© 2010 Erica Nehrling

THE EFFECT OF BUTYRATE-SUPPLEMENTED TOTAL PARENTERAL NUTRITION ON

MUC2 MRNA AND GOBLET CELL CHEMOTYPES IN A SHORT BOWEL SYNDROME NEONATAL PIGLET MODEL

BY

ERICA W. NEHRLING

THESIS

Submitted in partial fulfillment of the requirements for the degree of Master of Science in Nutritional Sciences

in the Graduate College of the University of Illinois at Urbana-Champaign, 2010

Urbana, Illinois

Master’s Committee: Professor Matthew Wallig, Chair Professor Kelly Tappenden, Director of Research Professor Rex Gaskins

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ABSTRACT

Background: Compromised barrier function and bacterial translocation are common

complications of short bowel syndrome (SBS) and parenteral nutrition (PN). Although mucins help

prevent intestinal bacteria translocation, total mucin production has been shown to decrease during

PN infusion. Butyrate administration may be a mucin fortifying strategy as it has been shown to

increase mucin production in vivo and ex vivo.

Objective: We tested the hypothesis that infusion of TPN supplemented with 9 mM butyrate will

enhance protective goblet cell chemotypes in the small intestine, specifically sulfomucins, that

enhancement of protective goblet cell chemotypes in the SCFA-supplemented group will be

observed, since this group also contains 9 mM butyrate, that butyrate-supplemented TPN will have a

greater effect on the small intestine than the large intestine, while butyrate supplementation at 60

mM will have a greater effect on the large intestine than the small intestine. We also hypothesize

that muc2 mRNA abundance will increase in the small intestine with butyrate and SCFA-

supplemented TPN infusion, whereas colonic muc2 mRNA abundance will increase in the colon

with 60 mM butyrate administration.

Methods: Forty-eight hour old piglets underwent 80% proximal jejunuoileal resection and were

randomized to one of four groups: control total parenteral nutrition (TPN), TPN supplemented

with 60 mM SCFA (36 mM acetate, 15 mM propionate, and 9 mM butyrate; SCFA), 9 mM butyrate

(9Bu) or 60 mM butyrate (60Bu). Animals were further randomized to an acute (12 hour) or chronic

(72 hour) time point. muc2 mRNA abundance, total goblet cell number, and total sulfomucin,

sialomucin, acidomucin, and neutral mucin goblet cell numbers were ascertained.

Results: Sulfomucin chemotypes were increased in the jejunal villi of the 9Bu group compared to

control (treatment main effect, p=0.047). Acidomucin chemotypes in ileal crypts were greater in the

60Bu group than the control group (treatment main effect, p=0.038). In the colonic crypts, SCFA

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groups tended to have greater acidomucin chemotypes than control at 12h while 60Bu group tended

to have greater acidomucin chemotypes per depth than control at 72h (p=0.060; Table 2.3). muc2

mRNA was increased in jejunal tissues in the 9Bu group compared to control with 270% and 30%

increases in muc2 mRNA at 12 and 72 hours, respectively (p=0.010).

Conclusion: The 9 mM Bu treatment increased protective goblet cell chemotypes and muc2 mRNA

in the jejunum while 60 mM Bu increased protective goblet cell chemotypes in the ileum and colon.

The 9Bu treatment was the most effective at upregulating muc2 mRNA abundance and sulfomucin

chemotypes. Butyrate-supplemented PN may be a therapeutic strategy for bolstering the epithelial

barrier by increasing mucin production in neonates with SBS on PN.

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ACKNOWLEDGEMENTS

First, I would like to thank my advisor, Dr. Kelly Tappenden, for her scientific brilliance and her

abounding energy which has served as such an inspiration to me. I also want to thank her for the

support and effort she put into developing my scientific mind over the last three years.

I would also like to thank Dr. Sharon Donovan, Director of Nutritional Sciences during most of my

time here at the University of Illinois, and Linda Barenthin, administrative secretary of the

Nutritional Sciences department until she retired a couple months ago. They fielded endless

questions, starting before I even ventured to the University of Illinois at Urbana-Champaign for

recruiting weekend in March 2007 and not ending until I deposited this thesis. As I have said in so

many emails and in person: thank you so much for your time.

I would also like to thank my committee members Dr. George Fahey, Dr. Rex Gaskins and Dr.

Matthew Wallig for all of their support in meeting with me when I needed help, for answering all of

my questions, and for directing me to great resources here on campus.

I would like to thank the members of the Tappenden lab, past and present, for their scientific savvy

and knowledge, energy in the many early mornings to help me with my piglet study, and time spent

discussing assays, science, statistics, and life.

I would like to thank my sister, Adrianne Nehrling Edwards, and my good friend and lab mate,

Jennifer Woodard for their support. They both offered me a wealth of scientific knowledge and

emotional support for the inevitable times at which science can be frustrating and troubleshooting

seems endless.

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Lastly, I would like to thank my parents, for their endless support in getting me from preschool to

elementary school to middle school to high school to undergraduate college to graduate school. I

love you two very much, and this thesis is a reflection of what wonderful and supportive parents you

are.

Thank you all so very much.

TABLE OF CONTENTS

LIST OF ABBREVIATIONS................................................................................................... VII

CHAPTER 1: LITERATURE REVIEW .................................................................................... 1

INTRODUCTION ..................................................................................................................... 1 THE NEONATAL INTESTINE ................................................................................................ 2 INTESTINAL FAILURE .......................................................................................................... 12 MANAGEMENT OF INTESTINAL FAILURE ...................................................................... 18 MODULATION OF MUCIN PRODUCTION ........................................................................ 20 RESEARCH OBJECTIVE ........................................................................................................ 27

CHAPTER 2: THE EFFECT OF BUTYRATE-SUPPLEMENTED TOTAL PARENTERAL NUTRITION ON MUC2 MRNA AND GOBLET CELL CHEMOTYPES IN A SHORT BOWEL SYNDROME NEONATAL PIGLET MODEL .............................. 30

ABSTRACT ............................................................................................................................... 30 INTRODUCTION ................................................................................................................... 31 MATERIALS AND METHODS............................................................................................... 33 RESULTS .................................................................................................................................. 41 DISCUSSION ........................................................................................................................... 42

CHAPTER 3: FUTURE DIRECTIONS .................................................................................. 70

REFERENCES ........................................................................................................................... 73

CURRICULUM VITAE ............................................................................................................. 90

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LIST OF ABBREVIATIONS

ANOVA analysis of variance CaCo2 human epithelial colorectal adenocarcinoma cell line CCD-18Co intestinal colonic myofibroblast cell line cDNA complementary deoxyribonucleic acid Cl.16E clonal derivative of HT29 cell line ddH20 double distilled water DEPC diethylpyrocarbonate DMEM dulbecco’s modified eagle medium DNA deoxyribonucleic acid dNTP deoxyribonucleotide triphosphate EGF epidermal growth factor EN enteral nutrition EST expressed sequence tag, NCBI database FAE follicle associated epithelium FITC fluorescein isothiocyanate FSR fractional synthetic rate GALT gut associated lymphoid tissue GH growth hormone GLP-2 glucagon-like peptide 2 Glut-2 glucose transporter 2 HIDAB high iron diamine alcian blue HT29 human colon adenocarcinoma cell line IBD inflammatory bowel disease IF intestinal failure ITF intestinal trefoil factor IL-10-/- interleukin-10 knockout IL-10 interleukin-10 JAM junction adhesion molecule LS174T human colon carcinoma cell line M cell microfold cell µg microgram µL microliter MLN mesenteric lymph nodes mM millimolar MODS multiple organ dysfunction syndrome mRNA messenger ribonucleic acid muc2 mucin 2 muc2-/- mucin 2 knockout Na+K+ATPase sodium potsassium adenosine triphosphatase Na S1 sodium sulfate transporter NaS-/- sodium sulfate transporter knockout NCBI National Center for Biotechnology Information PASAB periodic acid schiff’s alcian blue PGE1 prostaglandin E1 PGE2 prostaglandin E1 PN parenteral nutrition

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PNALD parenteral nutrition-associated liver disease RER rough endoplasmic reticulum RIN RNA integrity number RLMβ resistin-like molecule β RNA ribonucleic acid RNase ribonuclease rt-PCR real time polymerase chain reaction SBBO small bowel bacterial overgrowth SBS short bowel syndrome SCFA short chain fatty acid SE standard error SED subepithelial dome SGLT-1 sodium glucose transporter 1 sIgA secretory immunoglobulin A T84 human colon carcinoma cell line TPN total parenteral nutrition UV ultraviolet vWF von Willebrand factor wt wild type ZO-1 zona occludin-1

CHAPTER 1 LITERATURE REVIEW

INTRODUCTION

Short bowel syndrome (SBS) is the most common cause of intestinal failure in infants

(Goulet and Ruemmele, 2006) and occurs in 4.4% of preterm infants with only a 73% to 89.7%

survival rate (Quiros-Tejeira et al., 2004; Wales et al., 2004). Treatment often includes nutrition

management in the form of parenteral nutrition (PN), which can extend the life of a patient with

SBS for many years (Dudrick et al., 1968; Wilmore and Dudrick, 1968; Dudrick and Allen, 1971).

However, long-term administration of PN can compromise quality of life and can result in

potentially fatal primary complications such as parenteral nutrition-associated liver disease

(PNALD), recurrent sepsis, and uncontrolled fluid losses, or secondary complications such as

bacterial translocation, barrier dysfunction, muted intestinal adaptation, and multiple organ

dysfunction syndrome (MODS) (Johnson et al., 1975; Morin et al., 1978; Ford et al., 1984; Deitch,

1992; Inoue et al., 1993; Conour et al., 2002; Yang et al., 2002; Ziegler et al., 2002; Dahly et al., 2003;

Kansagra et al., 2003; Wiest and Rath, 2003; Kumpf, 2006; Grau et al., 2007; Jan, 2008).

Within the first line of defense against invading pathogens—innate immunity—is intestinal

mucous, made predominantly of the glycoprotein mucin. Decreases in mucin abundance or in

protective goblet cell chemotypes can leave the epithelial barrier exposed to bacteria and bacterial

products which can lead to bacterial translocation (Gork et al., 1999). In addition, direct contact of

harmful luminal contents to the epithelial lining promotes inflammation, which can further impair

barrier function (Yang et al., 2002; Yang et al., 2008) and increase the risk of bacterial translocation.

Further, it has been shown that parenteral nutrition-associated liver disease possibly results from

bacterial translocation (Sondheimer et al., 1998). Because the liver is responsible for clearing toxins

from the body, liver disease can lead to MODS. Bolstering one of the initial defenses, mucin, may

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improve the outcome of SBS patients by preventing bacteria-induced mucosal inflammation and

further complicatoins.

Mucin abundance is decreased by the typical nutritional management of SBS, PN

administration (Iiboshi et al., 1994; Bertolo et al., 1998; Law et al., 2007). However, PN increases

protective goblet cell chemotypes (Conour et al., 2002). The short chain fatty acid (SCFA) butyrate

enhances mucin production in vitro and ex vivo (Finnie et al., 1995; Willemsen et al., 2003; Hatayama

et al., 2007; Burger-van Paassen et al., 2009), but its direct effect on mucin in vivo and its effect via

parenteral administration has not yet been elucidated. This study analyzes the effect of parenteral

butyrate and SCFA administration on both mucin mRNA and goblet cell chemotypes in a SBS

model piglet receiving total parenteral nutrition (TPN).

THE NEONATAL INTESTINE

Intestinal Cells

Four main layers of tissue compose the intestinal wall: the mucosa, submucosa, muscularis

externa, and serosa (Saladin, 2004). The mucosa layer is closest to the lumen and is comprised of a

luminal layer of epithelial cells followed by the lamina propria and muscularis mucosa layers (Madara

and Trier, 1994). The anatomy of the small intestine, from the nature of the folds to the surface of

the cells, creates a vast area that allows for digestion and absorption of nutrients. The folds of

Kerckring occur along the mucosa of the small intestine, and upon this surface area are projections

called villi, fingerlike protrusions of the mucosal layer. Extending from the surface of each

absorptive cell of the villi are microvilli which contain the brush border enzymes necessary for

digestion. Crypts of Lieberkühn—similar to pores—descend from the epithelium to the muscularis

mucosa (Saladin, 2004).

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Stem cells located in the lower half of the crypts differentiate into one of five kinds of

epithelial cells: enterocytes, goblet cells, enteroendocrine cells, paneth cells, and M cells, also known

as microfold cells (Leedham et al., 2005). Epithelial cells migrate from the crypt to the tip of the

villus where they are sloughed off by luminal contents and consequently digested. Enterocytes are

responsible for continuous absorption of macromolecules and micronutrients from the lumen and

transportation of the contents to the circulation and lymph, and they are present in the villi and the

upper half of the crypt epithelium (Kimmich and Randles, 1975; Iuldashev et al., 1978; Saladin,

2004). Goblet cells are located in these same areas as enterocytes; however, they descend further

into the crypts and have the primary role of mucin production and secretion (Moe, 1953; Forstner et

al., 1973). Enteroendocrine cells are found along the epithelium and are responsible for releasing

peptide hormones of both endocrine and paracrine function. Paneth cells are the only epithelial cell

line present at the base of the crypts and extend about halfway up the crypt walls and secrete

defensins, lysozyme, and phospholipase to protect against harmful luminal contents (Trier, 1963; Qu

et al., 1996; Ayabe et al., 2000). Last, M cells are found in the follicle associated epithelium (FAE) of

Peyer’s patches in the ileum and are responsible for antigen sampling and transportation (see below;

Owen and Jones, 1974).

Epithelial cells are held together by tight junctions which are located between cells at the

apical end of the cell, and desmosomes, located directly beneath tight junctions. Tight junctions

restrict paracellular transport (Claude and Goodenough, 1973) and consist of the specific proteins

zonula occludens-1 (ZO-1; Stevenson et al., 1986), cingulin (Citi et al., 1988), junctional adhesion

molecules (JAM; Martin-Padura et al., 1998), claudin-1 and claudin-2 (Furuse et al., 1998), and

occludin (Furuse et al., 1993). Desmosomes are more important for cell to cell adhesion than

paracellular restriction and are comprised of the protein E-cadherin (Boller et al., 1985). Tight

junction structure is very plastic in nature and can be modulated by physiological, physical, and

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pathological occurrences. For example, intestinal inflammation has been shown to reduce barrier

function by causing tight junction transmembrane proteins to internalize and by disrupting the

protein’s membrane affiliations (Bruewer et al., 2003).

Enteric Immune System

The intestine has extensive immunological function, as it is in contact with both the external

environment (e.g. food and ingested bacteria) and the internal resident microbiota. The innate

immune system is a highly conserved type of immunity across organisms, and operates as both the

first line of defense against pathogens and as an activator of the acquired immune system, which is

present in more advanced organisms and involves recognition of foreign substances from previous

encounters (Yuan and Walker, 2004). The intestine bridges to the acquired immune system via M

cell and dendritic cell interaction.

The intestinal immune system is termed GALT, or gut associated lymphoid tissue, and

consists of oval bundles of lymphoid tissue known as Peyer’s patches which are covered by follicular

associated epithelium (FAE; Figure 1.1). Harmful luminal products or bacteria are endocytosed or

phagocytosed, respectively, by M cells in the FAE and are trafficked to the base of the cell (Owen

and Jones, 1974; Neutra et al., 1996). The bases of M cells are expanded to form a pocket like

invaginations containing T cells, B cells, and macrophages (Farstad et al., 1994); however, it has not

been clearly shown that these lymphocytes and macrophages are within the M cell plasma

membrane. In addition, the M cell pocket is only characteristic of a mature M cell; newborn piglets

have immature M cells that have not yet formed the pocket like invaginations (Kido et al., 2003).

