system stimulation treatment1 Treatment Treatment Item CON2 ENZ3 ISS4 SEM P-value 5 Dietary energy value , Mcal/kg DM 4.65 4.59 4.59 GE DE 4.07 3.99 4.00 0.02 0.051 ME 3.96 3.86 3.89 0.03 0.052 NE 3.29 3.30 3.11 0.19 0.748 ME/DE efficiency, % 97.31 96.92 97.35 0.37 0.457 NE/ME efficiency, % 83.09 85.31 80.03 4.42 0.701 1n = 7 pigs per treatment (CON and ENZ) and 6 pigs per treatment (ISS) 2Control treatment (CON) = pigs fed basal diet (0.0% β-mannanase) with saline injection 3Enzyme treatment (ENZ) = pigs fed enzyme diet (0.056% β-mannanase) with saline injection 4Immune system stimulation treatment (ISS) = pigs fed enzyme diet (0.056% β-mannanase) with LPS (Escherichia coli serotype O55:B5) injection 5Gross energy (GE) analyzed via bomb calorimetry; digestible energy (DE) = GE apparent total tract digestibility coefficient × diet GE; metabolizable energy (ME) = DE – (urinary energy + 0.4% of DE intake); net energy (NE) = (retained energy + fasting heat production)/DMI.

103

42.0

41.5 Injection 41.0

40.5

C CON

∘

, e

r 40.0

t a

r 39.5 e

p ENZ m

e 39.0 T 38.5

38.0 ISS 37.5

37.0 5 8 10 12 14 Experiment Day

Figure 1. Effect of treatment on young weaned pig (n = 10 per treatment) rectal temperature (C). Control treatment (CON) = pigs fed basal diet (0.0% β-mannanase) with saline injection. Enzyme treatment (ENZ) = pigs fed enzyme diet (0.056% β-mannanase) with saline injection. Immune system stimulation treatment (ISS) = pigs fed enzyme diet (0.056% β-mannanase) with LPS (Escherichia coli serotype O55:B5) injection. The arrows indicate days on which either a saline (CON and ENZ) or LPS (ISS) injection were administered at 1000 h. Rectal temperatures were measured 4 h post-challenge. Data points on d 5 and 8 represent average baseline pre- challenge temperature, and d 10, 12, and 14 represent post-challenge temperatures. Treatment by day interaction P < 0.0001; day P < 0.0001; treatment P < 0.0001; block P = 0.0015.

104

White blood cells Neutrophils 30 10

a a 9 a 25 a ab 8 ab a a b a 7

L /

/ 6

15 x 5

l e

e 4 b

C C 10 c 3

2 5 1

0 0 CON ENZ ISS CON ENZ ISS Treatment Treatment

Lymphocytes Monocytes 1.4 18 a a 16 a 1.2

14 ab ab b 1

12 ab ab ab L

/ 0.8 b

L /

0 10

x 0.6 l

C e 6 c C 0.4 4 0.2 c 2 0 0 CON ENZ ISS CON ENZ ISS -0.2 Treatment Treatment

Figure 2. Effect of treatment on complete blood count before (d 8) and after (d 10) challenge. Serum was collected at 1400 h each day (4 h post- challenge). Control treatment (CON) = pigs fed basal diet (0.0% β-mannanase) with saline injection. Enzyme treatment (ENZ) = pigs fed enzyme diet (0.056% β-mannanase) with saline injection. Immune system stimulation treatment (ISS) = pigs fed enzyme diet (0.056% β-mannanase) with LPS (Escherichia coli serotype O55:B5) injection. n=10 per treatment. a,bWithin a graph, bars without a common superscript differ, P < 0.05.

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Granulocyte-macrophage colony-stimulating factor Interleukin-1! (IL-1! ) (GM-CSF) a 60 a 2800

50 2300 40

1800 b

30 ab L b

L ab

m /

m b g

/ b

g b 1300 p 20 b b 800 b 10

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Interleukin-1 receptor antagonist (IL-1ra) Interleukin-6 (IL-6) a a 110000 1400

1200 90000 1000

70000

800

m /

/ b

g g 600

50000 p p b 400 b b 30000 b 200

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700

1900 600

500 1400

L 400

g p p 300 900 b b 200 b b b b b b 400 100 b b 0 CON ENZ ISS -100 CON ENZ ISS -100 Treatment Treatment

Figure 3. Effect of treatment on serum cytokine concentrations before (d 8) and after (d 10) challenge. Serum was collected at 1400 h each day (4 h post- challenge). Control treatment (CON) = pigs fed basal diet (0.0% β-mannanase) with saline injection. Enzyme treatment (ENZ) = pigs fed enzyme diet (0.056% β-mannanase) with saline injection. Immune system stimulation treatment (ISS) = pigs fed enzyme diet (0.056% β-mannanase) with LPS (Escherichia coli serotype O55:B5) injection. n=10 per treatment. a,bWithin a graph, bars without a common superscript differ, P < 0.05.

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CHAPTER III

BETA-MANNANASE SUPPLEMENTATION EFFECTS ON NURSERY PIG

GROWTH PERFORMANCE AND SERUM ACUTE PHASE PROTEIN

CONCENTRATIONS

Modified from a paper submitted to the Journal of Animal Science

N. F. Huntley*, S. A. Gould*, J. F. Patience*

*Department of Animal Science, Iowa State University, Ames, 50011, USA

Abstract

Dietary -mannans resemble carbohydrate structures on pathogen surfaces that may activate the innate immune system. Beta-mannanase supplementation has been proposed to prevent potential immune stimulation and spare energy for growth. The objective of this experiment was to evaluate the effects of dietary mannan level, - mannanase supplementation, and their interaction on growth performance, feed efficiency, and serum acute phase protein (APP) concentrations in nursery pigs. Pigs (n =

480, 10 pigs per pen) were blocked by initial BW and pens (n = 12 per treatment) were randomly assigned to 1 of 4 treatments in a 2  2 factorial arrangement for a 28-d experiment with 3 phases. Two levels of dietary mannan were achieved by replacing part of the corn and soybean meal in the low mannan diet (0.4% -mannan) with copra meal for a high mannan diet (2.8% -mannan). Each diet was fed with and without 0.05% endo-1,4--mannanase (Hemicell HT 1.5, Elanco Animal Health). Serum was collected

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(1 pig/pen) for haptoglobin and C-reactive protein (CRP) analysis prior to treatment initiation (d 0) and on d 28. Data were analyzed as a 2 x 2 factorial design with main effects of mannan level and -mannanase supplementation. There were no significant interactions between mannan level and -mannanase supplementation overall (d 0 to 28;

P = 0.852) or within each phase (P ≥ 0.106). Overall, high mannan diets decreased ADG

(0.37 vs. 0.38 kg/d; P = 0.027) and ADFI (0.52 vs. 0.54 kg/d; P = 0.024) compared to low mannan diets. There were no significant -mannanase effects on growth performance or feed efficiency (P > 0.10). Serum APP concentrations were similar among treatments at baseline. At d 28 haptoglobin concentrations were lower (P ≤ 0.0001) and CRP concentrations were greater (P ≤ 0.0001) compared to baseline values, but neither was affected by mannan level or -mannanase supplementation (P ≥ 0.160). Beta-mannanase supplementation tended to increase serum CRP in the high mannan diet (290.64 g/ml) compared to the low diet (243.49 g/ml; P = 0.0791) with no differences compared to when the enzyme was not supplemented (avg. = 273.91 g/ml). In conclusion, dietary - mannan concentrations as high as 2.8% did not induce a systemic innate immune response in nursery pigs. The levels of -mannan found in practical, commercial-type swine diets are not likely to induce an immune response and supplementation of - mannanase for this purpose is not warranted.

Introduction

Certain dietary carbohydrate components, such as β-mannans, have been hypothesized to induce innate immune system activation by mimicking bacterial cell wall structures [1–3]. Beta-mannans are found in feed ingredients such as soybean meal

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(SBM), palm kernel meal, and copra meal in varying concentrations [4]. If feed-induced immune stimulation occurs it may decrease dietary energy partitioning toward productive processes such as growth [5], and exogenous β-mannanase enzymes have been developed to theoretically inhibit this innate immune stimulation. In poultry, β-mannanase supplementation decreased serum acute phase proteins (APP), thus ameliorating systemic inflammatory stress in broilers [6]. However, this response has yet to be demonstrated in pigs.

Beta-mannanase supplementation responses in pigs have been variable [7–10].

Previous research in our laboratory evaluated β-mannanase supplementation in a corn,

SBM, and soy hulls-based diet and found no effects on diet digestibility, nitrogen balance, maintenance energy requirements, or systemic innate immune response variables

[5]. Thus, the results did not support the hypothesis that β-mannanase supplementation ameliorates an innate immune response induced by soybean β-mannans at a concentration of 1.3% of the diet. It is possible, however, that this concentration was too low to either effectively stimulate a systemic inflammatory response or to be able to measure the effects of β-mannanase supplementation. To test this hypothesis, the objectives of this study were to evaluate the impact and interactions of dietary β-mannan concentration and

β-mannanase supplementation on serum APP, as indicators of systemic immune activation, and on growth performance in nursery pigs.

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Materials and methods

All experimental procedures adhered to guidelines for the ethical and humane use of animals for research and were approved by the Iowa State University Institutional

Animal Care and Use Committee (IACUC #5-16-8263-S).

Animals, housing, and experimental design

A total of 480 weanling pigs (6.6 ± 0.4 kg BW; Genetiporc 6.0 × Genetiporc F25,

PIC, Inc., Hendersonville, TN) were purchased and transported to the Iowa State

University Swine Nutrition farm. Upon arrival, pigs were individually weighed, ear- tagged, and vaccinated for porcine reproductive and respiratory syndrome virus, and

Escherichia coli. Pigs were blocked by initial weight and pens were randomly assigned to

1 of 4 dietary treatments. There were 10 pigs per pen and 12 pens per treatment. Sexes were not separated but there were the same number of barrows and gilts per treatment within each block.

Diets and feeding

Pens (1.2 m × 2.4 m) had a four-space self-feeder and 2 nipple waterers to provide ad libitum access to feed and water. Pigs were fed a common diet for 7 d post weaning; the experimental diets were then fed in 3 phases over 28 d. Phases 1 and 2 each were offered for 7 d each and phase 3 for 14 d. Diets were formulated to meet or exceed NRC

(2012) nutrient recommendations and were formulated to provide equal ME:SID lysine concentrations. Treatments were arranged as a 2 × 2 factorial of dietary -mannan level

(low or high) and -mannanase supplementation (positive or negative). Two levels of dietary mannan were achieved by replacing a proportion of the corn and soybean meal in the low mannan diet (0.4% -mannan; Table 1) with copra meal for a high mannan diet

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(2.8% -mannan; Table 2). Diet -mannan concentration was calculated based on ingredient -mannan concentrations [12–14]. Beta-mannanase (Hemicell HT-1.5, Elanco

Animal Health, Greenfield, IN; endo-1,4-β-mannanase (EC 3.2.1.78; 160 x 106 units/kg) from Bacillus lentus) was supplemented at 0.05% of the diet. One β-mannanase unit of activity is defined as the amount of enzyme that generates 0.72 g of reducing sugars per min from a mannose-containing substrate at pH 7.0 and 40C.

Sample collection

Pig BW and feed intake were measured on d 0, 7, 14, 21, and 28 of the experiment to calculate ADG, ADFI, and G:F for each phase. On d 0 (baseline) and d 28 serum was collected from 1 barrow per pen (the same pig at each time point) via jugular venipuncture. Blood was allowed to clot, and serum was separated by centrifugation

(2000 × g for 15 min at 4°C), collected and divided into two subsamples, and stored at -

80°C until analysis. Diet subsamples were collected during mixing.

Analytical methods

Diets were ground to 1 mm particle size and analyzed in duplicate for DM

(method 930.15) and acid-hydrolyzed ether extract (aEE; method 2003.06) using standard methods [15]; and in triplicate for NDF [16] and ADF [17]. Diets were analyzed in duplicate for nitrogen (N; method 990.03 [15]; TruMac; LECO Corp., St. Joseph, MI).

