The Role of Adenosine Monophosphate Deaminase 1 (AMPD1) and Inflammatory

Cytokines in Mediating Muscle Weakness in a Mouse Model of Idiopathic

Inflammatory Myopathy

by William Coley

B.S. in Biology, May 2005, University of Virginia

A Dissertation submitted to

The Faculty of

Columbian College of Arts and Sciences

of The George Washington University

in partial fulfillment of the requirements

for the degree of Doctor of Philosophy

January 31st, 2013

Dissertation directed by

Kanneboyina Nagaraju

Professor of Integrative Systems Biology and Pediatrics

ii

The Columbian College of Arts and Sciences of The George Washington University

certifies that William D. Coley has passed the Final Examination for the degree of Doctor

of Philosophy as of August 24, 2012. This is the final and approved form of the

dissertation.

William D. Coley

Dissertation Research Committee:

Kanneboyina Nagaraju, Professor of Integrative Systems Biology and Pediatrics,

Dissertation Director

Robert Freishtat, Assistant Professor of Pediatrics and Emergency Medicine, Committee

Member

Joseph Devaney, Assistant Professor of Integrative Systems Biology and of Pediatrics,

Committee Member

The Role of Adenosine Monophosphate Deaminase 1 (AMPD1) and

Inflammatory Cytokines in Mediating Muscle Weakness in a Mouse

Model of Idiopathic Inflammatory Myopathy

iii

This dissertation is dedicated to my beautiful wife Catherine Larsen Coley, whose

unwavering support and kindness kept me from despairing in the worst of times. I also

must give thanks to my lord God, with prayers that this work will be a contribution to

medicine and an act of mercy to my neighbors.

iv

The author wishes to acknowledge several people who were instrumental in the completion

of the research presented in this dissertation. This work would not have been possible

without the assistance and support of Sree Rayavarapu, Dr. Jack van der Meulen, Arpana

Sali, Dr. Richard Sabina, Dr. Beryl Ampong, Sreedatta Mahadavi, Travis Kinder, Dr.

Rashmi Rawat, Dr. Ron Jubin, Dr. Gouri S. Pandey, Dr. Robert Wortmann, and Dr.

Heather Alger. The author further wishes to thank all of the faculty members who

dedicated their time to serve as thesis committee members while this research was

conducted. Those faculty members were Dr. Kanneboyinna Nagaraju (mentor), Dr.

Stephanie Constant (chair), Dr. Yi-Wen Chen (chair), Dr. Robert Freishtat (reader), Dr.

Joseph Devaney (reader), Dr. Sasa Radoja (member), Dr. Mary Rose (member), and Dr.

Philipe Andrade (outside examiner).

v

Abstract of Dissertation

The Role of AMPD1 and Inflammatory Cytokines in Mediating Muscle Weakness a

Mouse Model of Idiopathic Inflammatory Myopathy

PROBLEM: Myositis is a term used to describe a group of related autoimmune diseases

that affect skeletal muscle tissues. These diseases are also described as idiopathic

inflammatory myopathies, reflecting that they are characterized by significant muscle

weakness and lymphocyte infiltration into the muscle tissue due to unknown causes.

Patients with myositis will typically be diagnosed with one of the three most common

varieties: polymyositis, dermatomyositis, or inclusion body myositis. While there is no

question that there is an autoimmune reaction in these patients, it has been observed that

there is dissociation between autoimmune inflammation and muscle weakness. Treatment

with immunosuppressive compounds does not guarantee a recovery in muscle function, and

there is a poor correlation between degree of infiltration and muscle weakness. In essence,

there is a non-immune component that may cause muscle weakness in this disease.

OBJECTIVE: Prior investigations in humans have suggested a potential mechanism that is

independent of the action of autoreactive T-lymphocytes or autoantibodies to explain

muscle weakness in patients. It has been proposed that the persistent weakness is due to an

acquired deficiency of the metabolic enzyme adenosine monophosphate deaminase

(AMPD1). While there is substantial data to support this hypothesis, it cannot be directly

tested in humans. We were able test this hypothesis in an inducible transgenic mouse model

of myositis that closely mimics the human disease phenotype. Using these mice, we tested

the proposed hypothesis that a) AMPD1 is responsible for muscle weakness in myositis, b)

vi

supplements of D-ribose can replace the downstream metabolites of AMPD1 and

ameliorate disease symptoms and c) cytokines of the innate immune response modulate the

expression of AMPD1. This work required in vitro skeletal muscle cell culture techniques

and in vivo mouse phenotyping, histological, molecular, genetic and biochemical assays.

RESULTS: We observed that the onset of muscle weakness does indeed coincide with an

acquired deficiency of AMPD1 in skeletal muscle, in addition to a general suppression of

enzymes related to glycolysis. The loss of AMPD1 was specific to myositis, and not

observed in other myopathies. We made the novel observation that muscle weakness and

decreased AMPD1 activity occurred prior to the appearance of infiltrating lymphocytes in

skeletal muscle tissue. We found that the partial knockdown of AMPD1 in mice resulted in

significant muscle weakness in healthy mice, and corroborated this data with mice

heterozygous for the AMPD1 gene. We observed that metabolites downstream of AMPD1

(e.g. IMP and hypoxanthine) were significantly decreased. Attempts to replace these

metabolites with oral supplements of D-ribose proved to be ineffective as a treatment. We

found that innate immune cytokines such as Type I interferons and Interleukin-1 were

able to inhibit the expression of AMPD1 in vitro and induce weakness in vivo.