Below the FAE, in the lamina propria, is the subepithelial dome (SED) which contains strategically

placed dendritic cells, presumably to begin processing M cell admitted antigens for presentation to T

cells (Kelsall and Strober, 1996). Below the SED are lymphoid follicles (Owen, 1977) containing B

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lymphocyte germinal centers (Butcher et al., 1982). Between the follicles are the interfollicular

regions that are rich in T cells and macrophages (Kelsall and Strober, 1996). Typically, dendritic

cells are responsible for the connection between the innate immune system and the acquired

immune system by recognizing antigens engulfed by the M cells and presenting them to T cells

(Langhoff and Steinman, 1989) located in either the M cell pocket (Pappo, 1989; Pappo et al., 1991),

the lamina propria, or the nearest mesenteric lymph node (MLN; Holt et al., 1994). However, M

cells also have processes that protrude from their basal membrane and extend as far as 10 µm and

potentially make contact directly to lymphoid tissue (Giannasca et al., 1994). In addition, epithelial-

associated dendritic cells have been shown to extend between intestinal epithelial cells, forming tight

junctions with these cells, and directly sampling antigens from the lumen without the help of the M

cell (Rescigno et al., 2001; Niess et al., 2005). These epithelium associated dendritic cells are also

capable of traveling to the MLNs (Schulz et al., 2009).

Bacterial translocation has been established as a transcellular process, specifically through

FAE M cells (Grutzkau et al., 1990; Jones et al., 1994; Jensen et al., 1998); however, there has been

recent conjecture that bacterial translocation may also occur paracellularly (Nadler et al., 2000). In

this ex vivo study by Nadler and colleagues, pinocytosis and phagocytosis were alternatively inhibited

with no change noted in bacterial translocation meaning that paracellular transport is a likely

mechanism for bacterial translocation. Although, both mechanisms of transcellular transport were

not inhibited at the same time meaning that when one is restricted, the other mechanism may

compensate. Further research in the area of paracellular bacterial translocation is needed.

Because M cells function to transport bacteria and bacterial products for immune function,

they consequently have absent or minimal microvilli and greatly shortened glycocalyx (Bye et al.,

1984; Neutra et al., 1999). Mucous, a viscoelastic gel-like covering over the intestinal epithelium is

made mainly from goblet cell mucin secretion. Because goblet cell secretions could block M cell

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engulfment, they are rare in the FAE of human intestinal cells and have been precisely calculated to

account for only 0.5% of the cells in porcine FAE (Fujimura et al., 1992; Beier and Gebert, 1998). It

may seem that increases in muc2 mRNA, and potentially more Muc2 protein and mucous

production, or more protective goblet cell chemotypes would not likely affect bacterial translocation

since M cells are primed for bacterial engulfment and have few neighboring protective goblet cells.

However, mucous is still important in preventing bacteria-induced inflammation in the epithelium

apart from the FAE. As previously mentioned, inflammation can result in barrier function damage.

Furthermore, although definitive paracellular transport of bacteria and bacterial products has not

been established, protective mucin types and mucous abundance are correlated with decreased

bacterial translocation in vitro and in vivo, demonstrating the advantage of specific mucins in

protection against bacterial translocation (Gork et al., 1999; Sakamoto et al., 2004; Dawson et al.,

2009).

Mucins

Mucins and Intestinal Function

Mucous is a semi-permeable viscoelastic gel coating that protects the intestinal epithelium

from enzymatic, chemical, and mechanical damage by providing barrier function making it essential

in innate immunity. The main component of mucous is mucin, which is secreted from goblet cells;

however, mucous also consists of other secretory components that confer protection to the

intestine. Secretory immunoglobulin A (sIgA), a mucosal antibody secreted from intestinal plasma

cells (Perkkio and Savilahti, 1980), is present in the intestinal mucous (Hamer et al., 2009). Paneth

cells contribute to intestinal defense by secreting the microbiocidal peptides lysozyme (Fleming,

1922; Trier, 1963), defensins (Ayabe et al., 2000), and phospholipase (Qu et al., 1996). In addition to

mucin, goblet cells secrete intestinal trefoil factor (ITF; Podolsky et al., 1993), a peptide thought to

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safeguard the epithelial layer by encouraging healing upon insult (Thim, 1989; Wright et al., 1990;

Alison et al., 1995). In ITF-deficient mice, the administration of dextran sulfate sodium—an agent

that typically causes only mild intestinal injury—caused death from severe colitis (Mashimo et al.,

1996). Goblet cells also secrete resistin-like molecule β (RLMβ ), a protein that promotes

inflammation (Nair et al., 2008). While excessive inflammation can damage the epithelial layer, some

inflammation is necessary and is a natural reaction against pathogens.

Mucins are generally categorized into two groups: secreted mucins (gel-forming and non-gel-

forming) and membrane-anchored mucins. The majority of intestinal mucous is made of secretory

gel-forming mucins, and contains predominantly Muc2, a mucous protein encoded by the muc2 gene

(Gum et al., 1989; Griffiths et al., 1990). Muc2 is generally considered to be a secretory rather than a

membrane bound mucin, but some have found that Muc2 contributes to two different layers: one

fraction completely detached from the epithelium that is easily washed away with saline and a

fraction that is adherent to the epithelium (Johansson et al., 2008). The loose layer of mucous is

thickest in the colon and thinnest in the jejunum. The adherent layer is continuous and firm in the

colon but is thinner or non-existent in the small intestine (Atuma et al., 2001).

Mucins are heavily glycosylated proteins composed of approximately 75%-80% carbohydrate

and 20-25% amino acids (Carlstedt et al., 1993). While Muc2 contains a significant amount of

tandemly repeated, heavily O-glycosylated regions (Toribara et al., 1991) made mainly of serine,

threonine, and proline (Fahim et al., 1987; Huan et al., 1992), the mucin ends are unique and have

fewer serine and threonine residues and contain more cysteine residues (Gum et al., 1992). On

average, there are 4 monosaccharides on the carbohydrate side chains (Carlstedt et al., 1993), which

sometimes contain sulfate (Carlstedt et al., 1993) and sialic acid (Hansson et al., 1991) residues.

The structure of mucin is central to its function. The cysteine-rich ends of mucins serve to

aid polymerization of mucin monomers into oligomers, which has been demonstrated by treating

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mucins with disulfide-specific reducing agents and observing a subsequent loss of viscosity and gel-

forming ability in the mucin (Tabachnik et al., 1981). In addition, the cysteine-rich ends are very

similar to von Willebrand factor (vWF) (Gum et al., 1994), a glycoprotein found in endothelium,

essential in blood coagulation. Cysteine-rich ends are crucial to oligomerization and packaging of

vWF into granules, suggesting by analogy that cysteine-rich ends may be important to intestinal

mucin polymeriazation and packaging as well. (Xu et al., 1992).

Their heavily glycosylated core gives mucins their expanded and viscoelastic structures and

allows for solubility in the intestinal lumen. Removal of the carbohydrate moieties results in a

denatured protein, demonstrating that the glycosylated side chains are essential to mucin structure

(Shogren et al., 1989). In addition, it has been calculated that mucins with 2 or more carbohydrate

residues per side chain results in a 2.5-3 fold increase in size in solution compared to a non-

glycosylated mucin (Shogren et al., 1986). The expanded structure means that mucins overlap and

become entangled, giving rise to the viscoelastic properties of mucin (Shogren et al., 1989).

Viscoelasticity of intestinal mucous is necessary to withstand the mechanical stress of both the

muscular force of the intestine, such as peristalsis, and the movement of particles across the mucous

and epithelium. In other words, the viscoelasticity of mucous ultimately protects the epithelium

from mechanical damage (Bansil et al., 1995). Because mucous serves as a physical barrier, it also

protects the epithelium against chemical and enzymatic damage. Lastly, the sulfate and sialic acid

residues create a high density negative surface charge which binds water, further enhancing

viscoelastic properties.

In addition to the protection that mucin provides against the chemical, enzymatic, and

mechanical stress of normal gastrointestinal function, they also provide protection from bacteria

ingested or residing in the intestine. First, mucin protects the epithelium by trapping bacteria that

subsequently get excreted with the loose layer of mucous in the feces. In a study by Johansson and

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colleagues (2008), it was found that in the colon, the loose layer of mucin, which is excreted in the

feces, is colonized by bacteria, while the inner layer is almost devoid of bacteria, possibly reflecting

the effective barrier mucin provides. Further, in this study, muc2-/- mice had bacteria in direct

contact with the colonic epithelium, demonstrating the necessity of mucin in preventing bacterial

contact with the epithelium and possibly even preventing bacterial translocation (Johansson et al.,

2008).

Secondly, mucin serves as a physical barrier that is difficult for the bacteria to break down.

The type of carbohydrate side chains on mucin plays a role in the protectiveness of the mucin

against bacterial penetration. Upon bacterial introduction in vitro to germ-free rat mucin,

sulfomucins and neutral mucins were 40% and 55% degraded, respectively, at 4 hours while

sialomucins were 90% degraded at 1 hour and almost completely degraded at 4 hours (Fontaine et

al., 1998). Thus, sulfomucins and neutral mucins confer a more protective effect in the intestine

than sialomucins because they are more slowly degraded by bacteria, therefore the established

continuum of mucin protection is as follows: sulfomucins > neutral mucins >>> sialomucins.

An earlier study also suggested the benefit of acidomucins (both sialomucins and

sulfomucins) over neutral mucins. The colonization of germ-free rats with rat-specific intestinal

microbiota triggered an initial release of neutral mucins with a progressive shift towards acidic mucin

secretion observed over the following 4 months (Enss et al., 1992). In this study, the researchers

suggested that the replacement of neutral mucins with acidomucins over this time frame was in

response to the development of gastrointestinal microbiota and was possibly a form of maturation

of the gastrointestinal tract (Enss et al., 1992).

Several studies confirm the protective nature of sulfomucins (Sakata and von Engelhardt,

1981; Deplancke et al., 2000; Conour et al., 2002; Dawson et al., 2009). Mice deficient in sulfate

transporters had higher levels of sialomucin than sulfomucin levels compared to control mice, and

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these mice had greater susceptibility to toxin-induced colitis and exhibited decreased intestinal

barrier function (Dawson et al., 2009). This is most likely attributed to the lack of sulfate residues

provided intracellularly by sulfate transporters resulting in decreased sulfate addition during post-

translational mucin modification (Dawson et al., 2009). Further, it has been demonstrated that

sulfomucins increase in concentration from the duodenum to the colon, mirroring the same pattern

of bacterial concentration from proximal to distal bowel (Sakata and von Engelhardt, 1981;

Deplancke and Gaskins, 2001; Conour et al., 2002). The increase of sulfomucins in areas of higher

microbial density suggests the importance of sulfomucins in barrier defense against resident and

infectious microbial populations.

As previously mentioned, mucins serve to trap bacteria. The action of trapping bacteria also

allows bacteria to congregate which has been shown in vitro to aid in the growth of biofilms on fixed

Caco-2 human epithelial cells (Bollinger et al., 2003). While this may seem contrary to the purpose

of trapping bacteria, and while mucous can aid in the biofilm formation of harmful species

(Bollinger et al., 2003), biofilms have also been shown to be beneficial to the intestine and host

health (Jones and Versalovic, 2009). In addition to binding each other, bacteria bind receptor sites

on mucin which serve to compete with bacterial receptor sites on the underlying epithelium

(Wadolkowski et al., 1988; Conway et al., 1990). Scientists have demonstrated that some pathogens

have adapted to using these binding sites as temporary attachment while moving towards the

epithelial layer (Drumm et al., 1988; McCormick et al., 1988; Conway et al., 1990). These same

scientists also demonstrated that the pathogens lost their piliation, binding ability to epithelial cells,

or the ability to tumble and swim after colonizing intestinal mucous. So it may be that intestinal

mucous aids pathogens in reaching the epithelium, but may inhibit the bacteria’s ability to actually

bind to the epithelium. If enough mucous is present, it is likely that excretion of the pathogens

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while they are attached to the mucous can occur before the rate of replication of pathogens becomes

threatening (Forstner, 1994).

Synthesis and Excretion of Mucins

Muc2 production appears to occur solely in goblet cells (Van Klinken et al., 1995). Within

the goblet cells, mucins are synthesized similarly to most packaged secretory glycoproteins. They are

first synthesized on ribosomes and then transported to the rough endoplasmic reticulum (RER)

where they are folded and oligomerized. Carbohydrate side chain additions begin cotranslationally

via sulfotransferases, sialytransferases, and glycosyltransferases at the ribosomal level, continue at the

RER (Strous, 1979), and the main carbohydrate side chain additions occur within the Golgi complex

(Dekker and Strous, 1990). Condensing granules containing the mucins bud off from the trans-

Golgi and mucins are increasingly condensed with the help of increasing intragranular

concentrations of cationic calcium (Paz et al., 2003). As goblet cells mature, granules are separated

from the inner surface of the plasma membrane by a thin web of actin filaments. The mucins are

released when the granule’s membrane fuses with the cell plasma membrane, creating a fusion pore,

and then releasing the mucin via exocytosis through a dialating pore (Breckenridge and Almers,

1987). This process can be either constitutive or regulated.

In constitutive pathways, granules are transported directly to the plasma membrane and

release mucin without any signal, only requiring contact with the membrane. Several cell types have

been shown to release small amounts of glycoprotein continuously via the constitutive pathway

while also retaining the ability to upregulate mucin secretion in response to an environmental signal

(regulated pathway; Gumbiner and Kelly, 1982; Sporn et al., 1986; Arvan and Chang, 1987; Burgess

and Kelly, 1987). The mechanism regarding the movement of mucins out of granules is not fully

elucidated, but one hypothesis is that the smaller calcium ions escape first after the fusion pore

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opens, leaving highly negative mucins to strongly repel each other, which pushes them out of the

granule (Forstner, 1994). The hydrophilic mucins then become a viscous gel. Complete refilling of

the goblet cell takes approximately 60-120 minutes (Specian et al., 1990). Once full, the goblet cell is

then ready to release more mucin. The protection provided by mucous is most important in those

with intestinal problems, such as intestinal failure, since many of these conditions predispose the

intestine to further damaging events such as bacterial translocation into the systemic circulation or

organs.

INTESTINAL FAILURE

Etiology of Intestinal Failure

Intestinal failure (IF) is defined as the critical reduction of functional bowel below the

minimum amount needed for adequate absorption of the basic metabolic requirement without oral

or intravaneous supplementation (Jeejeebhoy, 2005). This decrease in function can result in severe

malabsorption without supplementation and can be caused either by short bowel syndrome (SBS)

which results from surgical resection of the bowel or intestinal failure which is non-functioning (but

not necrotic) sections of the bowel that don’t necessarily need to be removed. Other causes of IF,

besides SBS, include severe extended Hirschsprung’s disease (a motility disorder), chronic intestinal

pseudo-obstruction, mesenteric venous thrombosis, necrotizing enterocolitis, gastroschisis, midgut

volvulus, desmoids tumor, intestinal atresia, trauma, and Crohn’s disease (Hyman et al., 1994;

Nishida et al., 2002). Conditions like these in which functional surface area, motility and/or

obstruction are issues, intravenous supplementation is often necessary since enteral nutrition (EN)

would not be viable. In the past, severe malabsorption plagued individuals with IF. However,

patients with intestinal failure are now living longer due to the advent of PN in the mid-1970’s and

vast improvements to PN over the last 3 decades (Langnas, 2008).