An EDTA sample (9.56% N) was used as the standard for calibration and was determined to contain 9.55 ± 0.01% N. Crude protein was calculated as N  6.25. Gross energy was determined in duplicate using an isoperibol bomb calorimeter (model 6200; Parr

Instrument Co., Moline, IL). Benzoic acid (6318 kcal GE/kg; Parr Instrument Co.) was used as the standard for calibration and was determined to contain 6316 ± 0.9 kcal

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GE/kg. Diets were analyzed in duplicate for β-mannanase concentration (colorimetric determination, Elanco Animal Health, Gaithersburg, MD).

Serum subsamples were analyzed in duplicate for haptoglobin and C-reactive protein (CRP) using porcine specific commercially available kits according to the manufacturer’s instructions (Immunology Consultants Laboratory, Inc., Portland, OR).

The intra- and inter-assay coefficients of variation for haptoglobin and CRP were 4.9 and

16.0%, and 3.6 and 9.9%, respectively.

Statistical analysis

Data were analyzed as a 2 × 2 factorial in a randomized complete block design to test the main effects and interaction of dietary mannan level and -mannanase supplementation. Data were analyzed with pen as the experimental unit and block as a random variable using the MIXED procedure of SAS (SAS Institute Inc., Cary, NC).

Differences among treatments were determined using ANOVA and means were separated using the least square means statement and the PDIFF option. Serum APP data were log transformed and analyzed as repeated measures with compound symmetry covariance structure. Differences were considered significant if P was ≤ 0.05 and a tendency if P was

> 0.05 and ≤ 0.10.

Results

Growth performance

There were no significant interactions between mannan level and -mannanase supplementation overall (d 0 to 28) or within each phase (Table 6). Pig BW did not differ among treatments on d 0, 7, or 14, but pigs fed the low mannan diets, independent of -

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mannanase inclusion, tended to weigh more on d 21 and 28 (Table 6). This tendency for

BW differences during phase 3 was driven by increased ADG in the low mannan treatments during d 15 – 21 (P = 0.015; Table 6). During this time, feed intake was also greater (P = 0.007) for pigs fed low mannan diets, corresponding to no differences in G:F

(P = 0.834; Table 6).

Overall (d 0 – 28), pigs fed low mannan diets had greater ADG and ADFI compared to pigs fed high mannan diets (P ≤ 0.027) and feed efficiency did not differ (P

= 0.653; Table 6). Beta-mannanase inclusion did not impact BW or growth performance throughout the entire experiment or within each phase (P ≥ 0.106; Table 6).

Acute phase proteins

Serum APP concentrations were similar among treatments at baseline (P ≥ 0.20).

On d 28, haptoglobin concentrations were lower (P ≤ 0.0001; Fig. 1) and CRP concentrations were greater (P ≤ 0.0001; Fig. 2) compared to baseline values, but neither were affected by mannan level or -mannanase supplementation (P ≥ 0.160). Beta- mannanase supplementation tended to increase serum CRP in the high mannan diet

(290.64  20.10 g/ml) compared to the low mannan diet (243.49  20.10 g/ml) with no differences compared to when the enzyme was not supplemented (avg. = 273.91  20.10

g/ml; P = 0.0791).

Discussion

This experiment evaluated the interaction of dietary -mannan level and - mannanase supplementation on nursery pig growth performance and serum acute phase protein concentrations and tested the hypotheses that dietary -mannans induce an innate

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immune response in nursery pigs and that -mannanase supplementation alleviates this response [6,18]. Understanding if, or how, this response occurs in pigs is important because a perceived immune challenge can partition energy and nutrients away from productive processes such as muscle growth and negatively impact the efficiency and cost of meat production [5,19,20]. In the face of a true pathogen challenge, this is a desirable response to maintain animal health and survival. However, β-mannan in soybean, copra, and palm kernel meals resemble carbohydrate structures on pathogen surfaces [21] and may induce an unproductive and undesirable innate immune response, termed a feed-induced immune response (FIIR).

Beta-mannans are components of hemicellulose in the plant cell wall and are linear polysaccharides comprised of repeating β-(1,4)-linked D-mannose units with galactose and/or glucose substitutions [22]. Specifically, galactomannans (α-(1,6)-linked

D-galactose residues attached to mannose residues within the β-(1,4)-mannan backbone) are the type of mannans in the endosperm of legume plant seeds such as soybeans [22].

As a constituent of hemicellulose, β-mannans are not digested by mammalian endogenous enzymes. Thus, intact β-mannans may be available to bind carbohydrate recognition domains of pattern recognition receptors on innate immune cells surveying the intestinal epithelium for potential pathogens [1,23,24]. It is hypothesized that in this way, β-mannan may be capable of stimulating innate immune cells resulting in a nonproductive, energy-draining, inflammatory immune response [2,3,6]. One component of innate immune activation is the acute phase response, which is characterized by dramatic changes in the concentration of APP, such as haptoglobin and CRP.

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The initial FIIR descriptions were in poultry and reported increased macrophage and lymphocyte proliferation [25] and plasma APP concentrations [6] due to increasing dietary SBM inclusion. This response was then attenuated by the addition of - mannanase and was supported by improvements in feed efficiency [6,25]. It was hypothesized that the hydrolyzed manno-oligosaccharides no longer retained the degree of polymerization necessary to stimulate and cross-link multiple mannose receptors, thus reducing innate immune stimulation.

However, there is a paucity of data regarding immune responses of pigs when dietary -mannan and -mannanase are supplemented, and a -mannan-derived FIIR has yet to be demonstrated. This study provides novel data on the interaction of dietary β- mannan level and β-mannanase enzyme supplementation on nursery pig serum APP concentrations, as a measure of systemic immune stress. If an immune response to β- mannan had occurred, both CRP and haptoglobin concentrations would have been expected to increase substantially [26]. The main effect of dietary mannan level did not impact concentrations of either APP, indicating that an acute phase response was not detected in pigs fed the high mannan diet. The changes in APP concentrations from baseline to d 28 were likely due to time, age of the pig, or other environmental factors that equally affected each treatment.

Similarly, there was no interaction between the effects of mannan level and - mannanase supplementation on serum ACP concentrations. Had an innate immune response occurred in either or both diets, it would have been expected that -mannanase supplementation would prevent immune stimulation and decrease serum haptoglobin and

CRP, and this response would have been exaggerated in the high-mannan diets due to the

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greater amount of substrate for the enzyme. Thus, it was concluded that dietary -mannan concentrations as high as 2.8% did not induce a systemic innate immune response in nursery pigs. However, because diets with different -mannan concentrations were achieved by replacing corn and SBM with copra meal, a specific ingredient effect, separate from a -mannan nutrient effect, cannot be ruled out.

The results of this experiment corroborate data from Huntley et al. [5] which indicated that -mannanase supplementation did not impact nursery pig serum proinflammatory cytokines, haptoglobin, or mannose-binding lectin concentrations.

Furthermore, there were no differences in maintenance energy requirements between control and -mannanase supplemented pigs, supporting the conclusion that -mannanase did not prevent an energy-draining immune response. It is possible that a localized response may have occurred at the level of the intestines but was not detected systemically. However, if this did occur, the response did not impact energy partitioning or decrease growth or feed efficiency. Interpreted together, the data from Huntley et al.

[5] and the current study indicate that the levels of -mannan found in practical, commercial-type swine diets are not likely to induce a FIIR and supplementation of - mannanase for this purpose is not warranted.

While -mannan did not induce an innate immune response, increasing its inclusion in nursery diets certainly decreased ADFI and ADG in line with responses expected from feeding higher fiber diets [27]. Inclusion of high-fiber co-products in commercial swine diets is continually increasing and carbohydrase enzymes have been developed to reduce the detrimental effects of fiber on nutrient and energy digestibility, as well as improve growth and feed efficiency. For this purpose, -mannanase has had

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variable effects when supplemented in swine diets, and results seem to depend on pig age, diet ingredient composition, and enzyme inclusion level.

In this experiment, -mannanase supplementation did not affect growth performance or feed efficiency in either the high or low mannan diets. Similarly, other studies have reported no -mannanase effects on growth performance and feed efficiency

[7,10,28], or nutrient and energy digestibility [5,28,29]. Yet, others have reported improvements in these measurements due to -mannanase supplementation [8–10] and data from Kim et al. [30] indicate that -mannanase supplementation effectively improved DM, GE, and -mannan digestibility as well as growth performance and feed efficiency, regardless of the amount of -mannan in the diet. More research is needed to better understand the circumstances in which -mannanase supplementation is effective.

Furthermore, data from this experiment and others do not support the hypothesis that - mannanase prevents a -mannan-induced immune response.

Conclusions

In conclusion, the high mannan diet decreased ADFI and thus, decreased ADG by

2.7% compared to the low mannan diet with no apparent effect on serum haptoglobin or

CRP concentrations. This experiment provides immune data previously lacking in pig growth trials assessing -mannanase efficacy. Beta-mannanase supplementation had no impact on growth performance and did not affect haptoglobin or CRP concentrations differently in high compared to low mannan diets. Thus, the data do not support the hypothesis that the level of -mannan found in practical swine diets induces an innate immune response or that -mannanase supplementation may prevent such a response.

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Table 1. Ingredient composition (as-fed basis) of the low mannan diets for phases 1 - 31 Ingredient, % Phase 1 Phase 2 Phase 3 Corn 51.69 51.04 49.75 Reduced-oil corn DDGS 10.00 12.50 15.00 Fish meal 5.00 4.00 3.00 Soybean meal 25.00 25.00 25.00 Soybean oil 4.20 3.50 3.20 Limestone 1.10 1.15 1.20 Monocalcium phosphate 0.60 0.55 0.55 L-Lysine-HCl 0.50 0.43 0.45 DL-Methionine 0.08 0.06 0.10 L-Threonine 0.15 0.15 0.11 L-Tryptophan 0.05 0.04 0.04 L-Valine 0.05 0.00 0.00 Vitamin premix2 0.25 0.25 0.25 Trace mineral premix3 0.20 0.20 0.20 Denagard4 0.18 0.18 0.00 Aureomycin5 0.40 0.40 0.00 Mecadox 2.56 0.00 0.00 0.20 Salt 0.50 0.50 0.50 Choline chloride, 60% 0.05 0.05 0.05 Titanium dioxide 0.00 0.00 0.40 For the enzyme positive diet, -mannanase (Hemicell HT 1.5, Elanco Animal Health, Greenfield, IN; endo-1,4-β-D-mannanase from Bacillus lentus with not less than 237 million units per kg of product) was included at 0.05% at the expense of corn. 1Phase 1 = d 0 – 7; Phase 2 = d 8 – 14; Phase 3 = d 15 - 28 2Provided 6,614 IU vitamin A, 827 IU vitamin D, 26 IU vitamin E, 2.6 mg vitamin K, 29.8 mg niacin, 16.5 mg pantothenic acid, 5.0 mg riboflavin, and 0.023 mg vitamin B12 per kg of diet. 3Provided 165 mg Zn (zinc sulfate), 165 mg Fe (iron sulfate), 39 mg Mn (manganese sulfate), 17 mg Cu (copper sulfate), 0.3 mg I (calcium iodate), and 0.3 mg Se (sodium selenite) per kg of diet. 4Denagard 10 (tiamulin, 0.22 g per kg), Elanco Animal Health, Greenfield, IN 5Aureomycin (chlortetracycline (0.44 g per kg diet), Zoetis Inc., Kalamazoo, MI 6Mecadox 2.5 (carbadox), Phibro Animal Health Corp., Ridgefield Park, NJ