Additionally, we observed that IL-15 had a stimulatory effect on the expression of AMPD1

and that mice with myositis had significantly lower levels of the IL-15 receptor. These data

for the first time demonstrate a link between innate immune systems and energy generating

muscle metabolic pathways. Future experiments to modulate this link between innate

immune system and metabolic pathway would identify newer drugs that specifically target

muscle weakness and thus improve the quality of life for patients with myositis.

vii

Table of Contents

Dedication .......................................................................................................................... iii

Acknowledgments ............................................................................................................ iv

Abstract of Dissertation ................................................................................................... v

List of Figures ..................................................................................................................viii

List of Tables ...................................................................................................................... x

Chapter 1: Introduction ..................................................................................................... 1

Chapter 2: Characterization of muscle metabilic defects in myositis mice ......... 13

Chapter 3: The effects of D-ribose treatment in myositis mice ............................ 40

Chapter 4: The effects of inflammatory cytokines on AMPD1 expression ....... 60

Chapter 5: Conclusions and future directions ........................................................... 81

Chapter 6: Methods ........................................................................................................ 87

References ........................................................................................................................ 103

viii

List of Figures

Figure 1.1: AMPD1 mRNA levels are significantly reduced in myositis patients 7

Figure 2.1: The C34T allele is not present in HT mice 15

Figure 2.2: Overt symptoms are not visible until the infiltration stage 16

Figure 2.3: The EDL and soleus muscles of HT mice are severely weakened 18

Figure 2.4: HT mice show visible signs of myopathy at the infiltration stage 18

Figure 2.5: Loss of AMPD enzyme activity is detectable prior to disease onset 20

Figure 2.6: Diagram of the role of AMPD1 in purine catabolism 20

Figure 2.7: Loss of metabolites downstream of AMPD1 21

Figure 2.8: Infiltrated HT mice have lowered levels of AMPD1 protein 24

Figure 2.9: Loss of AMPD enzyme activity is specific to myositis 25

Figure 2.10: Muscles from myositis mice show a shift in fiber type composition 26

Figure 2.11: Morpholinos targeted against AMPD1 can reduce force generation 30

Figure 2.12: AMPD1 protein levels are significantly reduced in AMPD1+/-

mice 31

Figure 2.13: AMPD1+/-

mice show mild weakness in soleus muscle 32

Figure 3.1: AMP catabolism can generate free D-ribose 42

Figure 3.2: Mice with myositis are deficient for the breakdown metabolites 43

Figure 3.3: Treatment with oral D-ribose did not improve mouse bodyweight 46

Figure 3.4: Treatment with oral D-ribose did not improve voluntary movement 47

Figure 3.5: Histological analysis showed no difference between treated and untreated 50

Figure 3.6: Muscle force contraction analysis of the EDL showed no differences 51

Figure 3.7: Muscle force contraction analysis of the soleus showed no differences 52

ix

Figure 3.8: Treatment with D-ribose showed negligible effects on the fatigability 54

Figure 3.9: Quantitative PCR analysis suggests a lack of RBKS 55

Figure 4.1: A diagram of the AMPD1 promoter and its luciferase reporter 64

Figure 4.2: Expression from the AMPD1 promoter is affected by IFN and IL-15 67

Figure 4.3: Stably transfected myocytes also respond to IFN and IL-15 69

Figure 4.4: Receptors for IFN and IL-15 are expressed in mouse skeletal muscle 72

Figure 4.5: In vivo electroporation can specifically target expression plasmids 73

Figure 4.6: In vivo electroporation induces a low level of inflammation 74

Figure 4.7: Forced overexpression of IFN in vivo induces severe inflammation 74

Figure 4.8: Forced expression of IFN in vivo induces severe muscle weakness 77

Figure 4.9: Forced expression of IFN in vivo did not affect mRNA levels 78

x

List of Tables

Table 1: Twitch parameters in Type I (EDL) and Type II (soleus) muscles in HT mice 29

Table 2: Glycolysis related proteins downregulated in infiltrated HT myositis mice 35

Table 3: ER stress related proteins upregulated in infiltrated HT myositis mice 35

Table 4: IFN signature in HT mice 61

Table 5: Changes in gene expression for relevant cytokines and receptors 63

1

Chapter 1: Introduction

1.1 Background on Idiopathic Inflammatory Myopathies

The research summarized in this dissertation is focused on a set of closely related

autoimmune diseases that are collectively referred to as “idiopathic inflammatory

myopathy”, or simply “myositis”. This dissertation will focus more specifically on the

mechanisms of muscle weakness and their relationship to inflammation and cytokines of

the innate immune system in the muscle microenvironment. The term myositis usually

refers to one of three related diseases: polymyositis (PM), dermatomyositis (DM), and

inclusion body myositis (IBM). Less common form of myositis include juvenile

dermatomyositis and cancer-associated myositis. Each variation of the disease has defining

characteristics (e.g. the heliotrope rash of DM or the intramuscular rimmed vacuoles of

IBM), but there are some common characteristics. In PM and DM, patients typically

present clinically with diffuse, symmetrical muscle weakness that is worse in proximal

muscles than in distal muscles, whereas in IBM profound weakness may be present in both

proximal and distal muscles. Histologically, patients show muscle inflammation, fiber

degeneration, and overexpression of the MHC class I molecule 1-3

. This muscle pathology

is not always uniform, and many patients display isolated patches of inflammation and

degeneration in otherwise healthy muscle tissue. Patients with myositis are also commonly

diagnosed in interstitial lung disease 4. In affected patients, significant causes of morbidity

and mortality include difficulty in performing daily activities, dysphagia, and impaired

respiratory function 5,6

. Immunosuppressive therapies such as prednisone and methotrexate

are common first-line treatments, with cyclosporine and other powerful

2

immunosuppressants as second-line agents 7,8

. Unfortunately, these current therapies have

produced mixed results; for example, patients with DM are most likely to benefit from

glucocorticoid therapy (with a reduction in inflammation, and improvement in muscle

function), and most PM patients typically respond to immunosuppressive therapy9,10

. On

the other hand, patients with IBM (and a subset of PM patients) do not typically respond

either glucocorticoids or other immunosuppressant therapies 8. Studies have shown that the

therapeutic response to glucocorticoid treatment varies significantly; furthermore, the

degree of inflammatory infiltration and muscle function correlate poorly, suggesting a role

for other mechanisms in muscle dysfunction in these diseases 11,12

.