12

Intestinal Failure in Infants

IF is more common in preterm infants than term infants, as preterm infants typically suffer

from decreased gastric motility, fat malabsorption, and digestive enzyme deficiencies such as lactase

deficiency (Klaus and Fanaroff, 2001; American Academy of Pediatrics Committee on Nutrition,

2004; MacLean and Fink, 1980). This means the immature gastrointestinal system of a premature

infant is likely predisposed to intestinal failure. Intestinal insults requiring resection can occur

prenatally, neonatally, or postnatally. Prenatal etiologies of IF include atresia, malrotation, midgut

volvulus, segmental volvulus, abdominal wall defects, gastroschisis, and extensive Hirschsprung’s

disease. Midgut volvulus can occur during both neonatal and postnatal development, especially in a

preterm infant whose gastrointestinal system is still developing. Other neonatal etiologies of IF

include necrotizing enterocolitis, arterial thrombosis, and venous thrombosis. Postnatal origins of

IF include complicated intussusceptions, arterial thrombosis, inflammatory bowel disease, post-

traumatic resection, and extensive angioma (Goulet and Ruemmele, 2006).

Complications

Basic Complications

SBS has a relatively low incidence of about 3.1 per 1000 of term infants and 43.6 per 1000 of

premature infants (Wales et al., 2004). There are many complications of SBS that are life-

threatening, and the survival rates are alarmingly low, with only 73% to 89.7% of infants surviving

(Quiros-Tejeira et al., 2004; Wales et al., 2004; Goulet et al., 2005). Better survival rates are

significantly associated with remaining post-resection bowel lengths >38 cm, remaining ileocecal

valve, remaining entire colon, and with primary anastomosis rather than an ostomy performed

during resection (Quiros-Tejeira et al., 2004). Additionally, ostomy reversal is associated with

increased survival (Quiros-Tejeira et al., 2004).

13

One of the functional complications of SBS is reduced digestive and absorptive capacity.

SBS patients have decreased absorption of all nutrients, even easily digested carbohydrates (Olesen

et al., 1999). Oftentimes, management of decreased absorption involves PN since most premature

infants have neither the reflexes to suckle nor a mature gastrointestinal system to absorb nutrients

from enteral nutrition (EN) delivered via tube feedings. Unfortunately, though PN has prolonged

the lives of many infants suffering from SBS, it can cause further declines in intestinal function and

can create many other complications by its administration. PN is associated with both

morphological and functional decline of the intestine, termed intestinal atrophy (Johnson et al.,

1975; Inoue et al., 1993). PN can have life threatening complications associated with long term use

such as liver disease, septicemia or bacterial translocation, or loss of venous access (Pierro et al.,

1996; Sondheimer et al., 1998; de Buys Roessingh et al., 2008). Fluid balance is also an issue in SBS

patients depending on what part and how much of the intestine was resected. PNALD is most

commonly related to cholestasis in the pediatric population and has been shown to occur in 67% of

infants on long term PN after intestinal resection of which 17% progressed to liver failure

(Sondheimer et al., 1998). In this study, bacterial or fungal infection of the blood, urine, peritoneal

fluid, cellulitis aspirate, or cerebrospinal fluid preceded all instances of cholestasis, with most of

these bacteria implicated in infection being common gastrointestinal bacteria, leading one to believe

that bacterial translocation could have been the factor in some or many of these cases of cholestasis

and liver failure (Sondheimer et al., 1998).

Secondary Complications

Bacterial translocation can be a life-threatening secondary complication of either SBS and/or

PN administration and is the passage of viable or non-viable bacteria and bacterial products from

the intestinal lumen through the epithelium and into the mesenteric lymph nodes, liver, spleen,

14

kidney, and bloodstream (Wells et al., 1988; Berg, 1999). Some researchers have found increased

rates of bacterial translocation in rats receiving TPN compared to rats fed either rat chow ad libitum

or TPN solution orally (Alverdy et al., 1988; Eizaguirre et al., 2001). However, other groups have

found that TPN reduces barrier function but does not increase bacterial translocation in piglets

(Nakasaki et al., 1998; Kansagra et al., 2003). As mentioned above, Sondheimer and colleagues

(1998) speculated that bacterial translocation is a likely cause of cholestasis and PNALD. Bacterial

translocation has been associated with MODS in other studies as well (Deitch, 1990; Sullivan et al.,

1991; Zhi-Yong et al., 1992; Fukushima et al., 1995). The risk of bacterial translocation has been

proposed to increase due to decreased epithelial barrier function (see below) caused by PN

administration (Deitch et al., 1995) .

It has been recently demonstrated that bacterial translocation is also correlated with small

bowel bacterial overgrowth (SBBO; Cole et al., 2010). The mechanisms are not necessarily mutually

exclusive, although the relationship between barrier function, SBBO, and bacterial translocation has

not been clearly defined. SBBO is an increase in the total number of bacteria or of certain bacteria

and can result in nutrient or micronutrient malabsorption (e.g. Vitamin B12), gas generation with

bloating, abdominal pain, diarrhea, altered motility, and kidney stones. SBBO is a complication of

both SBS and PN and may decrease the effectiveness of weaning patients off of PN. Inflammation

is often a result of SBBO, either due to certain bacterial species igniting an inflammatory response

once entering the mucosa, or the host’s overreaction to absorbed bacterial antigens (Dibaise et al.,

2006). TPN has specifically been shown to predispose neonatal piglets to overgrowth of Clostridium

perfringens, a detrimental bacteria that has been implicated in bacterial translocation (Meddah et al.,

2001; Deplancke et al., 2002).

Mucosal inflammation may contribute to the pathogenesis of bacterial translocation. Direct

contact of harmful luminal contents to the epithelial lining promotes inflammation which can

15

further impair barrier function (Yang et al., 2002; Yang et al., 2008) and potentially increase the risk

of bacterial translocation. Activation of pro-inflammatory cytokines from leukocytes, macrophages,

and mast cells (Saladin, 2004) has been shown to impair mucosal integrity and barrier function by

negatively affecting tight junction proteins (Bruewer et al., 2003).

Adequate mucin content is essential for proper barrier function and decreased mucin

content can increase the risk of bacterial translocation. Dawson and colleagues (2009) found that

sulfate transporter (NaS1) deficient (NaS-/-) mice had a marked decrease in sulfomucin content.

Without a correctly functioning NaS1 transporter, there are fewer sulfate residues available for

mucin sulfation. NaS-/- mice had a significant 24% increase in macromolecule permeability. Upon

infection with Campylobacter jejuni, 60% of NaS1-/- mice experienced bacterial translocation presenting

with Campylobacter jejuni in the liver three days after infection while none of the NaS1+/+ mice

presented with bacterial translocation. Interestingly, the NaS1-/- and NaS1+/+ mice had similar

mucosal integrity against resident microbiota, indicating that the decrease in barrier function was

only in regards to pathogenic bacterial infection. Along with the decrease in sulfomucin content, an

increase in sialomucin content of the NaS1-/- mice was noted. However, mRNA levels of mucins

normally expressed in the mouse ileum were unchanged, demonstrating that any role that mucin

levels played in increased bacterial translocation and intestinal macromolecular permeability in the

NaS1-/- mice was likely related to a decrease in the sulfomucin to sialomucin ratio (Dawson et al.,

2009). However, total mucin production was not measured; thus, increases in bacterial translocation

in the NaS1-/- mice could be due to total decreased mucin abundance rather than solely decreased

relative sulfomucin content.

The relationship between inflammation and sulfomucins has also been investigated. Muc2-/-

mice downregulate several genes, including essential mucin synthesis enzymes such as

sulfotransferases (Yang et al., 2008). Downregulation of sulfotransferases will result in decreases in

16

sulfomucin content. This study also demonstrated that Muc2-/- mice developed tumors through an

inflammation linked pathway and that Muc2 deficiency resulted in subclinical chronic inflammation

(Yang et al., 2008). Makkink and colleagues (2002) discovered that IL-10-/- mice had sulfate depleted

mucins compared to control mice, and that IL-10-/- mice had correspondingly inflamed colons. IL-

10 is an anti-inflammatory cytokine first described in 1989 (Fiorentino et al., 1989). The lack of IL-

10 could easily cause inflammatory conditions such as inflammatory bowel disease (IBD) or colitis,

and Makkink and colleagues’ findings suggest that decreased mucin sulfation could be part of the

mechanism by which intestinal inflammatory conditions arise. Evidence also demonstrates that

administration of IL-10 can decrease TPN-associated declines in tight junction proteins in mice (Sun

et al., 2008), suggesting that a pro-inflammatory response is at least partly responsible for reduced

barrier function, either by tight junction dysfunction or by decreases in mucin sulfation.

Mucins, Bowel Resection, and Nutrition Support

The fractional synthetic rate (FSR) is the rate of incorporation of a precursor into a product

per unit of product mass and has been used in many studies to analyze protein synthesis and

incorporation. The FSR of Muc2 in enterally fed human preterm infants after small bowel resection

was 67.2%, which is one of the highest FSRs determined in humans to date. This was assessed by

analyzing labeled threonine incorporation into mucins (Schaart et al., 2009).

The effect of nutritional support on the rate of mucin synthesis is debated. Conour and

collegues (2002) found an increase in the number of acidomucin expressing goblet cells in newborn

piglets administered TPN. However, they did not quantify mucin abundance, so mucin decreases

may be happening concomitantly with goblet cell number stasis or increases.

Recent reports demonstrated definitive decreases in total mucin content or mucin

production with TPN administration. After administration of fluorescin isothiocyanate (FITC)-

17

dextran through a gastroduodenal tube, FITC-dextran descended between the villi in the TPN

administered rats while increased mucous gel in the chow-fed group prevented the FITC-dextran

dye from filling the inter-villi spaces and retained the dye to the lumen area (Iiboshi et al., 1994)

demonstrating the increased mucous gel in the chow fed rats. This group also found that FITC-

dextran plasma levels were elevated in TPN-administered rats compared to chow-fed rats, indicating

increased permeability in the intestines of rats receiving TPN. Another study in which rats were

administered TPN intravenously or fed with the elemental TPN solution orally, epithelial bound

mucin and mucin within the goblet cells was significantly decreased in the jejunum, with a similar

trend seen in the ileum (Spaeth et al., 1994). In addition to nutrition support, micronutrient

supplementation affects mucin production as well. Comparing adequate threonine EN piglets,

adequate threonine PN piglets, and deficient threonine EN piglets, PN resulted in lower mucin

production in the duodenum, jejunum, and colon (Law et al., 2007).

Lastly, if nutrition support is not tolerated, mucin production can ultimately be hindered in

an infant with SBS. If a preterm infant is in a state of protein-energy malnutrition, the infant can

have decreased mucin production, resulting in increased susceptibility to bacterial translocation and

infection (Sherman et al., 1985).

MANAGEMENT OF INTESTINAL FAILURE

The main goals in nutritional and medical management of intestinal failure are to maintain

fluid, electrolyte and nutrient balances and to reduce side effects of treatment. Managing SBS in

infants is critical as an infant is undergoing rapid development and has high nutritional requirements.

Further, preterm infants have increased nutritional and medical needs because they have to catch up

in growth to their term counterpart. Nutritional supplementation is based on the etiology of

intestinal failure, the extent and location of resection, if resected, and the clinical lab values of the

18

patient to ascertain vitamin, mineral, and macronutrient deficiencies. Medical management often

includes drug therapies such as anti-motility agents to allow the intestine adequate time for digestion

and absorption. Intestinal transplantation or bowel lengthening procedures may be considered

when long term TPN complications have occurred (O'Keefe et al., 2006; Jan, 2008).

EN in patients with compromised intestinal function is preferred, if tolerated, to avoid the

complications associated with PN. If an infant is placed on PN, then management should include as

much EN as possible to prevent PN-associated atrophy, PN dependence, and the aforementioned

PN complications. Intestinal adaptation is the ability of the intestine to increase absorptive area and

to recover functional abilities of this area to adapt for the loss of intestine post-resection (Althausen

et al., 1950; Dowling and Booth, 1966; Bell et al., 1973; Stein and Wise, 1978; Curtis et al., 1984).

Unfortunately, PN administration nullifies intestinal adaptation post-resection due to the lack of

luminal stimulation provided by oral or enteral feeding (Morin et al., 1978; Ford et al., 1984; Ziegler

et al., 2002). Dahly and colleagues (2003) demonstrated after 70% midjejunoileal resection that rats

administered TPN had significantly greater mucosal hyperplasia by 8 hours, continuing through 7

days post-resection compared to transection controls. However, orally fed resected rats had an

enhanced adaptive response compared to the rats administered TPN. This study is an example of

intestinal adaptation occurring even with the administration of PN, but it should be noted that the

orally fed rats still experienced greater adaptation demonstrating the benefit of luminal stimulation

that EN or oral diets provide.

Threonine is an extremely important part of management in the SBS infant. The mucin core

is made of mostly serine, threonine, and proline residues, thus these amino acids are essential for

adequate mucin production. Studies with rats have been conducted to test mucin production with

reduced threonine intake which have resulted in steady production of muc2 mRNA but decreases in

mucin protein abundance, demonstrating that the effect of reduced threonine availability is at the

19

translational level of mucin production (Faure et al., 2005). Deficient threonine administration in

piglets results in decreased mucin accumulation in the duodenum and colon compared to the

adequate threonine fed counterparts (Law et al., 2007). In this study, adequate threonine resulted in

stable numbers of goblet cells; however, these goblet cells were hypertrophied—or filled with more

mucin—compared to deficient threonine fed piglets. In addition, adequate threonine goblet cells

had more heterozygous chemotypes or contained a greater mix of mucins (Law et al., 2007). These

studies suggest that adequate threonine supplementation plays a role in appropriate mucin

production.

Altogether, PN has many basic complications; it reduces barrier function, promotes SBBO,

decreases mucin abundance and ultimately increases the risk of bacterial translocation across the

intestinal mucosa. The interconnected nature of inflammation, mucins, SBBO and barrier function

in the process of bacterial translocation is evident, and as in all disease, prevention is the ideal

management. Provision of EN in tolerable amounts is the best management for SBS as it will help

to preserve mucosal integrity, barrier function, and mucin production, which together will reduce the

risk of inflammation and bacterial translocation.

MODULATION OF MUCIN PRODUCTION

Modulation via Stress, Nutritional Supplementation, and Growth Factors

As just discussed, nutrition support and threonine can both affect the amount and type of

mucin produced. Mucin can be affected by stress and other dietary factors as well. Mice exposed to

repeated cold stress initially had an increase in mucin glycoprotein content after one stress exposure;

however, goblet cell numbers were significantly decreased after just one stress exposure (Pfeiffer et

al., 2001). This suggests that goblet cells can increase the rate in which they produce mucin in

response to stress and become hypertrophied, or engorged, with mucin. Because cell development,

20

differentiation, and travel take a couple of days, hypertrophy is the most expected response to

increase mucin in acute time frames in situations of normal proliferation. Even though stress elicits

a beneficial response in mucin production, stress also causes a decrease in total goblet cell number.

This study highlights the importance of reducing stress on an infant with intestinal failure so that

goblet cell number is not reduced. In addition, more research needs to be conducted on how to

increase mucin production without inducing goblet cell-reducing stress. One of these methods is by

butyrate administration which will be discussed shortly.

Fiber can modulate mucin production. Rats that were administered TPN intravenously or

fed with the TPN solution orally showed significantly decreased epithelial bound mucin and mucin

within the goblet cells in the jejunum and the ileum. However, upon administration of 3.3 g

cellulose per day, mucin concentration levels tend to increase, demonstrating that certain fiber may

upregulate mucin production. Celluose, however, decreased unadhered luminal mucin abundance,

probably because cellulose administration decreased transit time, removing unbound luminal mucin

(Spaeth et al., 1994).

As discussed above, adequate provision of dietary threonine is crucial in intestinal mucin

synthesis. Increasing oral threonine intake to 150% of requirements significantly increases mucin

production in the jejunum (Faure et al., 2005). Another potential dietary modulator in mucin

production is β-casomorphin-7, a milk degradation peptide resulting from the digestion of β-casein,

which has been show to increase mucin production in vitro in rat and human goblet cells,

demonstrating the potential of dairy products to improve mucosal protective function (Zoghbi et al.,

2006).