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Table 2. Ingredient composition (as-fed basis) of the high mannan diets for phases 1 - 31 Ingredient, % Phase 1 Phase 2 Phase 3 Corn 45.79 45.30 44.23 Reduced-oil corn DDGS 10.00 12.50 15.00 Fish meal 5.00 4.00 3.00 Soybean meal 20.00 20.00 20.00 Copra meal 10.00 10.00 10.00 Soybean oil 4.80 4.10 3.70 Limestone 1.20 1.15 1.10 Monocalcium phosphate 0.70 0.55 0.50 L-Lysine-HCl 0.60 0.55 0.57 DL-Methionine 0.07 0.07 0.09 L-Threonine 0.17 0.15 0.15 L-Tryptophan 0.06 0.05 0.06 L-Valine 0.03 0.00 0.00 Vitamin premix2 0.25 0.25 0.25 Trace mineral premix3 0.20 0.20 0.20 Denagard4 0.18 0.18 0.00 Aureomycin5 0.40 0.40 0.00 Mecadox 2.56 0.00 0.00 0.20 Salt 0.50 0.50 0.50 Choline chloride, 60% 0.05 0.05 0.05 Titanium dioxide 0.00 0.00 0.40 For the enzyme positive diet, -mannanase (Hemicell HT 1.5, Elanco Animal Health, Greenfield, IN; endo-1,4-β-D-mannanase from Bacillus lentus with not less than 237 million units per kg of product) was included at 0.05% at the expense of corn. 1Phase 1 = d 0 – 7; Phase 2 = d 8 – 14; Phase 3 = d 15 - 28 2Provided 6,614 IU vitamin A, 827 IU vitamin D, 26 IU vitamin E, 2.6 mg vitamin K, 29.8 mg niacin, 16.5 mg pantothenic acid, 5.0 mg riboflavin, and 0.023 mg vitamin B12 per kg of diet. 3 Provided 165 mg Zn (zinc sulfate), 165 mg Fe (iron sulfate), 39 mg Mn (manganese sulfate), 17 mg Cu (copper sulfate), 0.3 mg I (calcium iodate), and 0.3 mg Se (sodium selenite) per kg of diet. 4Denagard 10 (tiamulin, 0.22 g per kg), Elanco Animal Health, Greenfield, IN 5Aureomycin (chlortetracycline (0.44 g per kg diet), Zoetis Inc., Kalamazoo, MI 6Mecadox 2.5 (carbadox), Phibro Animal Health Corp., Ridgefield Park, NJ

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Table 3. Phase 1 diet formulated and analyzed energy and nutrient composition (as-fed basis) Low mannan diet High mannan diet - enzyme + enzyme - enzyme + enzyme Formulated composition ME, Mcal/kg 3.46 3.46 3.46 3.46 SID amino acid, % Lys 1.38 1.38 1.37 1.37 Met 0.42 0.42 0.40 0.40 Total sulfur AA 0.74 0.74 0.74 0.74 Thr 0.83 0.83 0.81 0.81 Trp 0.26 0.26 0.26 0.26 Ca, % 0.81 0.81 0.85 0.85 STTD P, % 0.41 0.41 0.44 0.44 Analyzed composition DM, % 90.43 89.99 90.93 90.91 GE, Mcal/kg 4.16 4.16 4.32 4.33 CP, % 22.70 22.15 21.77 20.85 aEE1, % 7.62 7.41 8.78 8.37 NDF, % 9.21 8.88 12.90 13.01 ADF, % 3.58 3.47 5.04 5.89 < detection < detection -mannanase2, 3, U/kg 345,000 355,000 limit limit 1Acid-hydrolyzed ether extract (aEE) 2Endo-1,4-β-mannanase (EC 3.2.1.78); 1 U = the amount of enzyme which generates 0.72 g of reducing sugars per min from a mannose-containing substrate at pH 7.0 and 40C 3The limit of detection for feed samples was 45,000 U/kg

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Table 4. Phase 2 diet formulated and analyzed energy and nutrient composition (as-fed basis) Low mannan diet High mannan diet - enzyme + enzyme - enzyme + enzyme Formulated composition ME, Mcal/kg 3.42 3.42 3.42 3.42 SID amino acid, % Lys 1.30 1.30 1.30 1.30 Met 0.40 0.40 0.40 0.40 Total sulfur AA 0.71 0.71 0.71 0.71 Thr 0.82 0.82 0.79 0.79 Trp 0.25 0.25 0.24 0.24 Ca, % 0.78 0.78 0.77 0.77 STTD P, % 0.39 0.39 0.40 0.40 Analyzed composition DM, % 89.50 89.79 90.82 90.88 GE, Mcal/kg 4.10 4.11 4.23 4.24 CP, % 21.52 21.88 21.06 20.57 aEE1, % 6.61 7.19 8.59 8.53 NDF, % 9.62 9.91 13.89 14.43 ADF, % 3.04 3.19 4.92 4.92 < detection < detection 304,000 316,000 -mannanase2,3, U/kg limit limit 1Acid-hydrolyzed ether extract (aEE) 2Endo-1,4-β-mannanase (EC 3.2.1.78); 1 U = the amount of enzyme which generates 0.72 g of reducing sugars per min from a mannose-containing substrate at pH 7.0 and 40C 3The limit of detection for feed samples was 45,000 U/kg

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Table 5. Phase 3 diet formulated and analyzed energy and nutrient composition (as-fed basis) Low mannan diet High mannan diet - enzyme + enzyme - enzyme + enzyme Formulated composition ME, Mcal/kg 3.39 3.39 3.39 3.39 SID amino acid, % Lys 1.28 1.28 1.28 1.28 Met 0.43 0.43 0.41 0.41 Total sulfur AA 0.72 0.72 0.70 0.70 Thr 0.78 0.78 0.78 0.78 Trp 0.25 0.25 0.25 0.25 Ca, % 0.75 0.75 0.70 0.70 STTD P, % 0.38 0.38 0.38 0.38 Analyzed composition DM, % 90.60 90.35 90.94 91.15 GE, Mcal/kg 4.17 4.15 4.27 4.27 CP, % 21.60 21.61 21.72 21.11 aEE1, % 6.98 6.99 8.16 8.47 NDF, % 10.38 10.83 14.87 15.00 ADF, % 3.27 3.39 5.70 5.80 < detection < detection limit 327,000 365,000 -mannanase2,3, U/kg limit 1Acid-hydrolyzed ether extract (aEE) 2Endo-1,4-β-mannanase (EC 3.2.1.78); 1 U = the amount of enzyme which generates 0.72 g of reducing sugars per min from a mannose-containing substrate at pH 7.0 and 40C 3The limit of detection for feed samples was 45,000 U/kg

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Table 6. Effects of dietary mannan level and -mannanase supplementation on nursery pig BW, growth rate, feed intake, and efficiency1 Enzyme Mannan level1 SEM P-value inclusion2 High Low - + ME Mannan Enzyme Overall d 0 BW, kg 6.64 6.65 6.64 6.65 0.36 0.583 0.736 0.790 d 28 BW 17.05 17.43 17.14 17.33 0.67 0.954 0.051 0.317 ADG, kg 0.37 0.38 0.37 0.38 0.01 0.852 0.027 0.354 ADFI, kg 0.52 0.54 0.53 0.53 0.02 0.922 0.024 0.922 G:F 0.71 0.71 0.70 0.71 0.01 1.000 0.653 0.106 d 0 - 7 ADG, kg 0.11 0.12 0.12 0.12 0.01 0.364 0.221 0.629 ADFI, kg 0.17 0.18 0.17 0.17 0.01 0.576 0.100 0.779 G:F 0.69 0.71 0.69 0.71 0.02 0.357 0.427 0.570 d 8 - 14 initial BW 7.44 7.53 7.47 7.50 0.38 0.396 0.202 0.680 ADG, kg 0.34 0.34 0.34 0.34 0.01 0.169 0.586 0.952 ADFI, kg 0.40 0.41 0.41 0.40 0.02 0.106 0.181 0.699 G:F 0.84 0.83 0.83 0.84 0.01 0.789 0.164 0.383 d 15 - 21 initial BW 9.79 9.90 9.84 9.86 0.46 0.203 0.254 0.859 ADG, kg 0.50 0.53 0.51 0.52 0.01 0.935 0.015 0.463 ADFI, kg 0.67 0.70 0.68 0.68 0.02 0.528 0.007 0.795 G:F 0.75 0.76 0.75 0.76 0.01 0.645 0.834 0.706 d 22 - 28 initial BW 13.33 13.61 13.41 13.53 0.56 0.328 0.058 0.429 ADG, kg 0.53 0.55 0.53 0.54 0.02 0.267 0.111 0.267 ADFI, kg 0.87 0.89 0.87 0.88 0.03 0.168 0.198 0.687 G:F 0.61 0.61 0.61 0.62 0.01 1.000 0.603 0.351 1n=12 pens per treatment with 10 pigs per pen 2Two levels of dietary mannan were achieved by replacing a proportion of the corn and soybean meal in the low mannan diet (0.4% -mannan) with copra meal for a high mannan diet (2.8% -mannan) 3For the enzyme positive diet, -mannanase (Hemicell HT 1.5, Elanco Animal Health, Greenfield, IN; endo-1,4-β-D-mannanase from Bacillus lentus with not less than 237 million units per kg of product) was included at 0.05%

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Haptoglobin 1200

l 1000

�

n 800

r t

n high mannan + enzyme e

c 600 n

o low mannan + enzyme

i high mannan - enzyme b

o 400 l

g low mannan - enzyme

p a

h 200

0 baseline d 28 Time

Figure 1. Effects of dietary mannan level and -mannanase supplementation on nursery pig serum haptoglobin concentration (휇g/ml) at baseline (d 0) and after 28 d on the experimental diet. Two levels of dietary mannan were achieved by replacing a proportion of the corn and soybean meal in the low mannan diet (0.4% -mannan) with copra meal for a high mannan diet (2.8% -mannan). For the enzyme positive diet, -mannanase (Hemicell HT 1.5, Elanco Animal Health, Greenfield, IN; endo-1,4-β-D-mannanase from Bacillus lentus with not less than 237 million units per kg of product) was included at 0.05%. The main effect of time (baseline or d 28) was significant (P < 0.0001) but time did not have any significant interactions with the effects of mannan, enzyme, or the combination of the 2 (P ≥ 0.437). Dietary mannan level, -mannanase supplementation, or the interaction of the two effects did not significantly impact serum haptoglobin concentrations (P ≥ 0.160).

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C-reactive protein 450

400 l

m 350

g �

300

o i

t 250 high mannan + enzyme

t n

e 200 low mannan + enzyme

c n

o high mannan - enzyme

c 150

P low mannan - enzyme

R 100 C 50 0 baseline d 28 Time Figure 2. Effects of dietary mannan level and -mannanase supplementation on nursery pig serum C-reactive protein concentration (휇g/ml) at baseline (d 0) and after 28 d on the experimental diet. Two levels of dietary mannan were achieved by replacing a proportion of the corn and soybean meal in the low mannan diet (0.4% -mannan) with copra meal for a high mannan diet (2.8% -mannan). For the enzyme positive diet, -mannanase (Hemicell HT 1.5, Elanco Animal Health, Greenfield, IN; endo-1,4-β-D-mannanase from Bacillus lentus with not less than 237 million units per kg of product) was included at 0.05%. The main effect of time (baseline or d 28) was significant (P < 0.0001) but time did not have any significant interactions with the effects of mannan, enzyme, or the combination of the two (P ≥ 0.332). Neither dietary mannan level nor -mannanase supplementation significantly impacted serum C-reactive protein concentrations (P ≥ 0.201). The interaction of mannan level and enzyme inclusion tended to affect C-reactive protein concentrations (P = 0.079).

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CHAPTER IV

XYLOSE METABOLISM IN THE PIG

Modified from a paper submitted to PLOS ONE

N. F. Huntley, and J. F. Patience*

*Department of Animal Science, Iowa State University, Ames, 50011, USA

Abstract

It is important to understand if, and to what extent, the pig can utilize xylose as an energy source if xylanase releases free xylose in the small intestine. The experimental objectives were to determine the effects of industry-relevant dietary xylose concentrations and adaptation time on xylose retention efficiency and metabolism, diet digestibility and energy value, nitrogen balance, and hindgut fermentation. Forty-eight pigs were housed in metabolism crates and randomly assigned to one of four treatments with increasing D-xylose levels (n=12/treatment) in 2 replications of a 22-d experiment with 3 collection periods. The control diet was xylose-free (0%), to which either 2, 4, or

8% D-xylose was added. Adaptation effects were assessed during three fecal and urine collection periods: d 5-7, 12-14, and 19-21. On d 22, pigs from the 0 and 8% treatments were euthanized; cecal and colon digesta were collected. Dietary xylose did not affect the total tract digestibility of dry matter, gross energy, or crude protein; digesta short chain fatty acids concentrations and molar proportions, and cecal pH were not different

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(P>0.10). This experiment utilized a targeted metabolomics approach to characterize and quantify urine xylose and metabolite excretion. Xylose retention decreased from 60% to

47% to 41% when pigs were fed diets containing 2, 4, or 8% xylose, respectively. A comprehensive pathway for xylose metabolism was proposed and D-threitol was confirmed as the major urinary metabolite of xylose. In conclusion, pigs can metabolize xylose, but with considerably lower efficiency than glucose, and can adapt with time to utilize xylose more efficiently.