1.2 Dissociation between Muscle Weakness and Inflammation

Idiopathic inflammatory myopathies are treated as autoimmune diseases, and it is generally

believed that the weakness seen in patients is solely due to the inflammatory and cytotoxic

actions of infiltrating autoreactive lymphocytes. However, multiple lines of evidence

suggest that muscle weakness in myositis arises from a pathology that is independent of the

inflammatory response driven by autoreactive lymphocytes. For example, studies have

shown that a) there is a lack of correlation between the degree of inflammation and the

degree of muscle weakness 13,14

, b) a subgroup of myositis patients do not respond to large

doses of steroids 15,16

, and c) in some patients steroid treatment effectively eliminates the

inflammatory cells in the myositis muscle tissue, with little improvement in clinical disease

in some patients 12

. Patients with chronic myositis show clinical disease without any

identifiable inflammation (as assessed by histological analysis or magnetic resonance

imaging) 17,18

. These observations suggest that the autoimmune response arises in parallel

3

with a myopathy that is not reversed by immunosuppression. Thus, the molecular

mechanisms that account for muscle weakness in the absence of inflammation are still

unknown.

One potential mechanistic explanation for the refractory muscle weakness seen in

myositis patients comes from the observation that there is an apparent disturbance in

patients‟ metabolism and ability to regulate ATP production 19

. Muscles that must function

for long periods of time, such as postural muscles, are composed of slow-twitch fibers that

are dependent on oxidative phosphorylation. Conversely, muscles composed of fast-twitch

fibers rely on glycolysis in order to carry out rapid, strenuous movements even under

ischemic conditions. However, in the case of myositis patients, there are indications that the

fast-twitch fibers are more prone to degenerate, and that these patients‟ ability to produce

ATP in the skeletal muscle may be impaired 20

. Mass spectrometry analysis of myositis

biopsies has shown a significant loss of type II (fast-twitch)-specific proteins (e.g., myosin

heavy chain 1, troponin T3, and actinin 3), and a modest increase in type I (slow-twitch)

muscle proteins (e.g., myosin heavy chain 7, troponin T1, and actinin 2) 21

. Similarly,

myositis patients have shown a modest decrease in enzymes required for glycolysis in the

muscle (e.g., glycogen debranching enzyme, muscle phosphofructokinase, fructose-1,6-

bisphosphatase isozyme 2, and phosphoglycerate mutase 2), with the most significant

decreases being found in patients with IBM 21

. These findings have been corroborated by

magnetic resonance spectroscopy (MRS) in patients with juvenile-onset DM; in that study,

the stable isotope 31

P was used to compare the levels of adenosine triphosphate (ATP),

adenosine diphosphate (ADP), adenosine monophosphate (AMP), and phosphocreatine in

the quadriceps of DM patients and healthy controls. The DM patients were found to have

4

significantly less ATP at rest (3.50±0.3 mmoles/kg), as compared to healthy controls

(5.04±0.03 mmoles/kg, p<0.001). The concentrations of ADP, AMP, and phosphocreatine

were similarly decreased in the DM patients 18

.

Multiple studies have indicated that the loss of another type II-specific muscle

enzyme, adenosine monophosphate deaminase 1 (AMPD1), is potentially responsible for

these observed disturbances in ATP metabolism and muscle weakness 22-24

. An deficiency

of AMPD1 in patient muscle biopsies was first described by Fishbein et al. in 1985 after an

antibody specific for AMPD1 was generated in that lab and used to differentiate slow

twitch (type I) muscle fibers from fast twitch (type II) muscle fibers 25

. It was observed that

patients with myositis showed a deficiency of the enzyme, with protein levels dropping to

between 1-10% of normal levels. In 1991 Sabina et al., it was suggested that the

symmetrical muscle weakness seen in myositis strongly resembles the pattern of weakness

seen in patients with an inherited deficiency of AMPD1 19

. This observation demands extra

attention, as a congenital loss of AMPD1 had previously been recognized as causing

muscle weakness and quick fatigability in AMPD1 deficient subjects 24,26

.

1.3 Association between AMPD1 and Muscle Weakness

AMPD1 is preferentially expressed at high levels in type II skeletal muscle, where it

influences the levels of inorganic phosphate (Pi), AMP, ADP, and phosphocreatine.

AMPD1 is the rate-limiting enzyme in the catabolism of AMP to inosine 5‟-phosphate

(IMP) and ammonia (NH3). In the skeletal muscle, this reaction is an important means of

removing excess AMP created by rapid contractions during exercise. Other tissues express

low levels of homologous enzymes (either AMPD2 or AMPD3) which function as a part of

5

the broader nucleotide salvage pathway 27-30

. The expression of AMPD1 is limited to

skeletal muscle tissues, where it binds to the myosin heavy chain protein in order to achieve

peak enzymatic activity 31

. Patients with an inherited defect in AMPD1 expression often

show significantly diminished muscle performance, suggesting that the purine nucleotide

catabolic pathway plays a role in short-term energy production 22,23

. Myositis patients have

been found to have low AMPD enzyme activity along with reduced levels of AMPD1

protein and mRNA 19,24,25

. Although this finding has been reported previously, we were

able to reproduce this finding in a small cohort of patients with polymyositis and

dermatomyositis (Figure 1.1B). Myositis patients show significantly reduced levels of

mRNA for AMPD1. Taken together, these observations suggest that the refractory

symptoms of muscle weakness in myositis patients might be explained by an acquired

deficiency of the AMPD1 enzyme.