Along with nutrition supplementation, growth factors—such as epidermal growth factor

(EGF)—are important for intestinal growth. EGF is important for cell growth, proliferation, and

differentiation (Cohen and Carpenter, 1975), and when provided luminally, decreases PN-associated

21

atrophy in rats administered TPN. In addition, EGF accelerates goblet cell maturation and mucin

production and normalizes expression of tight junction proteins, all of which improve intestinal

barrier function (Clark et al., 2006).

Modulation via the Short Chain Fatty Acid, Butyrate

Short chain fatty acids (SCFAs) are a subgroup of fatty acids including acetic, propionic,

isobutyric, butyric, isovaleric, valeric, and caproic acids. The salts or ester of these acids are acetate,

propionate, isobutyrate, butyrate, isovalerate, valerate, and caproate, respectively. The latter are

produced in vivo by fermentation of non-starch polysaccharides such as hemiceulloses, pectin, gums,

and mucilages by the anaerobic microflora of the cecum and colon. The main SCFAs produced are

acetate, propionate, and butyrate, which make up 83% of endogenous SCFAs. SCFAs in human

colon are generally found to be at a 60:25:15 molar ratio of acetate, propionate, and butyrate,

respectively (Cummings, 1984; Cummings and Englyst, 1987; Bourquin et al., 1992). Fermentative

end products, however, can be influenced by the type of fermentable substrate available (Mortensen

et al., 1988; Bourquin et al., 1992), the type of microbial species present in the intestine (Macfarlane

and Gibson, 1995), and by intestinal transit time (Macfarlane et al., 1992). The opposite is also true,

in that the type of fermentable substrate provided can influence the microbiota present (Wang and

Gibson, 1993). Thus, the type of fiber, a common source of SFCA production, chosen to

supplement EN or PN is important.

SCFAs can contribute about 10% of maintenance energy requirements in pigs (Imoto and

Namioka, 1978). The production is hypothesized to be similar in humans due to the similarity

between human and pig gastrointestinal tracts (Sangild, 2006). The energy contribution of SCFAs in

humans has been estimated at about 10-15% (McNeil, 1984). Butyrate is preferentially absorbed

22

over propionate and acetate at a ratio of 90:30:50, and approximately 70-90% of butyrate produced

in the intestines is used solely by the intestinal epithelium (Cummings, 1984).

Butryate has been shown in vitro to modulate muc2 expression (Augenlicht et al., 2003;

Willemsen et al., 2003; Gaudier et al., 2004; Hatayama et al., 2007; Burger-van Paassen et al., 2009).

Paradoxically, low levels (1-2 mM) of butyrate stimulate muc2 expression (Willemsen et al., 2003;

Hatayama et al., 2007; Burger-van Paassen et al., 2009), while higher levels of butyrate (5 mM-15

mM) either halt butyrate-stimulated increases of muc2 expression or even trigger mucous secretion

(Barcelo et al., 2000; Burger-van Paassen et al., 2009). Bathing the human colon cancer cell line,

LS174T, in 1-2 mM butyrate for 36 hours results in a 2-fold increase in muc2 mRNA and an 8-fold

increase in Muc2 protein (Hatayama et al., 2007). This was corroborated by Burger-van Passaan and

colleagues (2009) when 1 mM butyrate produced a 2-fold increase in muc2 mRNA levels when

administered to LS174T cells. Also in this study, 5-15 mM butyrate showed a return of muc2 mRNA

to basal levels. Additionally, 0.05 mM to 1 mM butyrate results in increased muc2 expression in

monolayers of T84 cells, and cocultures of T84 and LS174T cells with CCD-18Co cells (Willemsen

et al., 2003). In this study, butyrate was also shown to significantly increase prostaglandin E1

(PGE1) and decrease prostaglandin E2 (PGE2) concentrations in all dosage groups of cocultured

cells. Prostaglandins are lipid compounds that come from the eicosanoid family of paracrine

secretion (Saladin, 2004). They have been implicated in the regulation of mucous secretion

(Plaisancie et al., 1998) and have been shown to be released from myofibroblasts, or a cell similar to

both muscle and collagen, underneath the epithelial layer (Mahida et al., 1997). An increased

PGE1/PGE2 ratio has been thought to be beneficial because PGE1 is more potent than PGE2 at

stimulating muc2 expression in vitro (Willemsen et al., 2003).

Butyrate does not produce stimulatory effects in muc2 expression in all cell lines. muc2

expression is actually reduced upon butyrate bathing in Cl.16E cells (Augenlicht et al., 2003), a clonal

23

derivative of HT29 cells that are polarized and characterized by expression of the muc2 gene and

mucin synthesis and secretion (Augeron and Laboisse, 1984; Capon et al., 1992; Velcich et al., 1995).

Upon standard culturing of the Cl.16E cells, muc2 accumulated at day 7 and increased to a maximum

level at day 15. At this point, these cells were treated with 5 mM butyrate which resulted in a 50%

reduction in muc2 mRNA levels and a decrease in Muc2 apomucin after 24 hours (Augenlicht et al.,

2003). These results seem to indicate that butyrate has a negative effect on goblet cells; however, the

results can be explained by the butyrate concentration used and by referring to studies by others.

Augenlicht and colleagues consistently used butyrate at the concentration of 5 mM, which as

previously mentioned, has been shown to be inhibitory on muc2 expression (Burger-van Paassen et

al., 2009). Also, Gaudier and colleagues (2004) found that in a glucose rich medium, Cl.16E cells

show no increase in muc2 expression upon 2 mM butyrate treatment; however, in the absence of

glucose, muc2 expression increased 23-fold above basal level. Thus, the metabolism of butyrate as a

carbon source may play a role in the induction of Muc2 production. Augenlicht and colleagues

(2003) bathed their Cl.16E cells in Dulbecco’s Modified Eagle Medium (DMEM) which depending

on the solution, may contain glucose. Because the presence of glucose in butyrate administration

also effects muc2 expression (Gaudier et al., 2004), it is possible that the lack of effect of butyrate on

Muc2 levels found by Augenlicht and colleagues was due to the potential presence of glucose in the

cell medium.

Thus, while 1 and 2 mM butyrate levels increase muc2 expression, 5 mM butyrate

administration has been shown to reduce muc2 mRNA expression to baseline levels (Augenlicht et

al., 2003; Burger-van Paassen et al., 2009). In addition to muc2 expression and mucin production,

butyrate affects mucin secretion. Barcelo and colleagues (2000) demonstrated in vivo that 5 mM

luminal butyrate administration more than doubled mucin secretion in isolated vascularly perfused

rat colons. Mucins are secreted as protective measure in the intestine, and 5 mM butyrate appears to

24

trigger that protective response because that level is possibly perceived by the intestine as damaging

or butyrate may serve as a signal for regulated mucin secretion.

One ex vivo study analyzed the effects of butyrate administration on mucin production.

Butyrate administration of 0.1 mM resulted in a 476% mucin synthesis increase in comparison to the

control tissue in fresh human colonic biopsies. The increase was nullified in the presence of sodium

bromo-octanoate (an inhibitor of beta-oxidation), which reduces the ability for cells to metabolize

fatty acids such as butyrate, furthering the hypothesis that metabolism of butyrate by colonocytes is

an important mechanism in mucin synthesis (Finnie et al., 1995).

There have not been any in vivo studies conducted analyzing the effects of butyrate

administration on mucin production, nor have there been any studies analyzing parenteral butyrate

administration on mucin production, but butyrate has had strong positive effects on mucin

production when administered luminally in vitro and ex vivo. The one in vivo study by Barcelo and

colleagues did not ascertain the affects of luminal butyrate on mucin production or muc2 expression;

but studies in which fiber was administered in vivo (as previously discussed, fiber fermented by

enteric bacteria produce short chain fatty acids) mucin production was increased. Satchithanandam

and colleagues (1996) found that rats fed 10% psyllium had increased colonic luminal mucin, or

loosely bound mucin. They also suggest that total mucin levels—which include membrane adherent

mucins—were increased, but their determination methods of total mucin included protein assays of

the scrapings of the entire mucosal layer (Satchithanandam et al., 1996). Increases in protein content

in this case could be due to increases in villus height or other mucosal factors as butyrate positively

affects factors other than mucin when administered luminally in vivo (Scheppach et al., 1992;

Scheppach et al., 1997; Andoh et al., 1999; Vernia et al., 2000). Gaudier and colleagues have shown

that cells may need to metabolize butyrate to increase mucin production and muc2 expression;

however, since butyrate positively affects so many factors when administered luminally in vivo (in

25

26

which glucose availability cannot be eliminated), butyrate may still be able to produce positive effects

in the presence of glucose.

Administration of butyrate parenterally has been shown to enhance several indices of the

intestinal structure and function. Parenterally provided butyrate enchances glucose transporter-2 (glut-2)

mRNA, sodium glucose transporter-1(sglt-1) mRNA, Na+,K+-adenosine triphosphatase (Na+,K+ATPase)

mRNA (a critical basolateral enterocyte transporter), proglucagon mRNA (a precursor to GLP-2 and

incretins), plasma GLP-2 (an intestinotrophic peptide), glucose absorption, villus height,

proliferation and has been shown to decrease apoptosis (Tappenden et al., 1997; Tappenden and

McBurney, 1998; Bartholome et al., 2004). Because of the positive effects of butyrate seen in both

luminal and parenteral administration, parenteral butyrate and SCFA administration holds promise

as an intestinal mucin modulator as well, potentially providing protection from intestinal damage and

bacterial translocation. This is the first study to describe the effects of parenterally supplied butyrate

and SCFAs on muc2 mRNA expression and goblet cell chemotypes in a SBS model.

RESEARCH OBJECTIVE

Protective mucins are a crucial part of intestinal barrier defense. Mucins are even more

important in neonates with SBS on PN since both SBS and PN increase the risk for bacterial

translocation.

Several nutritional modulators of mucin production have been studied, with butyrate being

effective in upregulating mucin production in vitro and ex vivo; however the effect of butyrate in vivo is

unknown. This research aims to analyze the effect of parenteral butyrate administration on mucin-

producing goblet cell chemotypes in the small and large intestine using a PN-supported neonatal

piglet SBS model. The data from this research explores the following:

1) the effects of SCFA-supplemented TPN on goblet cell chemotypes and

muc2 mRNA in a massive small bowel resection neonatal piglet model;

2) the effects of butyrate-supplemented TPN, at doses of 9 mM and 60 mM, on

goblet cell chemotypesand muc2 mRNA in a massive small bowel resection neonatal

piglet model;

Because butyrate has been documented to improve so many factors of gastrointestinal

structure and function, we hypothesize that infusion of TPN supplemented with 9 mM butyrate will

enhance protective goblet cell chemotypes in the small intestine, specifically sulfomucins. Similarly,

we expect enhancement of protective goblet cell chemotypes in the SCFA-supplemented group too,

since this group also contains 9 mM butyrate. Further, we hypothesize that butyrate-supplemented

TPN will have a greater effect on the small intestine than the large intestine, while butyrate

supplementation at 60 mM will have a greater effect on the large intestine than the small intestine

because the large intestine is more accustomed to SCFA and may need a larger dose to confer a

benefit. As such, we expect muc2 mRNA abundance to increase in the small intestine with butyrate

27

28

and SCFA-supplemented TPN infusion, whereas we expect colonic muc2 mRNA abundance to

increase in the colon with 60 mM butyrate administration.

antigen

SED

Peyer’s Patch

= lymphoid follicle

= immature B lymphocyte

= macrophage

FAE

= mature B lymphocyte= mature B lymphocyte

= T cell

Afferent lymphatic

MLN

= dendritic cell

M cell

Figure 1.1: Peyer’s patch in the neonatal piglet. The M cell does not contain the basolateral pocket that is characteristic of mature M cells. The antigen is present at the luminal surface of the FAE, and upon M cell engulfment, a dendritic cell recognizes the antigen as a pathogen and transports the antigen to the lamina propria in or beyond the SED or to the nearest MLN via the afferent lymphatic system.

29

CHAPTER 2 THE EFFECT OF BUTYRATE-SUPPLEMENTED TOTAL PARENTERAL

NUTRITION ON muc2 mRNA AND GOBLET CELL CHEMOTYPES IN A SHORT BOWEL SYNDROME NEONATAL PIGLET MODEL

ABSTRACT

Background: Compromised barrier function and bacterial translocation are common

complications of short bowel syndrome (SBS) and parenteral nutrition (PN). Although mucins help

prevent intestinal bacteria translocation, total mucin production has been shown to decrease during

PN infusion. Butyrate administration may be a mucin fortifying strategy as it has been shown to

increase mucin production in vivo and ex vivo.

Objective: We tested the hypothesis that infusion of TPN supplemented with 9 mM butyrate will

enhance protective goblet cell chemotypes in the small intestine, specifically sulfomucins, that

enhancement of protective goblet cell chemotypes in the SCFA-supplemented group will be

observed, since this group also contains 9 mM butyrate, that butyrate-supplemented TPN will have a

greater effect on the small intestine than the large intestine, while butyrate supplementation at 60

mM will have a greater effect on the large intestine than the small intestine. We also hypothesize

that muc2 mRNA abundance will increase in the small intestine with butyrate and SCFA-

supplemented TPN infusion, whereas colonic muc2 mRNA abundance will increase in the colon

with 60 mM butyrate administration.

Methods: Forty-eight hour old piglets underwent 80% proximal jejunuoileal resection and were

randomized to one of four groups: control total parenteral nutrition (TPN), TPN supplemented

with 60 mM SCFA (36 mM acetate, 15 mM propionate, and 9 mM butyrate; SCFA), 9 mM butyrate

(9Bu) or 60 mM butyrate (60Bu). Animals were further randomized to an acute (12 hour) or chronic

(72 hour) time point. muc2 mRNA abundance, total goblet cell number, and total sulfomucin,

sialomucin, acidomucin, and neutral mucin goblet cell numbers were ascertained.

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Results: Sulfomucin chemotypes were increased in the jejunal villi of the 9Bu group compared to

control (treatment main effect, p=0.047). Acidomucin chemotypes in ileal crypts were greater in the

60Bu group than the control group (treatment main effect, p=0.038). In the colonic crypts, SCFA

groups tended to have greater acidomucin chemotypes than control at 12h while 60Bu group tended

to have greater acidomucin chemotypes per depth than control at 72h (p=0.060; Table 2.3). muc2

mRNA was increased in jejunal tissues in the 9Bu group compared to control with 270% and 30%

increases in muc2 mRNA at 12 and 72 hours, respectively (p=0.010).

Conclusion: The 9 mM Bu treatment increased protective goblet cell chemotypes and muc2 mRNA

in the jejunum while 60 mM Bu increased protective goblet cell chemotypes in the ileum and colon.

The 9Bu treatment was the most effective at upregulating muc2 mRNA abundance and sulfomucin

chemotypes. Butyrate-supplemented PN may be a therapeutic strategy for bolstering the epithelial

barrier by increasing mucin production in neonates with SBS on PN.

INTRODUCTION

Within the first line of defense against invading pathogens—innate immunity—is intestinal

mucous, the majority of which is secretory gel-forming mucins, predominantly Muc2, a mucous

protein encoded by the muc2 gene (Gum et al., 1989; Griffiths et al., 1990). Mucins are classified as

acidomucins or neutral mucins. Acidomucins have either sulfate or sialic acid residues on their

carbohydrate moieties and are called sulfomucins and sialomucins, respectively, whereas neutral

mucins have no such residues (Hansson et al., 1991; Carlstedt et al., 1993). Sulfomucins and neutral

mucins confer a more protective effect in the intestine than sialomucins because they are more

slowly degraded by bacteria, therefore the established continuum of mucin protection is as follows:

sulfomucins > neutral mucins >>> sialomucins (Fontaine et al., 1996).