Introduction

Xylose is a primary component of arabinoxylan in the hemicellulose fraction of carbohydrates in swine diets. Xylose is a potential energy source within the fiber structure of arabinoxylan, which is unavailable to the pig as a monosaccharide but can contribute some energy through fermentation. Xylanase hydrolyzes the -linked xylose backbone of arabinoxylan and releases a mixture of oligosaccharides, disaccharides, and monomeric pentose sugars [1–3] such as xylose. Free xylose can be absorbed by the small intestine and thus could potentially contribute energy to the pig; yet little information is available on xylose metabolism in pigs and mammals in general [4]. It is important to understand the energy available from xylose because nutritionists formulate diets to precisely balance the concentrations of energy and nutrients. Therefore, to effectively utilize xylanase to improve pig performance and efficiency when fed high fiber diets, we must understand how xylanase supplementation impacts dietary energy availability through the release of free xylose.

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Most research on xylose metabolism in pigs has been conducted utilizing diets with xylose concentrations ranging from 5 - 40% [4]. However, typical growing pig diets will contain only 3 - 5% xylose as arabinoxylan, and not all of this would be released as free xylose in the small intestine. Additionally, most studies have allowed only 4 - 7 d of adaptation to treatment diets, so it is yet to be determined if pigs develop increased capacity for xylose metabolism as the length of exposure increases. Therefore, the objectives of this experiment were to determine the effects of industry-relevant dietary xylose concentrations and adaptation time on xylose retention efficiency and metabolism, diet digestibility and energy value, nitrogen (N) balance, and hindgut fermentation, and to identify urinary metabolites and propose a metabolic pathway of xylose using a targeted metabolomics approach.

Materials and methods

All experimental procedures adhered to guidelines for the ethical and humane use of animals for research and were approved by the Iowa State University Institutional

Animal Care and Use Committee (2-17-8452-S).

Animals, housing, and experimental design

The experiment was conducted at the Iowa State University Swine Nutrition

Farm (Ames, IA) using 48 crossbred gilts (Genetiporc 6.0 × Genetiporc F25, PIC, Inc.,

Hendersonville, TN) with an initial body weight (BW) of 28.3  1.5 kg in two replications using 24 pigs each.

Prior to the experiment, pigs were housed individually for 4 d in pens (1.8  1.9 m) with slatted concrete floors for adaptation to individual housing and to the basal diet

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and feeding schedule. At the start of the trial, pigs were randomly allotted to one of four dietary treatments and were individually housed in metabolism crates (0.7  1.5 m) for the 22-d experiment. The metabolism crates allowed for separate collection of feces and urine and were equipped with a slatted floor, feeder, and nipple waterer that was attached to a modified water container to allow accurate measurement of water intake. Pigs were weighed on d 0 and 22. On the last day of the experiment (d 22) a total of 24 pigs, 12 from each replication (six each from the 0% and 8% treatments), were euthanized for tissue collection via captive bolt stunning and exsanguination.

Diets and feeding

Four dietary treatments with increasing levels of D-xylose were evaluated (Table

1). The basal control diet was xylose-free (0%) and the following three treatments consisted of either 2%, 4%, or 8% D-xylose (DuPont Nutrition & Health, Copenhagen,

Denmark) added to the basal diet at the expense of corn starch. All diets were formulated to meet or exceed NRC [5] (2012) requirements for growing pigs (Table 2) and were manufactured in mash form. The same batch of feed was used for both replications.

During diet mixing, representative diet samples were obtained by homogenizing 10 subsamples; these were stored at -20C for future analysis. Titanium dioxide (TiO2) was included in the diet as an indigestible marker to determine nutrient digestibility. Four days prior to initiation of the experiment, all pigs were offered the 0% diet. On d 0, pigs were transitioned to their assigned dietary treatments at 4% of the average initial BW.

The daily feed allotment was split into two daily feedings at 0800 and 1600 h. Any orts remaining after 1 h were collected and weighed. Pigs in both replicates received the same amount of feed. Pigs had ad libitum access to water throughout the entire trial.

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Sample collection

The effect of adaptation time was assessed by utilizing three fecal and urine collection periods throughout the 22 d experiment. Collection periods 1 (d 5-7; C1), 2 (d

12-14; C2) and 3 (d 19-21; C3) allowed pigs to adapted to the treatment diets for 4, 11, and 18 d prior to each collection, respectively.

Each collection period lasted 72 h and total urine and fresh fecal grab samples were collected at 0830 and 1630 h each day. Fecal samples were compiled and homogenized within each collection period and immediately stored at -20°C. Urine was collected quantitatively into 4-liter bottles containing 6 M-HCl to ensure that the pH was maintained below 2.0 to minimize nitrogen (N) volatilization. Total urine output was weighed and stored at -20°C. After each collection period was completed, urine was thawed, homogenized, strained through glass wool, subsampled, and again stored -20°C for later analysis. Urine specific gravity was measured by weighing a subsample of 500 

1 ml.

Water intake was also measured during each collection period. A 12-liter plastic container was modified to attach to the nipple waterers already installed in the metabolism crates. The containers were placed on top of each crate to allow water to be gravity-fed into each waterer. The containers were emptied at the beginning of each collection period and the volume of water added throughout the collection period was recorded. At the end of the 72-h period the volume of water remaining was recorded to calculate water disappearance. Correspondingly, at the beginning of each collection period, trays were placed underneath the front of each crate to collect and measure the

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volume of wasted water. Water intake was then calculated by subtracting water waste from water disappearance during each collection period.

On d 22, 24 total pigs (12 pigs from the 0% treatment and 12 pigs from the 8% treatment) were euthanized 60  3 min after feeding. The abdomen of the carcass was opened and, to minimize digesta movement, the gastrointestinal tract was clamped at the stomach/duodenum sphincter, the ileum/cecum junction, the cecum/large intestine junction, and at the rectum. The intestines were then removed from just below the stomach to near the anus. Digesta pH was measured at the cecum and mid-colon, and digesta from both sections was collected, and immediately frozen. Digesta samples were stored on dry ice for transportation to the lab and stored at -20°C until analysis.

Analytical methods and calculations

All diet, orts, and fecal samples were dried at 60°C to a constant weight and were ground to a particle size of 1 mm. Urine subsamples were thawed, mixed, and filtered through Whatman 41 filter paper (GE Healthcare Life Sciences, Chicago, IL, USA) prior to analysis. Urine, diet, and fecal samples were analyzed in duplicate for N (method

990.03 [6]; TruMac; LECO Corp., St. Joseph, MI, USA). Standard calibration was performed using EDTA (9.56% N) which was determined to contain 9.55 ± 0.01% N.

Crude protein (CP) was calculated as N 6.25.

Diet and fecal samples were analyzed in duplicate for dry matter (DM; method

930.15), TiO2 and gross energy (GE). Titanium dioxide was determine according to the colorimetric method of Leone (1973) [7]. For urine energy determination, 3 ml of urine was added to 0.5 g of dried cellulose (SolkaFloc, International Fiber Corporation, North

Tonawanda, NY, USA), subsequently dried at 50C for 72 h, and urine plus cellulose

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dried samples were analyzed in duplicate for GE. Gross energy was determined using a bomb calorimeter (model 6200; Parr Instrument Co., Moline, IL). Benzoic acid (6,318 kcal/kg; Parr Instrument Co.) was used as the standard for calibration and was determined to contain 6,315 ± 6.9 kcal/kg. Urinary energy was calculated from the difference of energy determined in cellulose and the energy determined in the samples containing both urine and cellulose.

Apparent total tract digestibility (ATTD) of dry matter, GE, and CP were calculated according to the equation of Oresanya et al. [8], as 100 – {100 ×

[concentration (g) of TiO2 in diet × concentration (g) of DM, GE, or CP in feces]/[concentration (g) of TiO2 in feces × concentration (g) of DM, GE, or CP in diet]}.

Diets and a representative portion of the fecal and colon digesta samples (n = 24) were also analyzed for D-xylose concentration using a commercially available kit

(Megazyme, Wicklow, Ireland). To prepare diet samples, they were ground to 0.5 mm particle size and acid hydrolyzed according to manufacturer’s instructions. Fecal and colon digesta samples used for D-xylose analysis were lyophilized, ground to 0.5 mm particle size, and clarified according to manufacturer’s instructions. In all fecal and colon digesta samples, the xylose concentration was determined to be < 1% (average = 0.24%), which agrees with previous literature that free xylose is nearly completely absorbed in the small intestine [9]. Therefore, the ATTD of xylose was not measured. Diets were also analyzed in duplicate for acid hydrolyzed ether extract (aEE; method 2003.06) and in triplicate for neutral and acid detergent fiber components (NDF [10] and ADF [11], respectively). The analyzed nutrient composition of the diets is presented in Table 2.

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Quantitative metabolomics analysis was completed for all urine samples using tandem GC/MS. Filtered urine samples were vortexed, 10 l was transferred to a 2-ml glass vial, and 50 l of ribitol (1 mg/ml in HPLC grade water) was added to each sample as an internal standard. The samples were then concentrated to complete dryness in a

SpeedVac concentrator (Savant, Thermo Fisher Scientific Inc., Waltham, MA). The dried samples were treated with methoxamine (20 mg/ml methoxamine HCl in dry pyridine) followed by sonication at ambient temperature for 30 min and incubation at 60C for 60 min. The samples were then derivatized with the addition of N,O-bis(trimethylsilyl)- trifluoroacetamide with 1% trimethylchlorosilane and incubated at 60C for 30 min.

Analysis was performed on an Agilent 19091s-433 GC/MS system equipped with a HP-5ms chromatographic column (30 m  250 m  0.25 m; Santa Clara, CA). The injection temperature set to 280C and the injected volume was 1 l with a 1:10 split. The helium flow rate was 1 ml/min. The column temperature was initially set at 90C for 2 min and then increased to 320C at a rate of 15C/min where it was held for 3 minutes.

Mass spectral analysis was performed in the scan-total ion monitoring mode after electron ionization. The MS quadrupole temperature was set to 150C and the ion source temperature was set to 230C. The mass data were collected in the range from m/z 40 to m/z 800. The acceleration voltage was turned on after a solvent delay of 7 min.

The AMDIS software (National Institute of Standards and Technology (NIST),

Gaithersburg, MD) was used for peak quantification. A hydrocarbon ladder was used for retention index calculation and standards of D-threitol (Sigma-Aldrich, Saint Louis, MO),

D-xylose, D-erythritol, xylitol, D-xylulose, and D-xylonic acid (Supelco, Bellefonte, PA) were used for compound identification. Additional compounds were identified using

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NIST14 Mass Spectral Library (version 2.2) based on mass spectral matching. The integrated peak areas of multiple derivative peaks belonging to the same compound were summed and considered as a single compound. Concentrations of identified compounds were calculated using the ratio of the peak area of each compound to the internal standard; and D-xylose, D-threitol, D-erythritol, xylitol, D-xylulose, and D-xylonic acid were quantified based on a standard curve regression.

Urine GE (kcal/l) contributed from xylose, threitol, xylitol, xylonic acid, and xylulose were calculated as the concentration of the compound (g/l)  GE of the compound (kcal/g). Urine GE contributed by N-containing compounds was estimated by applying an assumption that all urinary N was in the form of urea and was calculated as the urine N concentration (g/l)  2.52 (kcal/g, the GE of urea). Urine GE not allocated to xylose, metabolites, or N-containing compounds was calculated as urine GE (kcal/l) –

[urine GE from xylose + threitol + xylitol + xylonic acid + xylulose + N-containing compounds (kcal/l)].