Unfortunately, the association between myositis and AMPD1 deficiency in humans

has been confounded by the prevalence of a certain allele, sometimes referred to as the

C34T allele (rs17602729,). This C34T allele is found most frequently in Caucasian

population (13.3%) but is absent Asian populations (0% for both Chinese and Japanese)

according to the public HapMap database. This allele introduces a premature stop codon in

the second exon of AMPD1, thereby terminating the protein after 11 amino acids and

completely abolishing enzyme activity. However, it has been observed that two splice

variants occur in humans; the major isoform which includes all 16 exons, and a minor

isoform which drops exon 2 without affecting enzyme activity 32

. Many investigators have

ignored the fact that AMPD1 has two common splice variants, and consequently have

6

assumed that a homozygous C34T individual must lack all enzyme activity. To further

complicate matters, the C34T allele appears to escape nonsense mediated degradation, so

that a homozygous C34T individual may not appear to be deficient for AMPD1 if only

QPCR analysis is performed 24

. Such incomplete investigations forgo testing patients for

AMPD1 enzyme activity, thereby introducing uncertainty into the data and potentially

arrive at erroneous conclusions. When we had the opportunity to examine patient biopsies,

we observed a C34T allele frequency of 10%, as three of the subjects were heterozygous

for the C34T allele (Fig. 1.1A). It should also be noted that two different mutations

(R388W and R425H) of AMPD1 have been identified in Japanese patients with very low

AMPD1 enzyme activity and symptoms of myalgia, weakness, and exercise intolerance 26

.

The existence of these additional diseases-associated mutations lends strong support to the

hypothesis that AMPD1 activity does indeed augment muscle function, and opposes the

contention that AMPD1 deficiencies have no relevance to muscle weakness.

7

8

Unlike patients with myositis, a congenital loss of AMPD1 (called monoadenylate

deaminase deficiency, or MADD) is not associated with autoimmune diseases. Since

MADD patients do not suffer from inflammatory myopathies, it is unlikely that the

acquired deficiency of AMPD1 seen in myositis patients is responsible for causing the

infiltration of lymphocytes and the associated autoimmune reaction. Rather, it is more

likely that the AMPD1 deficiency arises in parallel with the inflammatory response. While

the myositis-associated deficiency of AMPD1 is well established, its functional relevance

has remained controversial in humans. Additionally, it is not yet known what mechanisms

are responsible for suppressing AMPD1 expression in idiopathic inflammatory myopathies.

1.4 Role of Innate Immune Signaling in Muscle Inflammation.

Considering that the muscle weakness in myositis patients is often symmetrical despite the

patchy inflammation, and even after successful immunosuppression, it is reasonable to

suspect that a soluble factor is responsible for inducing muscle weakness in a systemic

fashion. Currently there is no consensus regarding which factors may be responsible for

propagating the symptoms of muscle weakness in myositis, and while each most reports

focus on one or two cytokines of interest, no studies have progressed beyond the

descriptive stage. Discussions concerning cytokines often consider only lymphocytes as the

cytokines source, but it is important to consider cytokines that are produced from the

skeletal muscle itself 33,34

. Human skeletal muscle cells constitutively produce the cytokine

IL-6, and its expression can be increased by stimulation with IL-1, IL-1, tumor necrosis

factor-alpha (TNFα) and interferon gamma (IFNγ) in a dose-dependent manner 34,35

. IL-6

9

has a role in regulating metabolic rates in skeletal muscle, but it can also act as an

inflammatory molecule. Excessive secretion of IL-6 has previously been linked to other

autoimmune diseases (e.g. rheumatoid arthritis) and is known to promote the development

of potentially autoreactive TH-17 cells 36,37

. However, the pathogenic potential of IL-6 in

patients is untested, and a large scale survey of cytokine levels in patient serum found that

that IL-6 was significantly overexpressed only patients with dermatomyositis 38

. Other

cytokines, namely IL-8 and IL-15, are also known to be secreted from skeletal muscle and

are sometimes called “myokines” 39

. Despite the large body of literature describing how

these cytokines affect lymphocytes, little is known about how these myokines may

influence muscle development and function in health and disease. For example, IL-15 is

known to stimulate T-cell proliferation by binding to either the IL-2 or IL-15 receptor

complexes, but its effects on skeletal muscle fibers appears to have very different

consequences. Data obtained from mice lacking the IL-15 receptor (specifically the alpha

chain, IL15RA) suggests that IL-15 signaling may significantly alter the composition of

muscles by promoting the formation of fast twitch (type II) muscle fibers. These IL15RA

knockout mice were found to have an above average number of slow twitch (type I) fibers

which coincided with increased the animal‟s endurance during forced treadmill running 40

.

Apart from myokines like IL-6 and IL-15, recent investigations have confirmed that

Type I interferons play a prominent role in the pathogenesis of idiopathic inflammatory

myopathies (IIMs) 41-45

. Type I interferons include IFN and IFN, while IFN is classified

as a Type II interferon. Unlike IL-6 or IL-15, many tissues types will secrete IFN as part

of an innate immune response to various insults. Type I interferon responses are often

10

detected by measuring the increased expression of IFN target genes, the so called

interferon signature. For example, exposing cells to IFN will induce the simultaneous

expression of target genes such as Mx1, ISG15, and IFIT2. This pattern is also found in

patients with myositis, as strong intramuscular staining for these IFN targets has

previously been observed in biopsies from DM and PM patients 44

. An examination of

serum from a large sample of myositis patients showed that IFNwas significantly

upregulated in both DM and PM patients 38

. The clinical significance of the Type I

interferon response was underscored in a small study in which IIM patients failed to

respond to Infliximab, and a worsening in symptoms was correlated with an increase in

IFN serum levels 41

.