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Several studies confirm the protective nature of sulfomucins (Sakata and von Engelhardt,

1981; Deplancke et al., 2000; Conour et al., 2002; Dawson et al., 2009). It has been demonstrated

that sulfomucins increase in concentration from the duodenum to the colon, mirroring the pattern

of bacterial concentration from proximal to distal bowel (Sakata and von Engelhardt, 1981;

Deplancke and Gaskins, 2001; Conour et al., 2002). Sulfated mucins have been shown to be

essential in protecting against bacterial translocation (Dawson et al., 2009).

Muc2 is crucial in preventing bacterial contact with the epithelial lining (Johansson et al.,

2008). Decreases in mucin abundance or in protective goblet cell chemotypes leave the epithelial

barrier exposed to bacteria and bacterial products, which can lead to bacterial translocation (Gork et

al., 1999). Direct contact of harmful luminal contents to the epithelial lining promotes

inflammation, which can further impair barrier function (Yang et al., 2002; Yang et al., 2008) and

increase the risk of bacterial translocation. Bolstering mucin quality and abundance may improve

the outcome of SBS patients by preventing bacteria-induced mucosal inflammation and further

complications such as bacterial translocation.

Mucin abundance is decreased by PN administration, which is the primary nutritional

therapy for individuals with intestinal failure (Iiboshi et al., 1994; Bertolo et al., 1998; Law et al.,

2007). The SCFA butyrate enhances mucin production in vitro and ex vivo (Finnie et al., 1995;

Willemsen et al., 2003; Hatayama et al., 2007; Burger-van Paassen et al., 2009), but its direct effect

on mucin chemotype and abundance in vivo and systemically administered as a component of PN has

not been elucidated. This study analyzes the effect of parenteral butyrate and SCFA administration

on both muc2 mRNA abundance and goblet cell chemotypes in a TPN-supported neonatal piglet

with SBS. We hypothesized that infusion of TPN supplemented with 9 mM butyrate would

enhance protective goblet cell chemotypes in the small intestine, specifically sulfomucins. Similarly,

we expected enhancement of protective goblet cell chemotypes in the SCFA-supplemented group

32

too, since this group also contained 9 mM butyrate. Further, we hypothesized that butyrate-

supplemented TPN would have a greater effect on the small intestine than the large intestine, while

butyrate supplementation at 60 mM would have a greater effect on the large intestine than the small

intestine because the large intestine is more accustomed to SCFA and may need a larger dose to

confer a benefit. As such, we expected muc2 mRNA abundance to increase in the small intestine

with butyrate and SCFA-supplemented TPN infusion, whereas we expected colonic muc2 mRNA

abundance to increase in the colon with 60 mM butyrate administration.

MATERIALS AND METHODS

Experimental Design

Piglets were obtained at 48 hours of age from the Imported Swine Research Laboratory at

the University of Illinois at Urbana-Champaign. Piglets underwent superior vena cava cannulation,

swivel placement and 80% proximal jejunuoileal resection and were then randomized by initial body

weight (1.77 ± 0.16 kg, n = 120) to one of four groups: control TPN (TPN), TPN supplemented

with 60 mM SCFA (36 mM acetate, 15 mM propionate, and 9 mM butyrate; SCFA), 9 mM butyrate

(9Bu) or 60 mM butyrate (60Bu). Animals were futher randomized to an acute 12 hour (12h) or a

chronic 72 hour (72h) infusion time point, at which point samples were harvested and dependent

variables assessed.

Surgery

The surgical protocol was approved by the Laboratory Animal Care Advisory Committee at

the University of Illinois at Urbana-Champaign. Aseptic technique and sterile equipment were used

at all times. Piglets were anesthetized with isoflurane (98% oxygen, 2% Isoflurane; Baxter

Pharmaceutical Products, Inc., Deerfield, Illinois). Betadine (10% povidone-iodine; Purdue

Frederick Co., Norwalk, Connecticut) and lidocaine (2% lidocaine HCl, 20mg/ml; Abbott

33

Laboratories, North Chicago, Illinois) were applied to the surgical sites prior to jugular

catheterization, laporotomy, and catheter externalization. A 3.5 French polyvinyl chloride catheter

(Bergen-Brunswig, Lack Zurick, Illinois) was inserted into the external jugular vein through a 3-cm

incision in the right clavicle region. The catheter was inserted 6-cm into the external jugular into the

superior vena cava for TPN infusion. After placement, the catheter was flushed with 5 mL of

heparinized saline (20 U/ml; Sigma-Aldrich Co., St. Louis, Missouri) and subcutaneously tunneled

and externalized in the subscapular region.

All piglets also received an 80% proximal jejunoileal resection, as follows. A laparotomy was

performed to the right of the umbilicus and 15-cm of jejunum distal to the ligament of Treitz and

75-cm of ileum proximal to the ileoceccal junction were measured. Following vessel stricture, the

measured small intestine was excised, weighed and measured, and an anastomosis performed to

reestablish bowel continuity. The peritoneum, muscularis, and subcutaneous layers were sutured

separately. Piglets’ activity levels were monitored during surgical recovery. Piglets received Naxcel®

antibiotic (3 mg/kg BW; SmithKline Beeham Corp., Philadelphia, Pennsylvania) prior to surgery and

during the entire study period. Piglets also received buprenorphine analgesic (.01 mg/kg BW; Henry

Schein Inc., Melville, New York) intramuscularly every 12 hours for the first 24 hours post-surgery

to minimize discomfort.

Animal Care and Housing

Piglets were fitted with jackets and swivel tethers (Alice King Chatham Medical Arts,

Hawthorne, California) to protect infusion lines while allowing the piglets to move freely in their

cages. Piglets were individually housed in metabolic cages in the Edward R. Madigan laboratory

animal care facility at the University of Illinois at Urbana-Champaign. Radiant heaters were placed

at the top of the cages to maintain temperature of about 34°C.

34

Parenteral Nutrition Solutions

PN solutions were prepared and sterilized daily in a laminar flow hood. The solutions

contained dextrose, amino acids, lipids, and all essential minerals and vitamins (Table 2.1). SCFA

and butyrate were added to the PN solutions at the concentrations previously mentioned.

Volumetric infusion pumps (IMED 980; St. Louis, Missouri) were used to infuse PN. A stress

factor or 20% was included in calculating energy and protein needs resulting in an infusion of 307

Kcal/Kg/d and 15.5 g amino acids/kg/d. The piglets received nutrition support continuously

throughout the duration of the study. Nutrition needs were calculated daily and PN infusions rates

adjusted accordingly to ensure adequate supply of energy and nutrients for growth and recovery

from surgery.

Sample Collection

At euthanasia, the intestines were promptly removed. Proximal to the ligament of Treitz

was considered the duodenum. Distal from this ligament to the anastomosis was considered the

jejunum, and distal to the anastomosis and proximal to the ileoceccal valve was considered the

ileum. The colon was removed as well. All segments were flushed with ice-cold saline and samples

from each segment were fixed in 10% buffered formalin. Formalin-fixed tissue was dehydrated in

ethanol and infiltrated and embedded in paraffin wax. Blocks of tissue were sectioned to

approximately 5 µm thickness at the Veterinary Medicine Center for Microscopic Imaging and

sections were placed on polylysine slides for optimal adhesion during staining procedures.

Measurement of muc2 RNA

Total cellular RNA was isolated from jejunal, ileal, and colonic samples using the guanidium

isothiocyanate phenolchloroform method of Chomczynski (1993) and as previously described in our

35

lab (Bartholome et al., 2004). RNA concentration and purity after isolation were determined using a

Thermo Scientific NanoDrop 2000 (Thermo Scientific, Wilmington, DE). RNA quality was

determined by running all samples through ethidium bromide RNA gel electrophoresis and

examining under UV illumination using a Fotodyne Incorporated 3-3100 UV light box (Fotodyne

Incorporated, Hartland, WI). In addition, 12 samples with varying 18S and 28S band intensities in

gel electrophorosis were chosen for analysis in an Agilent 2100 bioanalyzer (Agilent Technologies,

Santa Clara, CA) at the W.M. Keck Center for Comparative and Functional Genomics (University of

Illinois at Urbana-Champaign, Urbana, IL) to further ascertain RNA integrity. The sample with the

weakest 18S and 28S band under UV illumination after RNA gel electrophoresis produced a RNA

integrity number (RIN) of 7.30 in the Agilent 2100 bioanalyzer. An RIN number is assigned by

Agilent Technologies’ bioanlyzer software and is a representation of the entire electrophoretic trace

of an RNA sample including degradation products. An RIN of greater than 7 is recommended for

optimal rt-PCR (real time-polymerase chain reaction) results. Thus, all samples were of high quality

and ideal for rtPCR.

Reverse transcription was carried out in a Gene Mate® Genius thermocycler (ISC

Biosexpress, Kaysville, Utah) as a 20 µL total volume containing 5µg RNA reconstituted in 10 µL

DEPC water, 1 µL random primer, 1 µL dNTP, 4 µL 5x First-Strand Buffer, 2 µL 0.1 M DTT, 1 µL

RNaseOut, and 1 µL SuperScript II Reverse Transcriptase. All of these reagents were obtained

from Invitrogen Corporation. rt-PCR consisted of a total reaction volume of 10 µL containing 2 µL

DEPC water, 2 µL cDNA template, 5 µL of TaqMan® Universal PCR Master Mix, 0.5 µL

eukaryotic 18S rRNA endogenous control primer and probe 20x (Applied Biosystems, Carlsbad,

CA), .5 µL Muc2 Custom Made Taqman® Gene Expression Assays PCR primer and probe

(Applied Biosystems, Foster City, CA). rt-PCR was completed using a Taqman ABI 7900 real time

PCR machine (Applied Biosystems, Carlsbad, CA).

36

The oligonucleotide primers specific for Muc2 cDNA detection were made by Applied

Biosystems Custom Made Taqman® Gene Expression Assays. The custom probe was designed

using the National Center for Biotechnology Information (NCBI) nucleotide and protein databases

and Primer Express 2.0 (Applied Biosystems, Carlsbad, CA). Because pig DNA for muc2 has not

yet been elucidated, a search was completed for mouse muc2 mRNA through the nucleotide

database in NCBI. Gi accession number 149258274 was chosen because it did not contain long

segments of repeated codons. Because mucins commonly have similar protein backbones, especially

in the center of the molecule, choosing a gene sequence with long segments of repeated codons

would decrease specificity of the designed primer. A nucleotide blast was then completed

comparing this chosen gene against sus scrofa expressed sequence tags (ESTs) with a low complexity

filter to mask repeat regions. An 81% identity was found with EST: BX671371 which was then

blasted against refseq RNA to establish if this EST matches muc2 mRNA already elucidated for

other species. A significant alignment of 88% identity was found with NM 002457 or Homo sapiens

muc2 mRNA. Then a blastx—protein comparison—of EST BX671371 against non-redundant

protein sequences was performed resulting in 25 matches of significant alignment with elucidated

muc2 proteins. The first 8 matches were 85%-87% similar to EST BX671371, with the 15%

difference likely due to species differences in mucin proteins. Accession numbers CAD54416

(Muc2 protein Mus muscularis-922 amino acids) and AAA59861.1 (Muc2 protein Homo sapiens-131

amino acids) were blasted against each other with only an 82% alignment, demonstrating the

interspecies variation of mucin proteins. BX671371 was entered into Primer Express 2.0 software,

with temperature, cytosine and guanine content, and primer length at default settings, and the primer

set with lowest penalty was chosen for custom primer and probe design. The resulting primer set

requested from Applied Biosystems is as follows:

Forward: 5’ – TCCAAGCCTCTCCCTGCAT – 3’

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Reverse: 5’ – ACAGCCACCAGGTCTCATTCTC – 3’

Probe: 5’ – CAGACTTCGATCCTCCCA – 3’

The rt-PCR product of pooled cDNA of all samples were run on a 1.8% agarose gel with a

50 base pair DNA ladder (New England Biolabs, Ipswich, MA) and visualized under UV

illumination on a Molecular Imager® ChemiDoc™ XRS with Quantity One 4.6.3 software (Bio-Rad

Laboratories, Hercules, CA) to determine the length of the amplicon to verify amplification of the

target. One band was present for 18S mRNA while no band was seen for muc2 (Figure 2.1).

Measurement of Mucin Protein: Acidic versus Neutral Mucins

Samples used for goblet cell chemotype counting were formalin-fixed, sliced to 5 µm

thickness and placed on polylysine slides. Samples were stained with periodic acid alcian blue

(PASAB) carbohydrate stain technique (Mowry, 1956; Carson, 1990). Sections were deparaffinized,

stained for 5 minutes with alcian blue, rinsed for 5 minutes in running tap water and 5 minutes in

ddH20. They were then incubated for 10 minutes in 1.0% periodic acid, and rinsed again for 5

minutes in running tap water and ddH20, sequentially. A 10-minute incubation in Schiff’s reagent

was conducted, followed by another sequential rinse of tap water, then ddH20. Lastly, slides were

counterstained with Harris’s hematoxylin with acetic acid for 15 seconds, rinsed in running tap water

for 10 minutes, dehydrated, and cover-slipped. PASAB stain colors neutral mucins pink and acidic

mucins blue (Figure 2.2). Another scientist blinded the histologist to the sample origin. Up to 10

well-oriented and intact villi and crypt units were identified in each sample, length and depth

measured, and goblet cells counted. Goblet cells were counted, then categorized to one of the

following 5 groups: 1.) 100% acidomucin; 2.) 75% acidomucin and 25% neutral mucin; 3.) 50%

acidomucin and 50% neutral mucin; 4.) 25% acidomucin and 75% neutral mucin, and; 5.) 100%

neutral mucin. The mixed goblet cells (Figure 2.3) were partitioned into the group that most

38

closely matched its composition. In addition, to obtain solely acidomucin and neutral mucin values

of all goblet cells, calculations of the mixed goblet cells—those containing both acidomucin and

neutral mucin components—were completed to distribute acidomucin and neutral mucin

components into either the 100% acidomucin chemotype or 100% neutral mucin chemotype

groups. Images were obtained using a Zeiss Axioskop 40 microscope and a Zeiss AxioCam MRc 5

camera (Carl Zeiss IMT Corporation, Maple Grove, MN). Length and depth measurements of

crypts and villi were obtained using AxiovisionLE Software (Release 4.7.2, Carl Zeiss). Images for

publication were obtained using an Olympus BX51 microscope, an Olympus DP70 digital camera

and DP Manager software (Olympus America Inc., Center Valley, PA).

Measurement of Sulfomucin versus Sialomucin Goblet Cell Chemotypes

Samples used for goblet cell chemotype counting were formalin-fixed, sliced to 5 µm

thickness and placed on polylysine slides. Samples were stained with high iron diamine alcian blue

(HIDAB) carbohydrate stain technique, as previously described (Spicer, 1965; Spicer et al., 1965;

Carson, 1990), except incubation in high iron diamine solution was limited to 16 hours rather than

18 hours to reduce non-specific staining. Following high iron diamine stain, slides were washed in

running tap water for 5 minutes and stained with alcian blue (pH 2.5) for 5 minutes, washed in

running tap water for 2-3 minutes, counterstained with Nuclear Fast Red (Sigma Aldrich, St. Louis,

MO) for 1-2 minutes, washed in running tap water for 5 minutes, dehydrated, and cover-slipped.

HIDAB stain colors sulfated mucins brown and sialated mucins blue (Figure 2.4). Another

scientist blinded the histologist to the sample origin. Up to 10 well-oriented and intact villi and

crypt units were identified in each sample, length and depth measured, and goblet cells counted.