Digesta samples from the cecum and colon were analyzed for short chain fatty acid (SCFA) concentrations. One gram of sample was diluted with 2.5 ml ddH2O and mixed with 0.2 ml of 25% metaphosphoric acid and 0.1 ml of isocaproic acid (48.3 mM;

Sigma-Aldrich, Saint Louis, MO) as an internal standard. A standard curve was generated using five concentrations of acetate, propionate, butyrate, valerate, isobutyrate, and isovalerate (Sigma-Aldrich, Saint Louis, MO). The internal standard (0.1 ml) was added to each set of standards. Samples and standards were centrifuged, and the supernatant was transferred to a glass vial for analysis. Gas chromatography (3800 Varian GC,

Agilent Technologies, Santa Clara, CA) was performed using a 15 m  0.25 mm  0.25

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μm fused silica capillary column (Supelco, Bellefonte, PA) with flame ionization detection. Helium was used as the carrier gas at the flow rate of 1 ml/min. The injector temperature was 250°C and the oven temperature held at 143°C until increasing to 200°C for 6 min. Concentrations of acetate, propionate, butyrate, valerate, isobutyrate, and isovalerate were calculated using the ratio of the peak area of each compound to the internal standard and standard curve regression. Molar proportions of SCFA were calculated as the individual SCFA concentration (mM) / total SCFA concentration (mM).

Statistical analyses

Data were analyzed as a three by four factorial design with pig as the experimental unit. The UNIVARIATE procedure of the Statistical Analysis Systems package version 9.4 (SAS Institute, Cary, NC, USA) was used to verify normality of residuals and homogeneity of variances and statistical outliers (> 3 standard deviations away from the mean) were removed. Water intake, urine output, and digesta SCFA molar proportions data were log transformed, and SCFA concentration data were reciprocal transformed to achieve a normal distribution. The statistical models for all SCFA data were fit with separate variances for each of the two treatments.

The fixed effects of dietary treatment (Trt), collection period (Col), and their interaction (Trt  Col), and the random effect of replication were analyzed using the

MIXED procedure of SAS. Data were analyzed as repeated measures with pig as the subject and an unstructured covariance structure was applied. For data collected only at one time period (i.e. growth performance and cecal and colon pH and SCFA), the fixed effect of dietary treatment was analyzed with replication as a random effect.

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Differences due to dietary treatment, collection period, and their interaction were determined using ANOVA and means were separated using the least square means statement and the PDIFF option. Linear and quadratic polynomial contrasts were used to determine the response to dietary xylose concentration. Differences were considered significant if P was ≤ 0.05 and a trend if P was > 0.05 and ≤ 0.10.

Urine metabolomics data were analyzed as grams of compound excreted per day to account for differences in urine volume. Multivariate analysis of log-transformed and range-scaled data was performed using MetaboAnalyst 4.0 [12] including repeated measures two-way ANOVA for the effects of dietary treatment and collection period, heat map generation, and the impact of dietary xylose concentration on metabolic pathways and metabolite set enrichment.

Results

Growth performance and water balance

All pigs remained healthy throughout the entire trial. Initial BW and d 22 BW did not differ (Trt P  0.6659; Table 3) and pigs on all dietary treatments had similar average daily gain (ADG) and feed efficiency (Trt P  0.1653; Table 3). Water intake increased in a linear fashion as dietary xylose concentration increased; in turn, this resulted in a linear increase in urine output (Table 4; Trt P < 0.01). However, urine specific gravity did not differ among treatments (Table 4). A tendency for differences among dietary treatments (P = 0.0656) and a significant collection period effect (P = 0.0087) were detected for urine specific gravity, but these were likely due to the very small standard error of the mean (SEM; 0.01). In any event, this magnitude of differences would not be

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biologically relevant. Collection period significantly impacted water balance as well as water waste, with pigs wasting less water during C2 and C3 compared to C1 (P = 0.0001;

Table 4)

Diet digestibility and energy value and nitrogen balance

The ATTD of DM, GE, and CP did not differ among treatments (Trt P  0.4074;

Table 5), but the ATTD was significantly impacted by collection period (Col P 

0.0014). There were significant interactions between treatment and collection period for

DM and GE ATTD (Trt  Col P  0.0003) and a trend for interaction for CP ATTD (Trt

 Col P = 0.0858; Table 5). Small differences in GE ATTD resulted in a tendency for an interaction of treatment and collection period on diet digestible energy (DE, Mcal/kg;

Table 5). Dietary treatment affected diet DE with pigs on the 2% xylose treatment having lower DE value compared to all other treatments (3.56 Mcal/kg vs. an average of 3.61

Mcal/kg; Trt P = 0.0004). However, the SEM for all digestibility variables was relatively low ( 0.50) which likely resulted in detection of the interaction.

Urine N (%) decreased linearly with increasing dietary xylose concentration (Trt

P = 0.009, linear P = 0.009; Table 5) but increased across collection periods as pigs had longer adaptation time to the diets (Col P < 0.0001). There tended to be an interaction between treatment and collection period for urine N with concentrations increasing across collection periods for all treatments, but to a lower magnitude in the 8% xylose treatment

(Table 5). Nitrogen balance (retained N and the percent of N excreted in the urine or feces) was not affected by dietary treatment (Trt P  0.5738; Table 5). As pigs had longer adaptation time, the proportion of N excreted in the feces increased (Col P < 0.0001) while the proportion excreted in the urine decreased (Col P < 0.0001).

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Urine energy concentration (kcal/l) linearly increased with increasing dietary xylose concentration (Trt P < 0.0001, linear P < 0.0001) and increased across collection periods as pigs had longer adaptation time to the treatment diets (Col P < 0.0001). This resulted in a significant treatment by collection period interaction for diet metabolizable energy (ME, Mcal/kg) value (Table 4.5).

Digesta short chain fatty acids

Short chain fatty acid concentrations (mM) and molar proportions in the cecum and colon did not differ between the 0 and 8% treatments (Trt P  0.2534; Table 6). The pH of the cecal digesta did not differ (Trt P = 0.3845) but colon digesta in pigs fed 8% xylose had a lower pH (Trt P = 0.0249; Table 6).

Urine metabolomics

Figure 1 displays the distribution patterns of compounds identified in urine from pigs fed increasing dietary xylose concentrations and among three collection periods representing increasing adaptation time to treatment diets. The Euclidean distance measure and the unweighted paired group average linkage technique for hierarchical clustering were applied. In total, 22 compounds were identified in the urine samples and

20 were determined to be differentially excreted due to dietary treatment, collection period, or their interaction (adjusted P < 0.05; Fig. 1). All compounds were impacted by collection period, except glutaric acid and malic acid which were significantly affected only by treatment. Erythrono-1,4-lactone was only affected by the main effect of collection period. In general, increasing dietary xylose concentration increased the excretion of metabolites in the urine with the exceptions of malic acid, glutaric acid,

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pseudouridine, citric acid, and p-cresol glucuronide for which excretion decreased (Trt adjusted P < 0.05).

Xylose metabolic pathway

Metabolomic analysis was used to identify metabolic pathways impacted by xylose consumption and determine metabolite set enrichment. According to the pathway enrichment analysis, the pentose and glucuronate interconversions pathway was the most significantly impacted (Trt P < 0.001) and contained the largest proportion of identified compounds (4/22) that were enriched in the pathway. Based on this analysis, a review of the literature on xylose metabolism in pigs [13–24] and other mammals [25–35] where data specifically in pigs were not available, and validation using KEGG metabolic pathways [36] and BRENDA comprehensive enzyme database [37], a pathway for the metabolism of xylose is proposed (Fig. 2). Clustering results (Fig. 1) and interpretation of the xylose metabolic pathway (Fig. 2) were used to determine that the most plausible direct metabolites of dietary xylose were considered to be D-threitol, xylitol, D-xylulose, and D-xylonic acid, in addition to xylose excreted in the urine unchanged. The excretion of all other identified compounds in the urine significantly affected by dietary xylose concentration were considered to have been indirectly affected as opposed to directly converted from dietary xylose.

Xylose metabolite excretion

The excretion of xylose and its metabolites are presented in Table 7. Xylose was the most abundant compound identified in urine from pigs on the 2, 4, and 8% xylose treatments. Xylose excretion increased as xylose consumption increased (Trt P < 0.0001, linear P < 0.0001, quadratic P < 0.0001). In the 0 and 2% xylose treatments, xylose

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excretion was similar across collection periods, but in the 4 and 8% treatments, excretion was decreased in C2 and C3 compared to C1 (Col P < 0.0001). In the 4% treatment, xylose excretion was not different between C2 and C3 but in the 8% treatment, xylose excretion was greater in C3 compared to C2, but still less than C1 (Trt  Col P < 0.0001;

Table 7). In pigs fed xylose-containing diets, the percent of consumed xylose excreted in the urine increased as dietary xylose concentration increased (Trt P < 0.0001, linear P <

0.0001; Table 8) but this proportion decreased after the first collection period (Col P <

0.0001). In the 8% treatment, the proportion was greater in C3 compared to C2 (Trt  Col

P = 0.0026, Table 8).

Threitol was the major metabolite of xylose. Urinary excretion of threitol increased as xylose consumption increased (Trt P < 0.0001, linear P < 0.0001, quadratic

P < 0.0001) and, similar to xylose, a treatment by collection period interaction was observed (Trt  Col P = 0.0008; Table 7). In pigs fed the xylose-containing diets, the percent of consumed xylose excreted in the urine as threitol increased as dietary xylose concentration increased (Trt P < 0.0001, linear P < 0.0001, Table 8). This proportion differed across all collection periods with C1 > C3 > C2 (Col P < 0.0001).

Xylitol, xylonic acid, and xylulose were minor metabolites of dietary xylose.

Urinary excretion of these metabolites also increased as xylose consumption increased

(Trt P < 0.0001, linear P < 0.0001) and significant treatment by collection period interactions were detected (Trt  Col P  0.0021; Table 7). The proportion of consumed xylose excreted as either xylitol, xylonic acid, or xylulose was also impacted by the interaction (Trt  Col P = 0.0133; Table 8), but in no instance did it exceed 1.1% of xylose intake.

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Allocation of urine GE

Because urine GE was found to significantly increase as dietary xylose concentration increased, it was of interest to determine where the excess urinary energy was coming from. The allocation of urine GE to xylose and its derivatives is presented in

Fig. 3; values are averaged across collection period. Xylose contributed a large portion of the excess urine energy; the remainder was explained by its metabolite threitol, and to a lesser extent xylitol, xylonic acid, and xylulose. In the 4 and 8% treatments the amount of urine GE coming from N-containing compounds was lower than the control treatment

(Trt P  0.0088; Fig. 3). Once the differences in energy coming from xylose, its metabolites, and N-containing compounds were accounted for, the remaining unassociated GE was similar among the 0, 2, and 4% treatments, but was lower in the 8% treatment (Trt P = 0.0420; Fig. 3). The underlying reason for this difference has yet to be determined.

Discussion

Growing pigs are commonly being fed diets with increasing dietary fiber levels.

To help mitigate the negative effects of fiber on growth performance and feed efficiency, exogenous carbohydrases that break down the fiber structure are often supplemented

[38,39]. Of the fiber that is found in cereal grains and co-products, a large proportion is hemicellulose and of this, a large fraction is arabinoxylan [40–42], which in turn is primarily composed of the pentose sugars xylose and arabinose. Xylanase releases a portion of the xylose from the xylan backbone [1–3]. This free xylose can then be absorbed in the small intestine [9,43–46] and potentially contribute energy to the pig.

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However, previous research indicates that free xylose is poorly utilized [4], so it is unknown whether xylose would contribute more energy when absorbed as a monosaccharide or through fermentation of arabinoxylan in the hindgut.

Energy is the most expensive component of swine diets [47], so nutritionists formulate diets to precisely balance the concentrations of available energy and nutrients.

Therefore, to effectively utilize xylanase to improve the efficiency of pig production, the amount of energy contributed by free xylose must be quantified. While the efficiency and extent of the release of free xylose by xylanase has yet to be determined, the concentration of xylose available for absorption is unlikely to be greater than 4% of diet

DM [4]. To our knowledge, only one previous study in pigs has been reported using dietary D-xylose concentrations below 5% [44], and the pig’s ability to adapt and improve its utilization of xylose has yet to be evaluated. To accomplish the experimental objectives and to begin to address the question “what is the energy value of xylose for pigs?”, this experiment utilized four diets starting with a xylose-free control diet (0%) to which either 2, 4, or 8% xylose was added. Samples were collected in three separate periods over 22 d to evaluate the effects of adaptation time.

Water balance

Water balance was clearly impacted by xylose consumption in this study.