Several non-traditional cytokines/chemokines have also been associated with the

onset of myositis in patients, although only one such protein, high mobility group box 1

(HMGB1), has previously been demonstrated to have inflammatory properties 46-48

. The

non-histone nuclear protein HMGB1 acts as a potent inflammatory molecule when released

from necrotic cells and appears to signal through the Toll-like receptor 4 (TLR4) protein.

Examinations of human muscle biopsies have revealed that cytoplasmic HMGB1

expression is widespread in PM and DM patients but absent from healthy controls.

Furthermore, when isolated primary muscles fibers are exposed to extracellular HMGB1 in

vitro, the fibers begin expressing major histocompatibility complex (MHC) class I antigens

on their surface and demonstrate an accelerated, dose-dependent efflux of calcium ions,

suggesting a role for HMGB1 in the perpetuation of inflammation and muscle fiber

degeneration even in the absence of infiltrating lymphocytes 49

.

11

1.5 The Inducible Transgenic Mouse Model of Myositis.

The work presented in this thesis is largely drawn from experiments performed in the

transgenic mouse model of myositis. These transgenic mice, which develop myositis, are

routinely referred to as HT mice throughout this dissertation. First developed and described

by Nagaraju et al. in 2000 50

the transgenic mouse model of myositis recapitulates several

unique features of the human myositis. As mentioned previously, the widespread and

persistent expression of MHC class I molecules within the skeletal muscle is one of the

defining, and most consistent, characteristics of myositis in patients 14,20,51,52

. Importantly,

healthy skeletal muscle fibers are one of the few cell types that express little or no MHC

class I molecules. Transient expression can be observed following physical damage or

exposure to inflammatory cytokines, but this eventually resolves. In contrast, patients with

myositis are suspected to experience sustained expression of MHC class I on the surface of

the skeletal muscle, and therefore the transgenic myositis mice were engineered to

conditionally overexpress MHC class I only in striated skeletal muscle cells 53

.

The inducible transgenic myositis mouse model is a disease model that requires two

components; a transgene and a tissue specific inducible regulator. Specifically, double

transgenic mice possess at least one copy of a tetracycline sensitive transactivator (tTA)

under the control of the muscle specific creatine kinase promoter (hereafter referred to as

the T gene) and at least one copy of an MHC class I gene (H-2Kb haplotype) under the

control of an octet of tetracycline response elements (hereafter referred to as the H gene).

The myositis mice are bred in a C57B6/j background, and the H-2Kb haplotype is

syngeneic to C57B6 mice. Doxycycline is used to suppress the activity of the T gene and

12

prevent the overexpression of the H gene (Tet-off system). Once doxycycline is removed

from the animals‟ drinking water, animals gradually begin to overexpress the MHC class I

gene specifically in skeletal muscle. While individual animals show different rates of

disease progression, male mice are more likely to have late onset and mild disease

symptoms. Female myositis mice reliably show visible disease symptoms around 10 weeks

after the withdrawal of doxycycline.

When the transgenic myositis mouse strain was first characterized, it was found to

exhibit many of the unique features seen myositis. Both patients and mice with myositis

experience muscle weakness, and both show the characteristic histological signs of

myositis such as muscle fiber degeneration and lymphocytes infiltration. Multiple

lymphocytes types are seen in the muscle tissue of HT mice; primarily macrophages, T

cells, and B cells, although there is some evidence of dendritic cells among the infiltrate.

However, the most striking similarity between patients and mice in this model of myositis

comes from the appearance of disease-specific autoantibodies in both patients and myositis

mice. For example, one of the diagnostic markers of polymyositis is presence of

autoantibodies that target the cytoplasmic histidyl-tRNA synthetase enzyme, termed “anti-

Jo-1 autoantibodies”. There are in fact several such autoantibodies like anti-Jo-1 (e.g. anti-

Mi2, anti-SRP) that are observed only in the idiopathic inflammatory myopathies.

Importantly, antibodies from the serum myositis mice, but not control littermates, have

been observed to bind histidyl-tRNA synthetase as well. The appearance of such disease-

specific autoantibodies suggests that the transgenic myositis mouse model closely mimics

the pathology of myositis in humans.

13

Chapter 2: Characterization of muscle metabolic defects in myositis mice.

2.1 Role of AMPD1 in the mouse model of myositis.

As discussed previously, patients with idiopathic inflammatory myopathies are routinely

treated with immunosuppressive treatments, but the successful suppression of infiltrating

lymphocytes does not guarantee a recovery in muscle function. This dissociation between

lymphocyte infiltration and muscle weakness was also previously observed in the

transgenic mouse model of myositis. These transgenic myositis mice, or HT mice, were

described in detail in Chapter 1.3. Prior to the initiation of this dissertation research, it was

observed that HT mice treated with prednisone did not show significant improvements in

muscle function (unpublished observations). In follow up experiments, HT mice were

crossed with Rag-/-

mice and the resulting offspring were given dichloromethylene

diphosphonate (to eliminate macrophages), in order to test the contribution of lymphocytes

to muscle weakness in HT mice. Even after the removal of T-cells, B-cells, and

macrophages, the diseased HT/ Rag-/-

mice were significantly weaker than healthy controls,

though not as weak as diseased HT mice (unpublished observations). The appearance of

weakness despite the elimination of these lymphocytes provided strong evidence that a

significant portion of the weakness seen in myositis arises in the skeletal muscle

independent of the action of autoreactive lymphocytes. In response to these results, we

sought possible mechanisms muscle weakness that were dependent on damage caused by

autoreactive lymphocytes and therefore chose to test the hypothesis that an acquired

deficiency of AMPD1 contributes to muscle weakness in myositis. Another important

14

feature of the mouse model of myositis is the absence of any corresponding C34T allele

which has caused such controversy in human studies. Sequencing of the murine AMPD1

gene confirmed that no similar mutation exists in the HT mouse colony, (Figure 2.1), nor

does such a mutation appear in public databases of the murine genome. The mouse gene is

highly homologous to the human protein (93% homology at the protein level). The other

advantage offered by the mouse model is the ability to systematically compare mice before

and after the onset of overt disease symptoms. By comparing the two different time points,

we were able to definitively say whether or not myositis mice acquired AMPD1 deficiency.