Goblet cells were counted in one of the 5 following groups: 1.) 100% sulfomucin; 2.) 75%

sulfomucin and 25% sialomucin; 3.) 50% sulfomucin and 50% sialomucin; 4.) 25% sulfomucin and

39

75% sialomucin; and 5.) 100% sialomucin. These mixed goblet cells (Figure 2.5) were partitioned

into the group that most closely matched its composition. Calculations were carried out and imaged

captured as described in the previous section.

Statistical Analysis

Data were analyzed using the mixed model in SAS statistical software (version 9.1; SAS

Institute, Cary, NC). Analysis was completed using analysis of variance (ANOVA) for a split-plot

design. Sources of variation include litter (block), time (main plot), and treatment (subplot).

Residuals were output from the mixed model and the Shapiro-Wilk test was used to test for

normality. Spearman correlation tests were performed on the residuals to test for homogeneity.

Log or reciprocal transformations were performed for any data that was not normal or had unequal

distribution of residuals. After p-values were obtained, data was retransformed to acquire the

correct mean estimate and the standard error of retransformed data is presented as an average of

standard error (SE) values calculated from both the upper and lower retransformed confidence

intervals of group means. If the main effects for time or treatment, or the interaction of

time*treatment were significant, the following post hoc comparisons were conducted: control versus

9Bu at 12h, control versus 60Bu at 12h, control versus SCFA at 12h, control versus 9Bu at 72h,

control versus 60Bu at 72h, and control versus SCFA at 72h. These comparisons were completed

for rt-PCR results, total goblet cell number, sulfomucin versus sialomucin chemotype abundance,

and acidomucins versus neutral mucins chemotype abundance. Significant data are identified as p≤

0.050 and trends identified by p≤ 0.100. Data were analyzed as one-tailed t-tests given the

directional hypothesis being tested.

40

RESULTS

Body and Organ Weights

As previously reported by our research group, piglet body weight did not differ between

control piglets and those receiving SCFA and butyrate supplementation at any point during the

study (Bartholome et al., 2004). In addition, organ weights did not differ between groups at the

conclusion of the study (Bartholome et al., 2004).

Total Goblet Cell Number

In the jejunal villi, 12h total goblet cell number per height tended to be greater than 72h total

goblet cell number (time main effect, p=0.074; Table 2.2). In the ileal crypts, 12h goblet cell

number per depth was significantly greater than that at 72h (time main effect, p=0.0004; Table 2.2).

No significance was noted in the colon (Table 2.2).

Acidomucin and Neutral Mucin Goblet Cell Chemotypes

Within the jejunal and ileal crypts, 12h neutral chemotypes per crypt depth (in µm) were

significantly greater than at 72h (p=0.005, p=0.012, respectively; Table 2.3). Within the ileal crypt,

12h acidomucin chemotypes per depth were also greater at 12h than at 72h (p=0.0003). Also within

the ileal crypt, acidomucin chemotypes per depth were greater in the 60Bu group than the control

group (treatment main effect, p=0.038; Figure 2.6). In the colonic crypts, SCFA groups tended to

have greater acidomucin chemotypes per depth than control at 12h while 60Bu group tended to

have greater acidomucin chemotypes per depth than control at 72h (p=0.060; Table 2.3).

Within the ileal villi, SCFA and 60Bu acidomucin chemotypes per villus height (in µm) were

greater than control at 12h; however, these differences were not sustained at 72h (p=0.031; Table

41

2.4; Figure 2.7). No significant differences or trends were noted in the jejunum for either neutral

mucin chemotypes or acidomucin chemotypes per height (Table 2.4).

Sulfomucin and Sialomucin Goblet Cell Chemotypes

Within the ileal crypts, sialomucin chemotypes per depth tended to be greater at 12h than at

72h (time main effect, p=0.054; Table 2.5). Within the colonic crypts, control sialomucin

chemotypes per depth were greater than 9Bu, 60Bu, and SCFA at 72h (p=0.039; Table 2.5; Figure

2.8). No differences were noted in the jejunal crypts.

Within the jejunal villi, 9Bu had greater sulfomucin chemotypes per height than control

(treatment main effect, p=0.047; Table 2.6; Figure 2.9). Also within the jejunal villi, sialomucin

chemotypes per height tended to be greater at 72h than at 12h (p=0.058; Table 2.6). Within the

ileal villi, control groups had greater sialomucin chemotypes than SCFA groups at 12h; however, this

difference was not seen at 72h (p=0.0004; Table 2.6).

Muc2 mRNA Abundance Within the jejunum, muc2 mRNA was greater in the 9Bu group than the control group

(treatment main effect, p=0.010; Table 2.7; Figure 2.10). Within the ileum, muc2 mRNA was

greater at 72h than at 12h (p=0.017; Table 2.7). No significance was noted in the colon.

DISCUSSION

The goal of this study was to investigate the effect of butyrate and SCFA administration via

parenteral nutrition on mucin production in the jejunum, ileum, and colon. Our study revealed

significant increases at the transcriptional level of mucin production and significant increases in

protective goblet cell chemotypes upon butyrate supplementation. This study used a parenterally-

42

supported SBS neonatal piglet model that is recognized as an ideal model for gastrointestinal disease

in the human infant for two reasons. First, the timing of gastrointestinal maturation is quicker than

humans, but clusters in a similar ratio, meaning the pig is a close reflection of human maturation

sequences but matures in much less time, naturally lending itself to the research setting (Sangild,

2006). Further, piglets have shown similar intestinal adaptation to humans post-resection

(Heemskerk et al., 1999). Thus, due to the interspecies similarities in the gastrointestinal tract, the

piglet’s mucin responses to resection, PN, and butyrate administration will likely be similar to

humans’ response.

In this study, the number of acidomucin goblet cell chemotypes per crypt were nearly double

that of the neutral mucin chemotypes in the colon of the control piglets which mirrors the natural

state of mucin ratios seen in neonatal life in humans (Filipe et al., 1989; Deplancke and Gaskins,

2001), further demonstrating the validity of the piglet model for study of goblet cell development

and chemotypes.

Gel electrophoresis of pooled cDNA samples produced a faint 18S band; however no muc2

band was present. The 18S band was estimated to be 18 ng while the lowest amount present in the

DNA ladder was approximately 27 ng. Even though 40 uL of rt-PCR amplicon was injected into

the sample well, the 18S band was still faint. rt-PCR demonstrated by CT counts that 18S was

exponentially greater in quantity than muc2. Thus, we believe that muc2 is at a concentration that is

high enough to be detected by rt-PCR, but not high enough to be visible on a gel. However, rt-PCR

verified amplification of muc2. Additionally, Taqman primer and probe sets are highly sensitive and

specific, and NCBI blasts confirmed that the primer and probe sets have significant alignment with

Sus scrofa ESTs and muc2 mRNA of other species demonstrating that the primer and probe sets

would amplify Sus scrofa muc2.

43

Sulfomucin goblet cell chemotypes increased within the jejunal villi of the 9Bu group. As

previously mentioned, sulfomucins are the most protective against bacterial degradation, with 40%

degraded at 4 hours after bacterial inoculation, followed by neutral mucins with 55% degraded at 4

hours post-inoculation, and followed lastly by sialomucins which are 90% degraded at 1 hour and

almost completely degraded at 4 hours (Fontaine et al., 1998). With a 37.3% increase of villus

sulfomucin chemotypes at 12h and a 10.7% increase at 72h, the 9Bu group likely has an improved

protective barrier against bacteria than the control group. The increase in sulfomucin chemotype in

the 9Bu group was mirrored by significant jejunal muc2 mRNA increases in the 9Bu group. A 270%

increase in muc2 mRNA was noted at 12h and a 30% increase was observed at 72h. Enhancement

by 9Bu at the transcriptional level and enhancement of protective goblet cell chemotypes were

greater at the acute time point (12h) than the chronic time point (72h), suggesting that butyrate

administration may be more beneficial for the jejunum immediately after small bowel resection.

Because 9Bu had positive effects but SCFA did not (SCFA is 60mM total concentration SCFA with

9mM butyrate), 60mM total concentration may be inhibitory in the jejunum. This will be discussed

in more depth shortly.

The distal small bowel did not show any significant improvements versus control with 9Bu

treatment. In the ileum, 60Bu significantly enhanced crypt acidomucin chemotypes compared to

control (treatment main effect). There was a 16.4% increase at 12h and a 32.6% increase at 72h. In

addition, 60Bu and SCFA enhanced villus acidomucin chemotypes at 12h by 75.5% and 44.7%,

respectively. These data indicate that butyrate supplementation of 60Bu is most beneficial for the

ileum and that benefits are seen at both acute and chronic time points. They also indicate that since

60Bu and SCFA—both 60mM treatments, but differing butyrate concentrations—had positive

effects, that concentration of short chain fatty acids, rather than type of short chain fatty acid may

be the mechanism by which the mucin is enhanced in the ileum. Conversely, since 60Bu produced a

44

greater increase in acidomucin chemotypes than the SCFA group, one could also argue that butyrate

was the source of mucin enhancement, and that the greater concentration of butyrate may have

increased the benefit.

Colonic goblet cell abundance and chemotype was not altered by the 9Bu treatment.

However, SCFA and 60Bu tended to increase acidomucin chemotypes at 12h and at 72h,

respectively. This again suggests that the distal bowel may need greater butyrate concentrations than

the jejunum to confer benefits.

There are a couple possibilities for why the jejunum may have responded positively to 9Bu

while the ileum and colon responded positively to 60Bu. The 9mM dosage of butyrate administered

to the piglets averaged 2.21mM/hour for the average piglet weight, while the 60mM butyrate dosage

averaged 14.9mM/hour. As discussed previously, 1-2 mM of butyrate has been shown to be

beneficial upon luminal administration, while 5-15 mM has been shown to be detrimental (Barcelo et

al., 2000; Willemsen et al., 2003; Hatayama et al., 2007; Burger-van Paassen et al., 2009). No studies

have been conducted on an upper limit of butyrate administered parenterally; however, it seems

there may be an upper limit of butyrate tolerance for the jejunum, at least. Because bacterial

concentrations increase from proximal to distal bowel, the ileum is accustomed to greater SCFA

concentration than the jejunum. Therefore, it is conceivable that a higher concentration of butyrate

is needed—even when administered parenterally—to stimulate beneficial effects in the ileum versus

the jejunum. Furthermore, because the jejunum is exposed to less fermentative products such as

butyrate, 60mM butyrate may be beyond the tolerable level of short chain fatty acids for the

jejunum. As previously shown, plasma concentrations of SCFA are not elevated in this piglet model

with administration of 60 mM SCFA, 9 mM butyrate, and 60 mM butyrate (Bartholome et al., 2004),

indicating that either the upper limit was not reached with these dosages, or an upper limit cannot be

defined by remaining plasma concentrations. Rather, upper limits may need to be defined by

45

measured structural and functional indices of the intestine such as protective goblet cell chemotype

abundance.

Sialomucins have been shown to be less protective than sulfomucins and neutral mucins

(Fontaine et al., 1998). In this study, control groups had greater sialomucin chemotypes than the

SCFA group at 12h in the ileum and control groups in the colon had greater sialomucin chemotypes

than all treatment groups at 72h. Sialomucin chemotypes only comprise 2% of the total colonic

goblet cells in these tissues, so although the decrease in sialomucin chemotypes was seen, it was a

negligible decrease as sialomucins are such a minor component of large intestinal mucin. Further,

humans have an even greater ratio of sulfomucins/sialomucins than piglets (Deplancke and Gaskins,

2001). Due to sialomucins not being as protective as other mucins, sialomucin chemotypes

composing a small percentage of total colonic goblet cells, and the greater sulfomucin/sialomucin

ratio in humans, decreases in sialomucins by SCFA and butyrate administration are statistically but

not clinically significant. Although neither a decrease nor an increase in sialomucins is clinically

significant, additional mucins no matter how easily degraded could only be beneficial as long as there

is no concomitant decrease in sulfomucins or neutral mucins. A significant increase in sialomucin

chemotypes was noted in the jejunal villus with 9Bu, which could only be a beneficial adaptation.

Significant time differences were noted in that jejunal crypt neutral mucin chemotypes and

ileal crypt neutral mucin, acidomucin and sialomucin chemotypes were greater at 12h than at 72h

(time main effect). Because there were no treatment differences, the apparent decrease in goblet cell

chemotypes from 12-72h were not due to SCFA or butyrate administration. Rather, there was a

trend for a decrease in jejunal crypt total goblet cells and a significant decrease in ileal crypt total

goblet cells, showing that this is likely a natural physiological change seen in piglets recovering from

massive bowel resection. The decrease of total goblet cell number at 72h can be explained by the

effect parenteral nutrition has on mucin production and other indices of intestinal functioning

46

previously mentioned. It may be that the upregulation in mucins provided by butyrate treatment or

adaptation cannot outweigh the increasingly damaging effects that PN administration has on

mucosal function and morphology over time. As we have previously reported, this piglet model

experiences an increased rate of crypt cell proliferation with SCFA and butyrate administration

compared to control piglets; however, all piglets have a decreased rate of crypt cell proliferation at

72h compared to 12h demonstrating that administration of SCFA and butyrate may not be able to

overcome the negative effects of PN administration over time (Bartholome et al., 2004).

The regulation of mucin expression has not been established in this study or in other

literature. In this study, ileal muc2 was greater at 72h than at 12h, but ileal crypt goblet cells were

greater at 12h than at 72h with no significant differences noted in the ileal villi. This suggests that

goblet cell chemotype development is regulated at the translational level. However, jejunal

regulation of mucin expression appears to be transcriptionally regulated. muc2 mRNA abundance in

the jejunum was increased by 9Bu treatment with a concomitant increase in jejunal sulfomucin and

sialomucin chemotypes, indicating that an increase in mucin gene transcription may lead to an

increase in mucin protein translation. Further study needs to be conducted to clarify the

mechanism(s) of mucin and goblet cell chemotype regulation and how these methods may vary

along the length of the intestinal tract.

In summary, with adequate nutritional provision, upregulations of protective goblet cell

chemotypes and muc2 expression occurred with butyrate and SCFA administration. The 9Bu

treatment produced the best improvement of the jejunal mucin profile, while SCFA and 60Bu

treatments improved the ileal mucin profile. Because of the discrepancy in benefit between

treatments, the ideal treatment may depend on the etiology of intestinal failure and resulting bowel

resection. A larger remaining jejunum than ileum post-resection may have greater benefit from the

47

48

9Bu treatment while a greater remaining ileum versus jejunum post-resection may receive greater

benefit from the SCFA or 60Bu treatments.

In conclusion, SCFA-supplemented TPN enhances protective goblet cell chemotypes in the

neonatal intestine of a SBS model piglet, and butyrate-supplemented TPN enhances both muc2

mRNA and protective goblet cell chemotypes in the neonatal intestine of a SBS model piglet.

Therefore, supplementation of TPN with SCFA or butyrate may benefit infants with SBS by

increasing protective goblet cell chemotypes and muc2 expression, potentially decreasing PN-

associated complications such as bacterial translocation.