Increasing xylose consumption increased water intake and urine output which agrees with previously reported xylose effects in pigs [9]. We hypothesize that more water was excreted in the urine to balance osmolarity as increasing amounts of xylose and other metabolites were excreted. This hypothesis is supported by the constancy of the specific gravity of urine across all treatments, even though the quantity of solutes excreted per

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day increased. As adaptation time increased, pigs excreted less xylose in the urine, and water intake and urine output were lower, accordingly. It would be interesting in future studies evaluating xylanase efficacy to measure water consumption and relate this to xylose release.

Digestibility, fermentation, and diet energy value

Contrary to the results of previous studies [9,43], dietary xylose concentration did not affect total tract digestibility of DM, GE, or CP in biologically relevant ways. Among all treatments and throughout the entire trial, the ATTD of DM ranged from 87.6 – 89.1% and diet DE was found to be slightly lower in the 2% treatment compared to the other three treatments. Based on previous research [9,43,44] and analysis of a subset of the colon digesta and fecal samples from pigs on this experiment, dietary xylose was assumed to be completely absorbed by the terminal ileum. Because possible microbial fermentation of xylose in the small intestine [9,44] was not evaluated, one potential drawback of this assumption is an overestimation of retained xylose. However, there is empirical evidence that xylose is readily absorbed in the duodenum and proximal jejunum in rats [48,49]. This suggests that the concentration of any unabsorbed xylose would likely be very low in the terminal ileum where microbial populations are more abundant.

The SCFA results from this experiment are consistent with the hypothesis that xylose was almost completely absorbed in the small intestine. Concentrations and molar proportions of SCFA did not differ between the 0 and 8% treatments, indicating that the nutrient composition of substrates fermented in the cecum and colon were similar [50].

In accordance, the pH of cecal digesta did not differ. Because no differences were

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detected in SCFA concentrations in either the cecum or colon, an explanation for a lower pH in the colon of pigs fed 8% xylose is not directly apparent.

Urine energy concentration linearly increased with increasing dietary xylose concentration, and diet ME decreased with xylose inclusion. Across all collection periods, the 8% treatment consistently provided the lowest diet ME. Although xylose did decrease the ME value of the diet, the relative differences of the 2 and 4% xylose diets compared to the control diet are quite low. For the 2% treatment across C1 – 3, the ME values were 98%, 97%, and 96% the value of the control diet and for the 4% treatment, the values were 98%, 98%, and 97%, respectively. The 8% xylose diets had the lowest relative ME value at 94%, 95%, and 93% of the control diet during C1 – 3, respectively.

Unlike previous studies using dietary xylose concentrations of 10% [43] or 18.7 and

37.4% [51], dietary xylose inclusion did not impact ADG or feed efficiency in pigs on this experiment. This result may be partially explained by the relatively small differences in diet ME value among treatments.

Xylose metabolism

Dietary xylose, adaptation time to the diets (i.e. the effect of collection period), and their interaction significantly impacted the profile of compounds excreted in the urine of pigs. Many of the identified compounds are related to general energy metabolism (ex. malate and citrate), amino acid metabolism (ex. glutaric acid), or are derivatives of microbial metabolism (ex. p-cresol glucuronide and hippuric acid) [52]. The excretion of these compounds was impacted by dietary xylose concentration and indicates that metabolic pathways beyond those directly related to xylose were affected.

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Interestingly, sucrose excretion was found to increase with increasing dietary xylose concentrations. Although total urinary sucrose excretion was low (0.03 – 0.12 g/d), it is uncommon for sucrose to be found in the urine of healthy animals. If sucrose is not hydrolyzed by sucrase at the intestinal brush border, it can be absorbed intact and is usually excreted as such in the urine [53,54]. The fact that urinary sucrose excretion increased due to xylose consumption, even though all diets contained the same amount of sucrose, may support the conclusions of previous studies that xylose inhibits intestinal sucrase activity [55] and reduces the post-prandial glycemic effect of sucrose-containing meals [56,57].

By far, xylose was the most concentrated compound identified in the urine of pigs fed xylose-containing diets. A small amount of xylose was excreted in the urine of the control pigs (0.61 g/d) which is comparable to basal xylose urinary excretion previously reported [44]. As expected, as dietary xylose concentration increased, urinary xylose excretion increased from 6.3 to 40.1 g/d. Adaptation was evident in the 4 and 8% treatments because xylose excretion was significantly lower in C2 and C3 compared to

C1. However, in the 8% treatment, xylose excretion was greater in C3 compared to C2.

To be able to compare xylose utilization efficiency on an equivalent basis across all treatments, the proportion of consumed xylose that was excreted in the urine was calculated (Table 8). As xylose consumption increased, the percent excreted in the urine as D-xylose increased from 27.6 to 39.2 to 43.9% in the 2, 4, and 8% xylose treatments, respectively. This was the first indication that the efficiency of xylose utilization decreases proportionally as dietary xylose concentration increases. These value are

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comparable to previous studies in pigs which reported that 35 – 53% of absorbed xylose is excreted in the urine [9,43,44].

Previous research has also shown that xylose can be metabolized to other compounds. Radioisotope studies in mammals demonstrated that xylose can be oxidized to xylonic acid [25], CO2 [25,58], and be converted to threitol [26,43]. Authors of previous studies have proposed metabolic pathways for some conversions; however, they have generally only been speculated upon for individual reactions [28] or short conversion pathways [25,26]. We have proposed a more comprehensive metabolic pathway to help explain the conversion of xylose to urinary metabolites identified in this study. Metabolic connections with the pentose phosphate pathway (PPP) were shown due to the common assumption that xylose, being a 5-carbon sugar, would be metabolized within the PPP. The connections to the PPP are also likely the means through which xylose can eventually be fully oxidized to CO2 and contribute energy to the pig. The proposed pathway was interpreted in conjunction with the cluster analysis of metabolomics data, that indicate which compounds were most similarly influenced by dietary treatment. As discussed below, research in pigs confirms the presence of the enzymes, mostly in the liver and kidneys, which can convert D-xylose into either D- xylonic acid or xylitol.

D-xylose is oxidized to D-xylonic acid by NADP+-linked D-xylose dehydrogenase (EC 1.1.1.179) with D-xylonolactone formed as an intermediate. This enzyme and metabolic conversion has been confirmed in pig liver [13,14,17] with lower activity in the kidney and lens, and even lower activity in the spleen, brain, heart and muscle [18]. The same pathway was demonstrated in guinea pigs using intraperitoneal

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injections of D-xylose-1-14C [25]. The same study then demonstrated that D-xylonic acid-

1-14C was metabolized much faster than D-xylose and a greater proportion was converted

14 to CO2. This may indicate that D-xylose dehydrogenase could be a rate-limiting step for xylose metabolism. In our study, D-xylonic acid was excreted in the urine at low levels

(< 0.6 g/d) but its excretion increased with increasing xylose consumption.

D-xylonic acid can then be converted to D-erythrulose by L-gulonate 3- dehydrogenase (EC 1.1.1.45); this enzyme activity has been confirmed in pig kidney[19].

D-erythrulose can then be reduced by D-erythrulose reductase (EC 1.1.1.162) to D- threitol, the most abundant xylose metabolite identified in the urine in our study. This enzyme (also known as D-threitol:NADP+ oxidoreductase) has not been reported specifically in the pig; however, it has been confirmed in the mouse kidney and liver [32] and cattle liver [29,30], with the highest enzyme activity reported in the kidney. If not converted to D-threitol, D-erythrulose can also be phosphorylated to D-erythrulose-4P, which links to the PPP.

In this study, D-threitol excretion increased with increasing xylose consumption - up to 16 g/d in the 8% treatment during C1. If D-threitol is assumed to be a metabolite of dietary xylose, urinary D-threitol represented 11, 13, or 15% of consumed xylose when the 2, 4, or 8% diet was fed, respectively. The identification of D-threitol as the major urine xylose metabolite corroborates previous studies in pigs [43] and humans [26] which both reported that 10-11% of ingested xylose was excreted as threitol. Pitkänen [26] specified that in the first 5 h after ingestion only 4% of the oral xylose dose was excreted as threitol; when the collection was extended to 24 h, the proportion increased to 10.6%.

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In the literature, the ME value of diets is consistently reduced by xylose inclusion

[4,9], but it is likely that in addition to excreted xylose, threitol also significantly accounts for the reduction in ME. This was apparent in our study and likely was an unidentified causative factor in the studies by Verstegen et al. [43] and Schutte et al. [9].

Furthermore, threitol likely explains the unidentified compound in guinea pig urine that contained 18% of an intraperitoneal injection of D-xylose-1-14C [25].

D-xylose can also be oxidized to xylitol via aldose/aldehyde reductase (EC

1.1.1.21). This conversion has been confirmed in the cortex of pig kidney [20,59], but the reaction seems to occur at a slow rate [20]. The enzyme has also been identified in the lens and skeletal muscle of pigs [21]. Xylitol was the second most abundantly excreted metabolite of xylose, but at most, only 0.41 g/d were excreted. In all xylose-containing treatments, xylitol excretion was the greatest during C3, potentially indicating a slow metabolic upregulation of aldose/aldehyde reductase as pigs adapted to the diets. Xylitol was also identified as a metabolite in humans infused with D-xylose-1-14C, but represented < 0.1% of the recovered 14C [58].

Xylitol can be converted to either the D- or L-isomer of xylulose. In this study, D- xylulose excretion was found to increase with increasing xylose consumption. Xylitol dehydrogenase (EC 1.1.1.14) oxidizes xylitol to D-xylulose and this enzyme is distributed in many mammalian tissues of which, the liver is the main organ [15]. Finally, in the liver and kidneys, D-xylulose can be phosphorylated by xylulokinase (EC 2.7.1.17) to D-xylulose-5P[27], an intermediate in the PPP. Xylitol, D-xylulose, and D-xylonic acid were determined to be minor metabolites of dietary xylose, and together represented

0.58 – 1.03% of consumed xylose.

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Taking into account xylose that was either excreted in the urine directly or excreted as threitol or a minor metabolite, the proportion of consumed xylose that was retained in the body ranged from 27 – 63%. Increased dietary xylose concentration decreased the retention efficiency across all collection periods. However, a larger proportion of xylose was retained during C2 and C3, i.e. after pigs had been adapted to the treatment diets for at least 11 d. This indicates that pigs are able to adapt metabolically to utilize xylose more efficiently. Further research is necessary to determine how this occurs, which mechanisms may be up or down regulated, and the final metabolic fate of retained xylose in pigs.

Dietary xylose concentration clearly impacts its retention efficiency. This is a consistent finding across studies in pigs and other species, as reviewed earlier this year by

Huntley and Patience [4]. Considering these data in the context of commercial pig production, it is unlikely that exogenous xylanase supplementation would release free D- xylose in the small intestine at concentrations > 4% of the total diet DM. Based on the results of this experiment, only about 50 – 63% of the xylose released would be retained and could potentially contribute energy to the pig through oxidation. Experiments must still be conducted to directly compare the net energetic value of xylose through metabolic oxidation verses fermentation. To speculate, based on the xylose retention efficiency measured in pigs fed 4% xylose in this study (51%), and using the metabolic energy efficiencies of xylose reported by Yule and Fuller [44] and reported energetic efficiencies of fiber fermentation in the pig [60–62], it is estimated that xylose would provide approximately 15% less energy if absorbed in the small intestine vs. through fermentation.

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Further research is needed to determine how much free xylose is released from arabinoxylan in the small intestine of pigs due to xylanase supplementation. Furthermore, it would be valuable to examine the liver and kidney for enzymes that may be differentially expressed and regulated in pigs consuming xylose. This will help determine how adaptation to xylose-containing diets occurs, and how other metabolic processes may be affected. Understanding potential effects on other metabolic processes may also help to explain the decreased proportion of urine GE not associated with xylose, its metabolites, or N-containing compounds in pigs fed 8% xylose diets.

Conclusions

In conclusion, pigs can metabolize xylose, but with considerably lower efficiency than glucose. Furthermore, pigs can adapt over time to utilize xylose more efficiently.

Xylose retention decreased from 60% to 47% to 41% when pigs were fed diets containing

2, 4, or 8% xylose, respectively. Thus, increasing dietary xylose concentration decreases diet ME. A comprehensive pathway for xylose metabolism in the pig is proposed and D- threitol was confirmed as the major urinary metabolite of xylose. Xylitol, D-xylonic acid, and D-xylulose were identified as minor metabolites.