Additionally, by utilizing the mouse model of myositis, we were able to test the

mechanistic link between AMPD1 expression and muscle weakness. The results of our

investigations in mice strongly support the hypotheses that myositis patients do acquire a

deficiency of AMPD1 which contributes to muscle weakness.

2.2 An acquired deficiency of AMPD1 and corresponding muscle weakness appear

prior to lymphocyte infiltration.

In the transgenic mouse model of myositis, the disease is induced by the withdrawal of

doxycycline from the drinking water. For all studies, doxycycline was withdrawn from all

mice at 5 weeks of age. Without doxycycline, double transgenic HT mice begin to

overexpress MHC class I in the skeletal muscle tissue, although visible and histological

disease symptoms are not apparent until about 15 weeks of age (roughly 10 weeks without

doxycycline). Histological hallmarks of myositis include infiltration of mononuclear cells

and muscle fiber degeneration. No sign of infiltration or degeneration was present in

healthy control mice (Fig. 2.2A, C) or in pre-infiltrated HT mice at 8 weeks of age (Fig.

15

2.2B). In contrast, 16 week old HT mice showed all of the histological hallmarks of

myositis (Fig. 2.2D) including mononuclear infiltration and extensive muscle fiber

degeneration. Infiltrated mice were also found to have below-average body weights, while

pre-infiltrated mice did not differ from controls (Fig. 2.2E - F). These results show that

lymphocyte infiltration and overt histological damage to skeletal muscle do not occur until

the infiltrated phase of the disease.

16

17

Importantly, we were able to detect muscle weakness in HT mice, but not control mice,

prior to the infiltration stage. Assessing muscle strength is best measured by in vitro force

contraction analysis on isolated muscles. Additionally, force contraction analysis can be

performed separately on predominantly fast-twitch (i.e., EDL) and slow-twitch (i.e., soleus)

muscles. Infiltrated HT mice showed significantly diminished EDL strength in terms of

both maximal force and specific force when compared to controls (Fig. 2.3E, G). When we

assessed the pre-infiltrated mice, we found significant weakness in the EDL (Fig. 2.3A)

despite the otherwise healthy appearance of the muscle histology at this age (Fig. 2.2C).

Conversely, the soleus muscles of pre-infiltrated mice were not weak (Fig. 2.3B,

D), but infiltrated mice showed a significantly reduced level of soleus specific force (Fig.

2.3H). There was a similar drop in the average soleus maximal force in infiltrated mice

versus control mice (Fig. 2.3F), but this decrease was not statistically significant.

HT mice did not show any significant changes in behavior until the infiltrated phase

of the disease. Specifically, infiltrated HT mice showed a significant loss of forelimb grip

strength (p<0.001) and hindlimb grip strength (p<0.01). These same mice were also fell

from the Rotarod more frequently (p<0.05), and stood standing on their back legs less often

(p<0.05) compared to control mice (Fig. 2.4C-F). Taken together, these results suggest that

a measurable loss of strength in the fast-twitch muscles precedes the infiltration of

mononuclear cells or obvious impairments in normal movement and the strength further

deteriorates during the infiltration phase of the disease.

18

19

AMPD1 expression is highest in the fast-twitch fibers (e.g., EDL, gastrocnemius) of

skeletal muscle. When we measured AMPD enzyme activity of the gastrocnemius,

infiltrated HT mice showed a severe decrease (-88.2%) in AMPD activity compared to

control mice (Fig. 2.5C), but the enzymatic activity was not significantly altered in the

slow-twitch soleus muscle (Fig. 2.5D). Pre-infiltrated mice also showed a significant

decrease (-39.7%) in AMPD activity in the gastrocnemius muscle fibers when compared to

the control mice (Fig. 2.5A). This pattern of AMPD enzyme deficiency coincided and

mimicked the pattern of muscle weakness seen in Figure 2.

We next examined the effects of AMDP1 deficiency on downstream metabolites by

quantifying both fumarate and hypoxanthine levels in skeletal muscle lysates. The

relationship between AMPD1 and these metabolites is illustrated in Figure 2.6. Fumarate is

a molecule that can fuel the TCA cycle, and healthy mice were found to have 42.26 ± 2.57

mmol of fumarate per 50 g of soluble protein, whereas infiltrated HT mice (n=10) were

observed to have significantly less fumarate; only 32.11 ± 3.14 mmol (Fig. 2.7A).

Similarly, healthy mice had 27.11 ± 3.51 mmol of hypoxanthine per 50 g of soluble

protein, while infiltrated HT mice had only 16.01 ± 1.52 mmol; a significant decrease (Fig.

2.7B). It is pertinent to note that hypoxanthine and ribose-1P (a metabolite for PPP

metabolism and glycolysis) are generated simultaneously during AMP catabolism (Fig.

2.6). The loss of these metabolites indicates that the loss of AMPD1 activity has a

significant impact on the production of metabolites that can fuel glycolysis and the TCA

cycle during muscle contractions.

20

21

22

We then analyzed AMPD1 expression by quantitative PCR and Western blotting.

An examination of mRNA levels for AMPD1 in pre-infiltrated mice did not show

significant changes in mRNA levels for AMPD1 (data not shown), but infiltrated mice

showed significant reduction (p<0.001) in AMPD1 expression in fast-twitch muscle (Fig.

2.8A) and a moderate but significant reduction (p<0.05) in slow-twitch muscle (Fig 2.8B).