Table 2.1: Composition of parenteral nutrient solutions Ingredients TPN1

TPN + 9Bu2 TPN + 60Bu2 TPN + SCFA2

Dextrose3,4 (ml/L) 235.3 231.9 216.2 224.2

Amino acids4,5 (ml/L) 594.0 594.0 594.0 594.0

Lipids4,6 (ml/L) 133.3 133.3 133.3 133.3

Acetic acid7 (mmol/L) __ __ __ 36.0

Propionic acid7 (mmol/L) __ __ __ 15.0

n-Butyric acid7 (mmol/L) __ 9.0 60.0 9.0

MTE-68,9 (µl/L) 100.0 100.0 100.0 100.0

Iron dextran10 (µl/L) 111.0 111.0 111.0 111.0

Calcium gluconate9,11 (µl/L) 2.2 2.2 2.2 2.2

Vitamin A + E + D12 (µl/L) 23.5 23.5 23.5 23.5

Vitamin K13 (µl/L) 20.0 20.0 20.0 20.0

Vitamin C14,15 (µl/L) 320.0 320.0 320.0 320.0

Vitamin B complex14,16 (µl/L)

200.0 200.0 200.0 200.0

Folic acid17 (µl/L) 28.0 28.0 28.0 28.0

Biotin7,18 (µl/L) 20.0 20.0 20.0 20.0

Sterile water (ml/L) 34.4 37.0 48.2 41.4

1TPN represents a standard total parenteral nutrient solution providing; energy, 1 Kcal/ml; carbohydrate, 100mg/ml; amino acids 50.5 mg/ml; lipid 40 mg/ml. 2TPN + 9Bu, TPN + 60Bu, and TPN + SCFA represent standard total parenteral nutrient solution plus the short-chain fatty acids listed. 350% dextrose (50 g dextrose/100 ml). 4Baxter Healthcare Corporation, Deerfield, Illinois. 58.5% Travasol®; (leucine 529 mg, phenylalanine 526 mg, lysine 492 mg, methionine 492 mg, isoleucine 406 mg, valine 390 mg, histidine 372 mg, threonine 356 mg, tryptophan 152 mg, alanine 176 mg, glycine 176 mg, arginine 880 mg, proline 356 mg, tyrosine 34 mg, Na+ 70 mM, K+ 60 mM, Mg2+ 5 mM, Cl- 70 mM, PO4

- 30 mM, acetate 130 mM); Baxter Healthcare Corporation. 630% Intralipid® (soybean oil 30g phospholipids 1.2g glycerine 1.7g)/100 ml; Baxter Healthcare Corporation. 7Sigma-Aldrich Co., St. Louis, Missouri. 8Contains: chromium .3 µmol/L; copper 25.2 µmol/L; iodine 1.9 µmol/L; manganese 21.6 µmol/L; selenium 1.0 µmol/L; zinc 492 µmol/L. 9Fujisawa USA, Inc. Deerfield, Illinois. 10Contains; elemental iron 100mg/ml; Rhone Merieux, Inc., Athens, Georgia. 11Contains: calcium 2.14g/100 ml.

49

50

Table 2.1, continued

12Vital E® -A+D Durvet; Contains: vitamin A 2331 IU; vitamin E 233 IU; vitamin D 7 IU; Boehringer Ingelheim Animal Health, Inc., St. Joseph, Missouri. 13Contains; phytonadione 10 mg/ml; K-Ject Vetus Animal Health; Burns Veterinary Supply Inc., Rockville Centre, New York. 14Vedco Inc., St. Joseph, Missouri. 15Contains: sodium ascorbate 250 mg/ml. 16Contains: thiamin hydrochloride (B1) 12.5 mg/ml; niacinamide (B3) 12.5 mg/ml; pyridoxine hydrochloride (B6); d-panthenol 5 mg/ml; riboflavin (B2; as riboflavin 5’ phosphate sodium) 2 mg/ml; cyanocobalamin (B12) 5 µg/ml. 17Contains: folic acid 5 mg/ml; American Pharmaceutical Partners, Inc., Los Angeles, California. 18Contains: d-biotin 20 mg/ml.

Table 2.2: Effect of short-chain fatty acid-supplemented total parenteral nutrition on total villus and crypt goblet cells following massive small bowel resection in neonatal piglets1,2

TREATMENT Control 9 Bu 60 Bu SCFA VILLUS Jejunum 12h 0.038 ± 0.006 0.042 ± 0.007 0.030 ± 0.004 0.036 ± 0.006 72h 0.036 ± 0.005 0.034 ± 0.005 0.039 ± 0.006 0.033 ± 0.005 Ileum 12h 0.025 ± 0.004 0.021 ± 0.006 0.021 ± 0.007 0.041 ± 0.007 72h 0.029 ± 0.006 0.036 ± 0.006 0.025 ± 0.005 0.027 ± 0.005 CRYPT Jejunum3

12h 0.093 ± 0.007 0.095 ± 0.007 0.085 ± 0.007 0.101 ± 0.010 72h 0.076 ± 0.008 0.082 ± 0.007 0.088 ± 0.008 0.091 ± 0.007 Ileum4

12h 0.096 ± 0.009 0.103 ± .00802 0.109 ± 0.008 0.106 ± 0.008 72h 0.073 ± 0.009 0.080 ± 0.008 0.094 ± 0.008 0.081 ± 0.008 Colon 12h 0.126 ± 0.011 0.113 ± 0.011 0.120 ± 0.012 0.137 ± 0.013 72h 0.116 ± 0.013 0.121 ± 0.012 0.133 ± 0.015 0.123 ± 0.013

1 All values are mean ± SEM expressed as relative goblet cell number per depth and is adjusted to include neutral and acidic fractions of mixed cells. Data were analyzed statistically as one-tailed t-tests in split-plot design using the mixed model in SAS 9.1 2 Abbreviations: 9 mM butyrate (9Bu), 60 mM butyrate (60Bu), 60 mM short chain fatty acid (SCFA). 3 Within this dependent variable, 12h tended to be greater than 72h (time main effect, p=0.074).

51

Table 2.2, continued 4 Within this dependent variable, 12h is greater than 72h (time main effect, p=0.0004).

52

Table 2.3: Effect of short-chain fatty acid-supplemented total parenteral nutrition on neutral and acidic goblet cell chemotypes following massive small bowel resection in neonatal piglets1,2

TREATMENT Control 9 Bu 60 Bu SCFA NEUTRAL Jejunum3

12h 0.0383 ± 0.0043 0.0377 ± 0.0043 0.0284 ± 0.0043 0.0405 ± 0.0062 72h 0.0243 ± 0.0054 0.0235 ± 0.0043 0.0264 ± 0.0048 0.0247 ± 0.0045 Ileum4

12h 0.0285 ± 0.0039 0.0274 ± 0.0034 0.0309 ± 0.0039 0.0271 ± 0.0037 72h 0.0230 ± 0.0028 0.0233 ± 0.0029 0.0258 ± 0.0033 0.0231 ± 0.0029 Colon 12h 0.0479 ± 0.0055 0.0380 ± 0.0055 0.0421 ± 0.0061 0.0410 ± 0.0070 72h 0.0356 ± 0.0068 0.0358 ± 0.0060 0.0296 ± 0.0080 0.0453 ± 0.0069 ACIDIC Jejunum 12h 0.0012 ± 0.0003 0.0013 ± 0.0003 0.0013 ± 0.0003 0.0015 ± 0.0006 72h 0.0010 ± 0.0003 0.0013 ± 0.0003 0.0011 ± 0.0003 0.0017 ± 0.0005 Ileum5,6

12h 0.0669 ± 0.0056 0.0746 ± 0.0051 0.0779 ± 0.0051 0.0691 ± 0.0056 72h 0.0516 ± 0.0056 0.0543 ± 0.0051 0.0684 ± 0.0051 0.0571 ± 0.0051 Colon7

12h 0.0779 ± 0.0070 0.0754 ± 0.0070 0.0779 ± 0.0078 0.0961 ± 0.0088 72h 0.0802 ± 0.0087 0.0856 ± 0.0078 0.1040 ± 0.0102 0.0785 ± 0.0090

53

54

Table 2.3, continued

1 All values are mean ± SEM expressed as relative goblet cell number per depth and is adjusted to include neutral and acidic fractions of mixed cells. Data were analyzed statistically as one-tailed t-tests in split-plot design using the mixed model in SAS 9.1 2 Abbreviations: 9 mM butyrate (9Bu), 60 mM butyrate (60Bu), 60 mM short chain fatty acid (SCFA). 3 Within this dependent variable, 12h was greater than 72h (time main effect, p=0.005). 4 Within this dependent variable, 12h was greater than 72h (time main effect, p=0.012). 5 Within this dependent variable, 12h was greater than 72h (time main effect, p=0.0003). 6 Within this dependent variable, 60Bu was greater than control (treatment main effect, p=0.038). 7 Within this dependent variable, SCFA tended to be greater than control at 12h and 60Bu tended to be greater than control at 72h (0.060).

Table 2.4: Effect of short-chain fatty acid-supplemented total parenteral nutrition on villus neutral mucin and acidomucin chemotypes per villus height following massive small bowel resection in neonatal piglets1,2

TREATMENT Control 9 Bu 60 Bu SCFA Mean Mean Mean Mean NEUTRAL Jej unum 12h 0.01861 ± 0.00451 0.02178 ± 0.00594 0.01210 ± 0.00266 0.01490 ± 0.00407 72h 0.01370 ± 0.00301 0.01406 ± 0.00309 0.01425 ± 0.00313 0.01294 ± 0.00284 Ileum 12h 0.00006 ± 0.00003 0.00009 ± 0.00005 0.00006 ± 0.00004 0.00011 ± 0.00006 72h 0.00003 ± 0.00002 0.00006 ± 0.00003 0.00006 ± 0.00004 0.00004 ± 0.00003 ACIDIC Jej unum 12h 0.01862 ± 0.00318 0.01821 ± 0.00349 0.01730 ± 0.00269 0.02037 ± 0.00390 72h 0.01994 ± 0.00310 0.01885 ± 0.00293 0.01929 ± 0.00299 0.01885 ± 0.00293 Ileum3

12h 0.00930 ± 0.00199 0.01346 ± 0.00320 0.01632 ± 0.00441 0.02161 ± 0.00462 72h 0.01622 ± 0.00387 0.02027 ± 0.00437 0.01032 ± 0.00280 0.01533 ± 0.00376

1 All values are mean ± SEM expressed as relative goblet cell number per depth and is adjusted to include neutral and acidic fractions of mixed cells. Data were analyzed statistically as one-tailed t-tests in split-plot design using the mixed model in SAS 9.1 2 Abbreviations: 9 mM butyrate (9Bu), 60 mM butyrate (60Bu), 60 mM short chain fatty acid (SCFA). 3 Within this dependent variable, SCFA and 60Bu were greater than control at 12h (p=0.031); however these differences were not sustained at 72h.

55

Table 2.5: Effect of short-chain fatty acid-supplemented total parenteral nutrition on crypt sulfomucin and sialomucin chemotypes per crypt depth following massive small bowel resection in neonatal piglets1,2

TREATMENT Control 9 Bu 60 Bu SCFA SULFATED Jej unum 12h 0.06551 ± 0.01052 0.06391 ± 0.01052 0.05886 ± 0.00969 0.07864 ± 0.00969 72h 0.06376 ± 0.00963 0.06069 ± 0.00963 0.06384 ± 0.00958 0.05311 ± 0.01003 I leum 12h 0.04090 ± 0.00991 0.03207 ± 0.00738 0.04613 ± 0.01605 0.04947 ± 0.01137 72h 0.04394 ± 0.01014 0.04626 ± 0.00995 0.04510 ± 0.00960 0.05370 ± 0.01213 Colon 12h 0.13490 ± 0.00945 0.13690 ± 0.01059 .13130 ± 0.01015 0.13350 ± 0.01015 72h 0.13470 ± 0.01189 0.13700 ± 0.01048 .12280 ± 0.01409 0.13180 ± 0.01074 SIALYLATED Jej unum 12h 0.00008 ± 0.00017 0.00027 ± 0.00035 0.00045 ± 0.00048 0.00010 ± 0.00018 72h 0.00004 ± 0.00011 0.00013 ± 0.00019 0.00001 ± 0.00009 0.00019 ± 0.00026 Ileum3

12h 0.05824 ± 0.00844 0.06547 ± 0.00844 0.05856 ± 0.00895 0.04849 ± 0.00844 72h 0.04580 ± 0.00799 0.04242 ± 0.00807 0.03523 ± 0.00867 0.04545 ± 0.00845 Colon4

12h 0.00101 ± 0.00057 0.00051 ± 0.00062 0.00114 ± 0.00062 0.00084 ± 0.00062 72h 0.00304 ± 0.00070 0.00082 ± 0.00062 0.00004 ± 0.00081 0.00004 ± 0.00065

56

57

Table 2.5, continued

1 All values are mean ± SEM expressed as relative goblet cell number per depth and is adjusted to include sulfated and sialylated fractions of mixed cells. Data were analyzed statistically as one-tailed t-tests in split-plot design using the mixed model in SAS 9.1 2 Abbreviations: 9 mM butyrate (9Bu), 60 mM butyrate (60Bu), 60 mM short chain fatty acid (SCFA). 3 Within this dependent variable, 12h tended to be greater than 72h (time main effect, p=0.054). 4 Within this dependent variable, control was greater than 9Bu, 60Bu, and SCFA at 72h (p=0.039).

Table 2.6: Effect of short-chain fatty acid-supplemented total parenteral nutrition on villus sulfomucin and sialomucin chemotypes per villus height following massive small bowel resection in neonatal piglets1,2

TREATMENT Control 9 Bu 60 Bu SCFA SULFATED Jejunum3

12h 0.0343 ± 0.0038 0.0471 ± 0.0038 0.0395 ± 0.0034 0.0371 ± 0.0034 72h 0.0364 ± 0.0034 0.0403 ± 0.0034 0.0340 ± 0.0038 0.0351 ± 0.0035 Ileum 12h 0.0321 ± 0.0043 0.0317 ± 0.0038 0.0383 ± 0.0057 0.0343 ± 0.0045 72h 0.0280 ± 0.0031 0.0315 ± 0.0037 0.0290 ± 0.0031 0.0334 ± 0.0044 SIALYLATED Jejunum4

12h 0.0004 ± 0.0003 0.0008 ± 0.0005 0.0005 ± 0.0004 0.0004 ± 0.0003 72h 0.0010 ± 0.0006 0.0017 ± 0.0009 0.0006 ± 0.0004 0.0021 ± 0.0012 Ileum5

12h 0.0027 ± 0.0006 0.0035 ± 0.0006 0.0019 ± 0.0006 0.0008 ± 0.0006 72h 0.0015 ± 0.0006 0.0005 ± 0.0005 0.0024 ± 0.0005 0.0021 ± 0.0006

1 All values are mean ± SEM expressed as relative goblet cell number per depth and is adjusted to include sulfated and sialylated fractions of mixed cells. Data were analyzed statistically as one-tailed t-tests in split-plot design using the mixed model in SAS 9.1 2 Abbreviations: 9 mM butyrate (9Bu), 60 mM butyrate (60Bu), 60 mM short chain fatty acid (SCFA). 3 Within this dependent variable, 9Bu was greater than control, (treatment main effect, p=0.047). 4 Within this dependent variable, 72h tended to be greater than 12h (time main effect, p=0.058). 5 Within this dependent variable, control was greater than SCFA at 12h; however this difference was not seen at 72h (p=0.0004).

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Table 2.7: Effect of short-chain fatty acid supplemented total parenteral nutrition on muc2 mRNA following massive small bowel resection in neonatal piglets1,2

TREATMENT Control 9 Bu 60 Bu SCFA Jejunum3

12h 0.571 ± 0.193 2.112 ± 0.788 0.775 ± 0.225 0.930 ± 0.269 72h 0.878 ± 0.254 1.138 ± 0.336 0.800 ± 0.232 0.578 ± 0.195 Ileum4

12h 0.885 ± 0.264 0.723 ± 0.264 1.004 ± 0.264 0.562 ± 0.264 72h 0.883 ± 0.264 1.494 ± 0.264 1.052 ± 0.262 1.138 ± 0.262 Colon 12h 0.508 ± 0.187 0.461 ± 0.226 0.671 ± 0.083 0.593 ± 0.156 72h 0.636 ± 0.167 0.712 ± 0.187 0.446 ± 0.118 0.556 ± 0.146

1 All values are mean ± SEM expressed as muc2 mRNA/18S mRNA. Data were analyzed statistically as one-tailed t-tests in split-plot design using the mixed model in SAS 9.1. 2 Abbreviations: 9 mM butyrate (9Bu), 60 mM butyrate (60Bu), 60 mM short chain fatty acid (SCFA). 3 Within this dependent variable, 9Bu was greater than control (treatment main effect, p=0.010). 4 Within this dependent variable, 72h was greater than 12h (p=0.017).