This experiment provides novel data regarding xylose metabolism when fed at relevant concentrations for typical swine diets. To fully understand these data in the context of practical swine nutrition, further research is needed to determine how much free xylose is released from arabinoxylan with xylanase supplementation. Because of the importance and cost of energy in swine diets, assigning an energy release value to

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xylanase is of great interest. These data indicate that xylose’s energy contribution to the pig will depend on the amount of xylose released and the length of adaptation to the diet.

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Table 1. Diet ingredient composition (as-fed) Dietary treatment, % xylose inclusion

Item, % 0 2 4 8 Corn starch 57.00 55.00 53.00 49.00 Sucrose 8.60 8.60 8.60 8.60 Cellulose 9.00 9.00 9.00 9.00 Casein 5.50 5.50 5.50 5.50 Bovine plasma 5.00 5.00 5.00 5.00 Fish meal 4.50 4.50 4.50 4.50 Skim milk powder 3.00 3.00 3.00 3.00 Whey powder 3.00 3.00 3.00 3.00 Soybean oil 1.00 1.00 1.00 1.00 D-Xylose1 0.00 2.00 4.00 8.00 Limestone 0.90 0.90 0.90 0.90 Monocalcium phosphate 0.35 0.35 0.35 0.35 Choline chloride 0.05 0.05 0.05 0.05 Potassium carbonate 0.50 0.50 0.50 0.50 Magnesium oxide 0.07 0.07 0.07 0.07 L-Lysine HCl 0.05 0.05 0.05 0.05 DL-Methionine 0.10 0.10 0.10 0.10 L-Threonine 0.03 0.03 0.03 0.03 Vitamin premix2 0.25 0.25 0.25 0.25 Trace mineral premix3 0.20 0.20 0.20 0.20 Salt 0.50 0.50 0.50 0.50 Titanium dioxide 0.40 0.40 0.40 0.40 1DuPont Nutrition & Health, Copenhagen, Denmark 2Provided 6,614 IU vitamin A, 827 IU vitamin D, 26 IU vitamin E, 2.6 mg vitamin K, 29.8 mg niacin, 16.5 mg pantothenic acid, 5.0 mg riboflavin, and 0.023 mg vitamin B12 per kg of diet. 3Provided 165 mg Zn (zinc sulfate), 165 mg Fe (iron sulfate), 39 mg Mn (manganese sulfate), 17 mg Cu (copper sulfate), 0.3 mg I (calcium iodate), and 0.3 mg Se (sodium selenite) per kg of diet.

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Table 2. Formulated and analyzed diet energy and nutrient composition (as-fed) Dietary treatment, % xylose inclusion

Item 0 2 4 8 Formulated composition SID amino acid, % Lys 1.02 1.02 1.02 1.02 Met 0.36 0.36 0.36 0.36 Total sulfur AA 0.56 0.56 0.56 0.56 Thr 0.61 0.61 0.61 0.61 Trp 0.18 0.18 0.18 0.18 Ca, % 0.66 0.66 0.66 0.66 STTD P, % 0.33 0.33 0.33 0.33 Analyzed composition DM, % 94.02 94.90 95.26 95.58 GE, Mcal/kg 3.92 3.91 3.95 3.96 CP, % 14.05 14.03 14.07 14.03 aEE, % 1.62 1.57 1.76 1.52 NDF, % 10.31 10.40 10.49 10.56 ADF, % 7.62 7.71 7.62 8.00 Xylose, % 0.00 1.99 4.02 8.02 Pigs (n = 12/treatment) were housed individually in metabolism crates and fed diets containing either 0, 2, 4, or 8% D-xylose at 4% of BW and fecal samples were collected during 3 different periods representing increasing adaptation time to treatment diets. SID, standardized ileal digestible; AA, amino acids; STTD, standardized total tract digestible; GE, gross energy; CP, crude protein; aEE, acid hydrolyzed diethyl ether extract; NDF, neutral detergent fiber; ADF, acid detergent fiber.

Table 3. Effect of dietary xylose concentration on pig growth performance Dietary treatment, % xylose inclusion

Item 0 2 4 8 SEM P-value BW, kg d 0 28.18 28.27 28.38 28.20 0.46 0.9895 d 22 40.90 40.23 40.75 40.28 0.46 0.6659 ADG, g/d 606 570 589 576 20 0.1653 Gain: feed 0.53 0.50 0.52 0.51 0.02 0.1660 Pigs (n = 12/treatment) were housed individually in metabolism crates and fed diets containing either 0, 2, 4, or 8% D-xylose at 4% of BW for 22 d. BW, body weight; ADG, average daily gain

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Table 4. Effect of dietary xylose concentration and collection period on water balance

Dietary treatment, % xylose inclusion Collection period T×C1

Item 0 2 4 8 SEM P-value d 5-7 d 12-14 d 19-21 SEM P-value P-value Water waste, ml/d 853 796 1024 1265 321 0.3633 1404x 712y 837y 294 0.0001 0.4096 Water intake‡, ml/d 2447b 2532b 2762ab 3017a 222 0.0424 3025x 2551y 2493y 176 <0.0001 0.4352 Urine output‡, ml/d 984c 1159bc 1286ab 1566a 129 0.0032 1465x 1196y 1085z 74 <0.0001 0.7898 Urine specific gravity 1.034 1.031 1.025 1.029 0.009 0.0656 1.029xy 1.028y 1.032x 0.009 0.0087 0.2528 Pigs (n = 12/treatment) were housed individually in metabolism crates and fed diets containing either 0, 2, 4, or 8% D-xylose at 4% of BW. Water intake and urine output were measured during 3 different collection periods representing increasing adaptation time to treatment diets. a-cTreatment means without a common superscript differ (P  0.05) x-zCollection period means without a common superscript differ (P  0.05)

1 1 132 Dietary treatment × collection period interaction (T×C) 60 ‡

Treatment linear contrast is significant at P < 0.01

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Table 5. Effect of dietary xylose concentration and collection period on diet digestibility and energy value and on nitrogen balance Collection 5 - 7 12 - 14 19 - 21 period, d Dietary Pooled T×C1 treatment, % 0 2 4 8 0 2 4 8 0 2 4 8 SEM P-value xylose inclusion ATTD, % DM† 87.82de 88.49abcd 88.39abcde 88.12bcde 88.14cde 89.10a 88.89ab 88.58abcd 88.70abc 87.56e 88.80abc 88.06cde 0.30 0.0003 GE† 88.98ef 89.45bce 89.43abcef 89.33bcef 89.39ce 90.18a 90.05ab 89.69abce 90.00abd 88.62f 90.01abc 89.19cdef 0.29 <0.0001 CP† 90.86 91.16 91.29 90.74 91.96 92.18 91.78 91.64 92.62 91.95 92.57 91.59 0.50 0.0858 Urine GE*‡†, 171 212 220 289 213 241 273 312 224 249 300 328 17 0.5803 kcal/l Diet energy value, Mcal/kg, as-fed DE*#† 3.56 3.54 3.61 3.59 3.61 3.58 3.63 3.63 3.63 3.57 3.66 3.62 0.02 0.0857 ME*‡† 3.30bc 3.24de 3.23e 3.11g 3.33b 328bc 3.27cde 3.17f 3.37a 3.24e 3.28cd 3.14fg 0.02 0.0116

Urine N*‡†, % 0.28 0.31 0.20 0.21 0.43 0.39 0.33 0.29 0.50 0.44 0.41 0.34 0.03 0.0962 1

Nitrogen balance Retained, % 76.70 75.08 78.72 75.98 76.32 75.97 76.03 75.90 76.25 75.85 75.69 73.19 1.62 0.1497 of intake N in feces†, % 59.98 63.08 58.52 61.03 65.30 66.25 64.86 66.02 67.97 66.12 69.22 69.18 2.03 0.2949 of excreted N N in urine†, % 40.02 36.92 41.48 38.97 34.70 33.75 35.14 33.98 32.03 33.88 30.78 30.82 2.03 0.2949 of excreted N Pigs (n = 12/treatment) were housed individually in metabolism crates and fed diets containing either 0, 2, 4, or 8% D-xylose at 4% of BW. Feces and urine were collected during 3 different periods representing increasing adaptation time to treatment diets. ATTD, apparent total tract digestibility; GE, gross energy; CP, crude protein; ME, metabolizable energy; N, nitrogen a-fWithin a row, means without a common superscript differ (P  0.05) 1Dietary treatment × collection period interaction (T×C) *Treatment is significant at P < 0.01 #Treatment linear contrast is significant at P < 0.05 ‡Treatment linear contrast is significant at P < 0.01 †Collection period is significant at P < 0.01

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Table 6. Effect of dietary xylose inclusion on cecum and colon digesta pH and SCFA concentration and molar proportions Dietary treatment, % xylose inclusion

0 SEM 8 SEM P-value Digesta pH

Cecum 6.64 0.11 6.54 0.11 0.3845 Colon 6.52 0.10 6.17 0.10 0.0249 Cecal digesta SCFA concentration, mmol/g Acetate 22.55 1.76 22.55 1.11 0.8394 Propionate 8.82 0.70 8.90 0.62 0.9281 Butyrate 2.13 0.16 2.46 0.29 0.4230 Valerate 0.87 0.06 0.91 0.09 0.7377 Isobutyrate 0.68 0.04 0.68 0.04 0.8756 Isovalerate 1.14 0.09 1.10 0.05 0.6613 Total 36.20 2.42 37.03 1.62 0.5341 Cecal digesta SCFA molar proportion, % Acetate 61.97 1.78 62.02 1.40 0.9819 Propionate 24.41 1.24 23.97 1.18 0.7782 Butyrate 6.00 0.65 6.59 0.78 0.4075 Valerate 2.43 0.12 2.51 0.24 0.9100 Isobutyrate 1.96 0.16 1.89 0.15 0.7435 Isovalerate 3.23 0.27 3.02 0.14 0.4632 Colon digesta SCFA concentration, mmol/g Acetate 22.20 2.09 22.12 2.19 0.9731 Propionate 8.39 0.73 8.91 0.79 0.7775 Butyrate 3.65 0.41 3.60 0.34 0.9181 Valerate 1.00 0.11 1.01 0.11 0.8466 Isobutyrate 0.72 0.06 0.64 0.06 0.3509 Isovalerate 1.38 0.09 1.22 0.09 0.2534 Total 37.34 1.97 37.51 2.11 0.9474 Colon digesta SCFA molar proportion, %

Acetate 59.06 2.73 58.55 2.97 0.8741 Propionate 22.57 1.68 23.90 2.07 0.6321 Butyrate 9.85 1.34 9.62 1.18 0.9392 Valerate 2.70 0.29 2.73 0.31 0.9706 Isobutyrate 2.04 0.23 1.80 0.24 0.4379 Isovalerate 3.79 0.33 3.40 0.40 0.4664 Pigs (n = 12/treatment) were housed individually in metabolism crates and fed diets containing either 0 or 8% D-xylose at 4% of BW for 22 d. SCFA, short chain fatty acids

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Table 7. Effect of dietary xylose concentration and collection period on urine xylose and metabolite excretion Collection period, d 5 - 7 12 - 14 19 - 21

Dietary treatment, % Pooled T×C1 0 2 4 8 0 2 4 8 0 2 4 8 xylose inclusion SEM P-value Urinary excretion, g/d Xylose*‡† 0.51g 7.04f 21.59d 50.06a 0.49g 6.01f 16.25e 31.53c 0.83g 5.73f 16.06e 38.58b 1.28 < 0.0001 Threitol*‡† 0.12f 2.73e 6.53d 15.96a 0.08f 2.36e 5.61d 10.95c 0.27f 2.46e 6.01d 13.46b 0.45 0.0008 Xylitol*‡† 0.01f 0.07e 0.16cd 0.27b 0.01f 0.06ef 0.14d 0.23b 0.04ef 0.05ef 0.21bc 0.41a 0.02 0.0021 Xylulose*‡ 0.02e 0.08d 0.12c 0.20a 0.02e 0.08d 0.13c 0.17b 0.02e 0.07d 0.13c 0.21a 0.007 0.0021 Xylonic acid‡† 0.000cd 0.001cd 0.009cd 0.059a 0.000d 0.001de 0.009c 0.033b 0.000d 0.001d 0.008ce 0.035b 0.003 < 0.0001 Pigs (n = 12/treatment) were housed individually in metabolism crates and fed diets containing either 0, 2, 4, or 8% D-xylose at 4% of BW. Urine was collected during 3 different periods representing increasing adaptation time to treatment diets. a-gWithin a row, means without a common superscript differ (P  0.05) 1Dietary treatment × collection period interaction (T×C)