A similar loss of AMPD1 protein in the fast-twitch muscle of infiltrated mice was observed

by Western blotting (Fig. 2.8C).

It was also important to verify that the loss of AMPD1 was specific to myositis, and

not a general feature of all muscle diseases. Therefore, we assessed AMPD activity in the

mouse model of muscular dystrophy (the mdx mouse) and the mouse model of limb-girdle

muscular dystrophy 2B (the SJL mouse) (Fig. 2.9). The results indicated that there was no

significant difference in AMPD activity between the control C57/BL10, mdx, or SJL mice

in either the fast-twitch (gastrocnemius) or slow-twitch (soleus) muscle fibers. These

results support the conclusion that the loss of AMPD activity is specific to myositis.

2.3 Fiber type switching occurs during the disease in the mouse model of myositis.

Since AMDP1 is expressed preferentially in type II (fast-twitch) muscle fibers, the results

seen in Section 2.2 could potentially be the result of fiber type conversion. In order to

address this possible confounding factor, we quantified the number of type I (slow) fibers

in the EDL and subjected the EDL to twitch parameter analysis. Quantification of slow-

twitch muscle fibers was performed by staining with antibodies that recognize either the

fast or slow myosin heavy chain protein. The results for pre-infiltrated mice showed no

23

significant difference from control values in terms of the percentage of MHC-positive

fibers within either the fast-twitch (Fig. 2.10A) or the slow-twitch muscle (Fig. 2.10B).

However, infiltrated mice showed a significant increase in the number of slow MHC-

positive fibers in both the gastrocnemius (Fig. 2.10C) and soleus (Fig. 2.10D).

In order to verify the functional consequences of fiber type conversion, we

performed twitch parameter analysis in the range of 50-200 Hz on EDL and soleus muscles

from infiltrated mice. The EDL muscles showed a significant reduction in the kinetics of

peak force (39% weaker) and dtpt values (44% lower) when compared to the control mouse

EDL muscles (Table 1). There was also a significant drop in peak force (29% weaker) for

the soleus muscle when compared to that in control mice. Neither the EDL nor the soleus

showed changes in the recovery half-rate time. Overall, the data are consistent with a loss

of type II fibers in the EDL in the infiltration stage, as shown in Figure 2.10. Importantly,

no significant changes in fiber type proportions are seen in the pre-infiltrated mice.

2.4 Decreasing AMPD1 protein expression results in muscle weakness

Next, we tested the hypothesis that knocking down AMPD1 levels would induce muscle

weakness in healthy mice. Vivo-morpholinos were used to knock down AMPD1

expression in vivo. Vivo-morpholinos resemble siRNA oligomers in some ways, but in

morpholinos, the phosphodiester bond is replaced by phosphorodiamidate linkage and the

ribose backbone replaced by a morpholino ring. Due to this altered chemistry, morpholinos

cannot be degraded by any extant nucleases, and are incapable of inducing a PKR-mediated

innate immune reaction. However, morpholinos retain the ability to base pair with nucleic

acids and can block protein translation or interfere with pre-mRNA splicing 54

. The Vivo-

24

morpholino we used was targeted to occlude a 24-bp region centered around the start codon

of the AMPD1 mRNA in order to block ribosome assembly and protein translation. This

inhibition did not affect mRNA levels but did lower protein levels of the targeted gene.

25

26

27

Healthy C57BL/6 mice were injected intravenously with either control (Ctrl mo) or

AMPD1-targeting morpholinos (moAMPD1). Western blot analysis revealed that

moAMPD1 was able to partially block protein translation in the quadriceps, while the

control mice showed normal levels of AMPD1 (Fig 2.11A). Densitometry analysis of

confirmed that the moAMPD1 knockdown resulted in a significant drop in AMPD1 protein

levels (Fig 2.11B). The amount of inhibition achieved was sufficient to induce weakness in

the soleus muscle (Fig. 2.11E), where basal AMPD1 activity is low. Conversely, the EDL

muscles (where AMPD1 is abundant) were not significantly weakened by this treatment

(Fig 2.11A, B).

2.5 Examination of muscle function in AMPD1 knockout mice

We also attempted to test the functional contribution of AMPD1 to muscle strength

by obtaining and examining AMPD1-/-

mice. While an acquired loss of AMPD1 would be

best modeled by the previously described in vivo knockdown experiments, we anticipated

that Vivo-morpholinos may not be sufficient to achieve >90% knockdown of protein

levels. Therefore we sought to confirm and expand upon our knockdown experiments by

examining mice that lacked AMPD1 from birth.

While we were able to obtain AMPD1+/-

mice, we determined that AMPD1 null

mice were not viable after three generations of breeding failed to produce any AMPD1-/-

offspring. Nevertheless, mice which were heterozygous for the AMPD1 gene were found to

possess significantly lower levels of AMPD1 protein compared to wild type littermates

(Fig. 2.12). When the muscle function of 12 week old mice was assessed by in vitro force

contraction, the AMPD1+/-

mice were observed to possess a pattern of weakness which

28

resembled the weakness seen in mice given Vivo-morpholinos against AMPD1 (Fig. 2.13).

Specifically, the only observed differences between wild type mice and AMPD1+/-

mice

was seen in the soleus. No significant changes were seen in the function of the EDL

between wild type and heterozygous mice at 12 weeks of age (data not shown). The

maximal force for the soleus, as well as the weight of the soleus, was on average lower

when compared to control littermates (Fig. 2.13A – B). The specific force for the soleus did

not appear to differ between wild type and heterozygous mice (Fig. 2.13C). However, this

same pattern of changes in the soleus mimics the results from the injection of anti AMPD1

vivo-morpholinos into healthy mice (compare Fig. 2.13A – C with 2.13 D – F).