59

Figure 2.1: Gel electrophoresis. DNA ladder is present in the left lane with the bottom four bands being 50 base pairs, 100 base pairs, 150 base pairs, and 200 base pairs. The product of rt-PCR of the cDNA of all samples pooled is in the right lane and contains both 18S and muc2 amplicon. The 18S DNA amplicon is 187 base pairs which is present as a faint band. The muc2 amplicon is 64 base pairs and is not visible in the right lane.

60

Acidic mucin chemotype

Neutral mucin chemotype

Figure 2.2: Periodic acid schiff’s alcian blue (PASAB) stain of ileal tissue from a 72h piglet receiving 9Bu treatment. Image is at 10x magnification. PASAB stain colors neutral mucins pink and acidomucins blue.

61

62

Figure 2.3: PASAB stain of ileal villi from a 72h piglet receiving 9Bu treatment. Image is at 40x magnification. PASAB colors neutral mucins pink and acidomucins blue. Goblet cells were counted in one of the 5 following groups: 1.) 100% acidic mucin; 2.) 75% acidic mucin and 25% neutral mucin; 3.) 50% acidic mucin and 50% neutral mucin; 4.) 25% acidic and 75% neutral mucin; and 5.) 100% neutral mucin. These mixed goblet cells were assigned to the group that most closely matched their composition.

75% acidic mucin, 25% neutral mucin

25% acidic mucin, 75% neutral mucin

Sulfomucin chemotype

Sialomucin chemotype

Figure 2.4: High iron diamine alcian blue (HIDAB) stain of jejunal tissue from a 12h control piglet. Image is at 10x magnification. HIDAB stain colors sulfomucins brown and sialomucins blue.

63

75% sulfomucin; 25% sialomucin

50% sulfomucin; 50% sialomucin

Figure 2.5: High iron diamine alcian blue (HIDAB) stain of jejunal crypts of a 12h control piglet. Image is at 40x magnification. HIDAB stain colors sulfomucins brown and sialomucins blue. Goblet cells were counted in one of the 5 following groups: 1.) 100% sulfomucin; 2.) 75% sulfomucin and 25% sialomucin; 3.) 50% sulfomucin and 50% sialomucin; 4.) 25% sulfomucin and 75% sialomucin; and 5.) 100% sialomucin. These mixed goblet cells were assigned to the group that most closely matched their composition.

64

0.00

0.03

0.05

0.08

0.10

Treatment

Ilea

l C

ryp

t A

cid

omu

cin

Gob

let

Cel

l C

hem

otyp

es p

er D

epth

Control 9 Bu Control 9 Bu60 Bu 60 BuSCFA SCFA

12 h 72 h

* 60Bu > Control, treatment main effect

Figure 2.6: Effect of SCFA and Bu supplementation on ileal crypt acidomucin chemotypes. Data is expressed as relative goblet cell number per depth and is adjusted to include neutral and acidic fractions of mixed cells. *Adjusted acidic goblet cell number was greater in the 60Bu treatment group than control (treatment main effect, p=0.038). Data were analyzed statistically as one-tailed t-tests in split-plot design using the mixed model in SAS 9.1.

65

0.000

0.008

0.015

0.023

0.030

Treatment

Ilea

l V

illu

s A

cid

ic G

oble

t C

ell

Ch

emot

ypes

per

Hei

gh

t

Control 9 Bu Control 9 Bu60 Bu 60 BuSCFA SCFA

12 h 72 h

*

*

Figure 2.7: Effect of SCFA and Bu supplementation ileal villus acidomucin chemotypes. Data is expressed as relative goblet cell number per height and is adjusted to include neutral and acidic fractions of mixed cells. *Significantly different than respective control group (p=0.031). Data were analyzed statistically as one-tailed t-tests in split-plot design using the mixed model in SAS 9.1.

66

0

0.001

0.002

0.003

0.004

Treatment

Col

onic

Cry

pt

Sial

omu

cin

Gob

let

Cel

l C

hem

otp

yes

per

Dep

th

*

Control 9 Bu Control 9 Bu60 Bu 60 BuSCFA SCFA

12 h 72 h

**

Figure 2.8: Effect of SCFA and Bu supplementation on colonic crypt sialomucin chemotypes. Data is expressed as relative goblet cell number per depth and is adjusted to include sulfated and sialated fractions of mixed cells. *Significantly different than respective control group (p=0.039). Data were analyzed statistically as one-tailed t-tests in split-plot design using the mixed model in SAS 9.1.

67

0.00

0.02

0.03

0.05

0.06

Treatment

Jeju

nal

Vil

lus

Sulf

omu

cin

Gob

let

Cel

l C

hem

otyp

e p

er H

eig

ht

Control 9 Bu Control 9 Bu60 Bu 60 BuSCFA SCFA

12 h 72 h

* 9Bu > Control, treatment main effect

Figure 2.9: Effect of SCFA and Bu supplementation on jejunal villus sulfomucin chemotypes. Data is expressed as relative goblet cell number per height and is adjusted to include sulfated and sialated fractions of mixed cells. *Sulfomucin chemotypes are greater in 9 Bu than control (treatment main effect, p=0.047). Data were analyzed statistically as one-tailed t-tests in split-plot design using the mixed model in SAS 9.1.

68

0.00

0.50

1.00

1.50

2.00

2.50

3.00

3.50

Treatment

Jeju

nal

muc

2 m

RN

A/

18S

mR

NA

exp

ress

ed a

s fo

ld c

han

ge

Control 9 Bu Control 9 Bu60 Bu 60 BuSCFA SCFA

12 h 72 h

* 9Bu > Control, treatment main effect

Figure 2.10: Effect of SCFA and Bu supplementation on jejunal muc2 mRNA. Data is expressed as muc2 mRNA/18s mRNA. *The 9Bu group had greater muc2 mRNA than control, (treatment main effect, p=0.010). Data were analyzed statistically as one-tailed t-tests in split-plot design using the mixed model in SAS 9.1.

69

CHAPTER 3 FUTURE DIRECTIONS

SBS and PN administration both contribute to intestinal barrier dysfunction and bacterial

translocation, and these data demonstrate butyrate’s effectiveness in improving innate barrier

defense and presents a promising treatment for decreasing rates of bacterial translocation. Within

this study, exploring bacterial infection in the surrounding lymph nodes and the liver and kidneys

would be an interesting avenue of study to elucidate whether the increases in protective goblet cell

chemotypes observed in piglets receiving SCFA and butyrate supplementation decreased bacterial

translocation. Additionally, to be sure that any decreases in bacterial infection are attributed to

increases in protective mucins, tight junction protein abundance and mRNA expression could also

be assessed.

Since PN is much more common than TPN, the effect of butyrate and SCFA administration

on goblet cell chemotypes and muc2 in the presence of EN is an important relationship to establish.

This avenue would also be important to explore since practitioners try to avoid TPN with most SBS

infants and instead try to administer some EN to maintain as much bowel function as possible.

Because butyrate has been shown to have profound effects on mucin production when administered

luminally (Finnie et al., 1995; Willemsen et al., 2003; Gaudier et al., 2004; Hatayama et al., 2007), and

PN is associated with decreased mucin production (Iiboshi et al., 1994; Bertolo et al., 1998; Law et

al., 2007), the positive effects of PEN in conjunction with parenteral butyrate or SCFA

administration on protective goblet cell chemotypes and muc2 mRNA could be additive.

To study the effects of butyrate administration in addition to EN, butyrate could clinically

only be supplied parenterally, as butyrate is not tolerable as an oral solution. However, oral

administration of tributyrin is a possibility. Tributyrin is a prodrug form of butyrate which means

that it is inactive until metabolized, at which time it becomes an active metabolite. Tributyrin has

70

been shown to be tolerated in an adult clinical trial and 3 daily oral dosings can attain in vitro levels of

butyrate seen in many studies (Edelman et al., 2003). Tributyrin confers positive effects on intestinal

morphology by increasing villus height and crypt depth when administered with a fermentable

substrate in pigs (Piva et al., 2002; Piva et al., 2008). Further, administration of a 1:1 mixture of

triacetin and tributyrin in rats with 60% bowel resection and cecectomy has been demonstrated to

significantly increase jejunal mucosa, all intestinal segmental weights, and all intestinal segmental

protein levels (Kripke et al., 1991). While in this study it cannot be determined if the positive effects

on intestinal adaptation are due to tributyrin rather than triacetin, this is expected to be the case due

to all the literature stating specifically butyrate’s positive effects on intestinal adaptation (Tappenden

et al., 1997; Bartholome et al., 2004; Hamer et al., 2008) and recent findings supporting butyrate’s

relationship with proglucagon and intestinotrophic GLP-2 (Woodard and Tappenden, 2008).

Another method to increase butyrate availability to the intestinal epithelium is to increase its

endogenous production. This can be done by feeding fermentable fiber along with PEN that is then

digested by the resident microbiota to create SCFAs, including butyrate (Bourquin et al., 1993;

Kapadia et al., 1995). Additionally, some fibers increase butyrate production more than others

(Kapadia et al., 1995). This method may be less direct than administration of a prodrug; however, it

has other benefits. Feeding certain fibers not only increases SCFA production, but can also change

the composition of the intestinal microbiota by increasing the density of the resident microbes,

which can decrease the growth of harmful microbes such as Clostridium difficile (May et al., 1994).

Increasing butyrate production and potentially protective mucins could also decrease growth of

harmful bacteria, which would be obviously beneficial in a short bowel syndrome infant at risk for

bacterial translocation. Both animal studies and clinical studies exploring the effects of tributyrin

and fiber administration are needed to validate the effectiveness of these methods on mucin

production and on possible subsequent prevention of bacterial translocation.

71

72

In addition, the mechanism by which butyrate increases mucin production needs to be

explored. It may be that butyrate increases or decreases an intermediate factor that affects mucin

production such as affecting mRNA or protein expression of sulfotransferases or sialotransferases.

If there is an intermediate factor, its discovery would allow expedited clinical targeting of increasing

mucin production.

In summary, the area of mucin modulatioin via SCFAs and butyrate is novel and much is yet

to be discovered. Butryate is a promising supplement to potentially improve the clinical outcomes

of the SBS pediatric population via improved barrier defense.

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CURRICULUM VITAE

E  RICA WHITNEY NEHRLING

U  Division of Nutritional Sciences 

n n42   iversity of Illinois at Urbana‐Champaig0 Bevie win AveUrbana,  – 0776 

r Hall, 905 South Good IL 61820     (217) 265enehrli2@illinois.edu 

  FORMAL EDUCATION  

Masters student in Di ional Sciencniv nois at aign 

6/2007 – 8/2010  vision of Nutrit Urbana Champ

Expe

es       U ersity of Illi

cteGPA: 3.8

      d Graduation: August 2010       8/4.00       Advisor: Kelly A. Tappenden, PhD, RD 

Thesis research:  The Effect of Butyrate‐Supplemented Total Parenteral Nutrition on muc2 mRNA and Goblet Cell Chemotypes in a Short Bowel Syndrome Neonatal Piglet Model.  Dietetic Intern University of Illinois at Urbana Champaign 

8/2009 – 5/2010    

Graduated May 2010        2001 ‐ 2005    d Consumer Sciences Bachelor of Science in Family an

Cum L  Honors Univ rgia, Athens, GA 

      aude with

Majo        ersity of Geo

    r: Dietetics    GPA: 3.67/4.00 

     OTHER EDUCATION  The ACES Academy of Teaching Excellence  College of ACES Teaching College Course 2009 Graduated November 2009  The University of Illinois at Urbana Champaign, College of Business Certificate in Business Administration Graduated Spring 2009   

90

SCHOLARSHIPS, AWARDS, AND HONORS  University  Illinoi  at Urbana­Champaign 

   2009   

of  s2010  William Rose Endowed Award 2007‐ Jonathan Baldwin Turner Fellowship 2009  Abbott Nutrition Scholarship for Certificate in Business 

Administration  University 001‐2005 

of Georgia, Athens, GA 2 HOPE Scholarship (full tuition) 

te Scholar olar (  semesters) 

National Collegia  Presidential Sch 42004  Top 5% of class    RESEARCH EXPERIENCE  University of Illinois at Urbana Champaign 

appenden, PLaboratory of Kelly A. T hD, RD 

0   une 2007 – June 20 9   Research Fellow uly 200  – Pr sent      Research Assistant JJ 9 e Studies:    

ion on Mucin Production and Barrier Function in g TPN 

2009: Effect of Butyrate Supplementat Receivinf Health 

Short Bowel Syndrome PigletsFunding: National Institutes oPI: Kelly Tappenden, PhD, RD Selected for Oral Presentation at Experimental Biology Conference 2010 for the merican Society for Nutrition symposia “Animal Models in Gastrointestinal Health 

sease” Aand Di 2008:  Evaluation of GATTEX Effects on Intestinal Development in Neonatal Piglets Funding: NPS Pharmaceuticals PI: Kelly Tappenden, PhD, RD 

ry on the Response to Dietary Prebiotics in Neonatal Piglets  

e2008: Impact of Route of DelivFunding: Bristol‐Myers Squibb I: Sharon Donovan, PhD, RD PRole: Intestinal electrophysiology, ion transport and ion secretion  

 and Early Nutrition on Intestinal Development in the 2007: Effect of Mode of DeliveryNeonatal Piglet 

91

Funding: Bristol‐Myers Squibb PI: Sharon Donovan, PhD, RD Role: Intestinal electrophysiology, ion transport and ion secretion 

92

TEACHING EXPERIENCE niversity of Illinois at Urbana Champa

entor 2008      ign U

Research Apprentice Program M

hools  

005 – 2006    Rockdale County Public Sc2      Substitute Teacher 

004 – 005  

 2   University of Georgia gy Labs I and II 

2      Teaching Assistant: Anatomy and Physiolo

002 – 2004    Athens Clarke County Mentoring Program  2  PROFESSIONAL EXPERIENCE 4­H Illini Food Science and Human Nutrition Summer Academy  University of Illinois, Department of Food Science and Human Nutrition Co­Coordinator            January 2009 ­ July 2009 FSHN Summer Academy dates        June 29th – July 1st 2009 

• udget, recruited  

Developed and implemented creative FHSN programming, devised b

• professors Delivered lessons on anatomy, physiology, and molecular nutrition Generated interest for University of Illinois and for FSHN through program 

• f •

Worked with 4‐H Extension Specialist, Deb Stocker, to create IRB to evaluate efficacy oprogram 

• Students who participated in IRB evaluation marked “agreed” or “strongly agreed” on areas of high enjoyment of academy, retention of FSHN concepts from academy, and recommendation of academy to peers. 

 Women, Infants, Children Program Eas Covington, GA t Metro Health District, Newton County Health Department; 

• Nutritionist              July 2006 – May 2007 

Assess nutritional risk and supply nutrition education to clients • h Provide nutrition education to community through Nutrition Education Outreac

Program • Involved  in statewide committee devoted to changing WIC counseling process  PROFESSIONAL ORGANIZATIONS American Dietetic Association 

n American Society of Nutrition American Society of Parenteral and Enteral Nutritioniversity of Georgia Alumni Association utritional Sciences Graduate Student Association, Secretary, 2008 

UN