*Treatment is significant at P < 0.01 1 ‡ 63

Treatment linear contrast is significant at P < 0.01 †Collection period is significant at P < 0.01

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Table 8. Effect of dietary xylose concentration and collection period on urine energy concentration and urine sugar concentration and excretion Collection period, d 5 - 7 12 - 14 19 - 21

Dietary treatment, % Pooled T×C1 2 4 8 2 4 8 2 4 8 xylose inclusion SEM P-value Consumed xylose excreted in urine, % As xylose*‡† 31.09cd 47.17b 54.87a 26.53de 35.50c 34.56c 25.32e 35.08c 42.29b 2.34 0.0026 As threitol*‡† 12.05 14.27 17.49 10.40 12.26 12.00 10.88 13.13 14.75 0.71 0.1037 As xylitol, xylulose, or 0.94abc 0.89abc 0.80cd 0.92abc 0.90bc 0.73d 0.86abcd 1.03a 0.98ab 0.06 0.0180 xylonic acid*# Retained xylose*‡†, % 55.92bc 37.67d 26.84e 62.15ab 51.35c 52.77c 62.94a 50.77c 42.01d 2.85 0.0063 a-eWithin a row, means without a common superscript differ (P  0.05) 1Dietary treatment × collection period interaction (T×C) *Treatment is significant at P < 0.01 ‡ Treatment linear contrast is significant at P < 0.01 1 †Collection period is significant at P < 0.01 64

#Collection period is significant at P < 0.05 Pigs (n = 12/treatment) were housed individually in metabolism crates and fed diets containing either 0, 2, 4, or 8% D-xylose at 4% of BW. Urine was collected during 3 different periods representing increasing adaptation time to treatment diets.

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Figure 1. Two-way heat map visualization and hierarchical clustering of urine metabolite distribution. Pigs (n = 12/treatment) were fed diets containing either 0, 2, 4, or 8% D- xylose and 3 different collection periods were utilized to represent increasing adaptation time to treatment diets. The rows display the metabolites and the columns represent individual samples. Relative urinary metabolite excretion (g/d, log transformed) is represented in the heat map by colors, which correspond to the magnitude of difference when compared with the average value for the metabolite. Only metabolites significantly (adjusted P  0.05) impacted by treatment (% dietary xylose inclusion; 0, 2, 4, or 8%), collection period (1 = d 5-7, 2 = d 12-14, 3 = d 19-21), or their interaction are displayed. Significance was determined using two-way ANOVA.

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Figure 2. Proposed xylose metabolic pathway. Dietary D-xylose concentration (0, 2, 4, or 8%) linearly increased the urinary excretion of the compounds in bold font (P < 0.0001). Non-bolded compounds were not detected in urine samples. The color of the block arrow indicates average excretion amount of the compound in pigs from the 8% xylose treatment; red: 40 g/d, orange: 13 g/d, yellow: 0.3 g/d, white: < 0.2 g/d. Solid connecting arrows represent reactions confirmed in mammals and dashed connecting lines represent presumed reactions based on reactions occurring in microorganisms. Highlighted enzymes: 1) D-xylose 1-dehydrogenase (1.1.1.179), 2) L-gulonate 3- dehydrogenase (1.1.1.45), 3) D-erythrulose reductase (1.1.1.162), 4) aldose/aldehyde reductase (1.1.1.21), 5) L-xylulose reductase (1.1.1.10), 6) xylitol dehydrogenase (1.1.1.14), 7) xylulokinase (2.7.1.17), and 8) D-ribose dehydrogenase (1.1.1.115). The boxes indicate KEGG metabolic pathway classifications [36].

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Allocation of urine gross energy

350

300 xylitol + xylonic acid +

250 xylulose

t i

l threitol /

l 200

, 150 xylose E

G 100 N-containing compounds 50 unassociated 0 0 2 4 8 Dietary xylose inclusion, %

Figure 3. Allocation of urine gross energy (GE) from xylose, metabolites, and nitrogen (N)-containing compounds. Pigs (n = 12/treatment) were fed diets containing either 0, 2, 4, or 8% D-xylose. The presented values are treatment averages across 3 different collection periods. The effect of treatment was significant for all variables (P  0.042). Dietary xylose concentration linearly impacted each variable (linear contrast P  0.012) except the unassociated fraction. Increasing dietary xylose concentration increased the urine GE (kcal/l) contributed from xylose (SEM = 4.16), threitol (SEM = 2.05), and the combination of all lesser metabolites (xylitol, xylonic acid, and xylulose; SEM = 0.08), but decreased the GE contributed from N-containing compounds (SEM = 4.07). The amount of urine GE unexplained by the xylose, its metabolites, or N-containing compounds (SEM = 10.83) was not different among the 0, 2, and 4% treatments (P  0.416) but was lower in the 8% treatment (P = 0.042).

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CHAPTER V

INTEGRATIVE SUMMARY

General discussion

Xylose and mannose are two of the primary carbohydrates of grain and legume fiber; yet, their direct and indirect influences on swine energy metabolism are poorly understood.

As -mannanase and xylanase enzymes use continues to increase, it is important to evaluate their metabolic impact to accurately estimate enzyme effects on dietary energy availability. This information will allow nutritionists to improve the accuracy of diet formulations when high-fiber ingredients and carbohydrase enzymes are utilized.

Therefore, the overall objective of this dissertation work was to evaluate the impacts of

-mannan and xylose on energy metabolism in the pig.

-Mannanase can be added to swine diets with the objective of inhibiting a - mannan-induced immune response. This hypothesis was developed from and supported by experiments in broilers which reported that -mannanase supplementation alleviated a soybean-meal derived immune response and spared that energy for growth [1]. However, prior to our experiments, only growth trials had been published regarding the effect of - mannanase in swine [2–5]; as such, evidence supporting this dietary -mannan-induced immune response and amelioration due to -mannanase supplementation was lacking.

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Therefore, the objective of Experiment 1 (Chapter 2) was to evaluate the - mannan-induced immune response hypothesis in pigs by testing -mannanase supplementation influence of on maintenance energy requirements and to characterize systemic immune response and nutrient partitioning effects. The data from this experiment did not support the hypothesis that β-mannanase supplementation ameliorates an innate immune response induced by soybean β-mannans at a concentration of 1.3%. It is possible, however, that this concentration was too low to either effectively generate a systemic inflammatory response or to be able to measure β-mannanase supplementation effects. Therefore, Experiment 2 (Chapter 3) was a follow-up trial to evaluate the impact and interactions of dietary β-mannan concentration and β-mannanase supplementation on serum acute phase proteins, as indicators of systemic immune activation, and on growth performance in nursery pigs. This experiment provided immune data previously lacking in pig growth trials assessing -mannanase efficacy and confirmed the conclusions from the initial experiment.

In order to truly understand what immune stimulation would look like in the context of Experiment 1, a positive control treatment using LPS was included.

Importantly, this treatment allowed us to evaluate the impact of an innate immune challenge on maintenance energy requirements. Prior to this experiment, few studies had addressed comprehensive changes in energy partitioning during an immune response.

Innate immune challenge elevated pigs’ maintenance energy requirements by 23.3% and this shift in energy partitioning toward the immune response limited lipid deposition by

30.2%, leading to an 18.3% decrease in ADG during the immune challenge.

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It is important to understand that in the face of a true pathogen challenge, increased maintenance energy requirement compounds the issue of decreased feed – and thus energy - intake typically observed during disease outbreaks. This means that during health challenges, energy available for growth is likely even more restricted than previously thought. The results of this research are most valuable when interpreted in context with the findings of other Iowa State researchers that immune stimulation increases glucose requirement [6], and pathogen challenges increase the lysine:energy requirement [7] in addition to decreasing the digestibility of specific amino acids [8].

Clearly, immune responses drastically affect energy and nutrient requirements and nutritional strategies could be implemented to alleviate resource competition and improve animal health and productivity during health challenges.

The use of exogenous carbohydrases to improve fiber utilization in swine diets presents multiple issues for nutritionists to consider, such as poor response consistency and appropriate enzyme application (ex. -mannanase). Furthermore, understanding how enzyme hydrolysis products contribute to energy availability and affect overall metabolism adds additional complexity. As interest in xylanase supplementation continues to increase, an improved understanding of xylose metabolism in the pig and its energetic value is essential for effective xylanase utilization. Based on older experiments that utilized pigs fed unrealistically high dietary concentrations [9–11], it was commonly assumed that pigs cannot effectively catabolize xylose for energy. It is possible that these high xylose concentrations overwhelmed the pig’s metabolic capacity and led to incorrect conclusions. Therefore, Experiment 3 (Chapter 4) evaluated more industry-relevant dietary xylose concentrations and tested for possible adaptation mechanisms. It was

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found that the pig can utilize xylose but does so less efficiently as more xylose is consumed; only 40 – 60% of consumed xylose was retained. Furthermore, pigs can adapt over time to improve xylose utilization efficiently. By applying metabolomic approaches, urinary metabolites of xylose were identified and quantified. This information facilitated construction of a comprehensive pathway for xylose metabolism in the pig.

Overall, this dissertation improved our understanding of how -mannanase and xylanase may impact energy metabolism in the pig. This research also furthered the understanding of how immune activation can repartition energy and increase maintenance energy requirements. Understanding the extent to which energy requirements and nutrient deposition change in pigs experiencing sustained immune stress may help develop more effective feeding strategies for health-challenged herds and encourage appreciation for the economic benefits of maintaining high-health populations. Furthermore, this research was the first to address xylose metabolism in pigs using industry-relevant dietary concentrations and will substantially further the scientific community’s understanding of xylose metabolism in mammals, in addition to helping swine nutritionists determine the energetic value of fiber in pig diets.

Recommendations for future research

Future directions to expand upon this work include evaluating xylose effects on overall energy metabolism and regulation mechanisms by measuring differential gene expression and protein abundance in key metabolic tissues, such as the liver and kidney

[12,13]. This approach would also be of interest in the muscle, which may be able to utilize or accumulate xylose [13–15], and the intestinal mucosa, to potentially identify mechanisms of absorption [16]. Such data would further inform how xylose is

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metabolized and potentially provide more empirical evidence to support the proposed xylose metabolic pathway. Additionally, there is a need for more comprehensive understanding of xylanase vivo mode of action and hydrolysis products. Specifically, quantifying the xylanase release efficiency of free xylose in the small intestine would provide clearer context in which to interpret the results of Experiment 3. With a greater understanding of the nutritional and metabolic influences of dietary carbohydrates and carbohydrases, we can improve the productivity and efficiency of pork production.

Literature cited

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2. Yoon SY, Yang YX, Shinde PL, Choi JY, Kim JS, Kim YW, et al. Effects of mannanase and distillers dried grain with solubles on growth performance, nutrient digestibility, and carcass characteristics of grower-finisher pigs. J. Anim. Sci. 2010;88:181–91.

3. Cho JH, Kim IH. Effects of beta mannanase and xylanase supplementation in low energy density diets on performances, nutrient digestibility, blood profiles and meat quality in finishing pigs. Asian J. Anim. Vet. Adv. 2013. p. 622–30.

4. Pettey LA, Carter SD, Senne BW, Shriver JA. Effects of beta-mannanase addition to corn-soybean meal diets on growth performance, carcass traits, and nutrient digestibility of weanling and growing-finishing pigs. J. Anim. Sci. 2002;80:1012– 9.

5. Lv JN, Chen YQ, Guo XJ, Piao XS, Cao YH, Dong B. Effects of supplementation of β-mannanase in corn-soybean meal diets on performance and nutrient digestibility in growing pigs. Asian-Australasian J. Anim. Sci. 2013;26:579–87.

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15. Gibson WR, Bourne AR, Sernia C. D-Xylose transport in isolated skeletal muscle of chickens: Effects of insulin and tolbutamide. Comp. Biochem. Physiol. C. 1980;67:41–7.

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