2.6 Genomic and proteomic examinations of the HT mouse model of myositis

Given our observation that AMPD1 enzymatic activity decreased significantly before the

onset of overt symptoms, we chose to examine the mRNA profile of pre-infiltrated mice

and infiltrated mice using a gene expression array. We later followed up on our expression

array data by taking advantage of an existing proteomic project in our laboratory to

quantify differences in protein levels between H mice and HT mice.

29

30

31

32

33

Expression array data from the pre-infiltrated mice showed no detectable levels of

mRNA for CD3zeta, CD3epsilon, CD4, CD8, CD20, elastase, CD83, or CD94, suggesting

that the muscle tissue from pre-infiltrated mice contained no significant populations of T

cells, B cells, neutrophils, or NK cells. In contrast, expression array data from infiltrated

mice showed a significant increase in levels of CD3 and CD8 transcripts in the skeletal

muscle of infiltrated mice. Macrophage-restricted genes (e.g. EMR1, LGALS3) were

detected in both in all muscle tissue samples, although HT mice showed increased levels of

macrophage transcripts compared to healthy age matched controls. Nevertheless, the

absence of lymphocyte-specific transcripts is consistent with our histological analyses (Fig.

2.2) and supports the hypothesis that significant muscle weakness seen in pre-infiltrated

mice is not a result of the action of autoreactive lymphocytes.

An examination of the genes required for both glycolysis and oxidative

phosphorylation revealed that there was a moderate but significant downregulation of these

two energy production pathways in both pre-infiltrated and infiltrated mice. Specifically,

we found a significant (-fold change <-1.5; p < 0.05) decrease in the expression of

transcripts for many muscle-specific transcripts including AMPD1, muscle

phosphofructokinase (PFKM), glycogen phosphorylase (PYGM), aldolase (ALDOA),

phosphoglycerate mutase (PGAM2), enolase (ENO3), pyruvate kinase (PKM2), and lactate

dehydrogenase (LDHA). Known mitochondrial enzymes were also significantly

downregulated, but very few transcripts for mitochondrial enzymes exceeded the (-fold

change <-1.5) threshold. Cumulatively, these changes in mRNA regulation are consistent

with other observations that a metabolic deficiency exists in pre-infiltrated muscles.

34

As mentioned previously, our lab undertook a large program, called the SILAM

project, to generate tissues that would allow for the quantification of differences between

large sets of proteins identified via mass spectroscopy. SILAM (an acronym for Stable

Isotope Labeling in Mice) tissues were dissected from C57Bl6 mice that had incorporated

13C

into >94% of all lysine residues on every protein. By comparing muscle lysates from

age matched H mice and HT mice to muscle lysates from SILAM animals we confirmed

that AMPD1 was abnormally low in infiltrated HT mice, with only 23.1% remaining in

diseased mice compared to healthy controls. Table 2 shows a list of other enzymes related

to glycolysis were found to significantly diminished, including ENO3, ALDOA, PYGM,

PFKM, and LDHA. This drop in protein levels coincided with a similar decrease in mRNA

levels for these proteins, but few other genes showed positive correlations between changes

in mRNA and protein levels. No significant changes were seen in mitochondrial related

proteins, while the most dramatically upregulated proteins were associated with ER stress

and protein ubiquitination (Table 3). Overall, this data reinforced the conclusion the onset

of myositis coincides with a dramatic loss of AMPD1 protein, and accompanying changes

in other enzymes related to glycolysis.

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36

2.7 Conclusions and Discussion

This chapter focuses on the AMPD1 enzyme as a potential mechanism underlying

muscle weakness in the absence of lymphocyte-mediated inflammation. Foremost among

them is the fact that an acquired deficiency of AMPD1 may provide a mechanistic

explanation for the observed muscle weakness that persists even after successful

immunosuppression. However, this potential mechanism is a very complicated task in

humans, but had never before been attempted in mice. The presented results support the

hypothesis that an acquired AMPD1 deficiency could contribute to the muscle weakness

seen in myositis. The data obtained from our enzymatic assays and gene expression array

also indicated that muscle weakness begins early in the pathologic process.

This work also provides evidence that an acquired deficiency of AMPD1 results in

significant muscle weakness. Force contraction analysis revealed weakness in both type I

and type II fibers, although type II muscles (e.g., EDL) suffer worse weakness and

degeneration than do type I fibers (e.g., soleus) in mice. Importantly, the signs of weakness

can be detected during the pre-infiltrated stage of the disease. Healthy mice have a very

tight distribution of values for both EDL maximal force and specific force, and the onset of

disease causes a significant loss of muscle strength in HT mice. However, the progression

of myositis is not uniform even between littermates, leading to a wider distribution of

values for muscle strength in HT mice. Nevertheless, comparisons between pre-infiltrated

and infiltrated mice make it clear that muscle weakness in the HT mice is detectable even

before the appearance of infiltrating lymphocytes and other histological signs of disease.

To date, no other published findings have described any attempts at in vivo

knockdown in mice in order to verify the mechanistic link between AMPD1 and muscle

37

function. We used systemic administration of Vivo-morpholinos to specifically block the

translation of AMPD1. AMPD activity is more than 10 times higher in fast-twitch muscles

(EDL, quadriceps) than in slow-twitch muscles (soleus). Consequently, the degree of

knockdown achieved was sufficient to produce observable weakness in slow-twitch

muscles because the enzyme is expressed at lower levels in these muscles. When we

examined mice heterozygous for the AMPD1 gene, we observed a similar pattern of

weakness, although the differences between AMPD1+/+

and AMPD1+/-

mice were not

statistically significant. However, it is not appropriate to directly compare a congenital loss

of AMPD1, and an acute loss of AMPD1. Future examinations of AMPD1+/-

mice will

need to test for the possible compensatory upregulation of AMPD3. Based on our data, we

anticipate that an 80-90% knockdown of AMPD1 could have produced weakness in the

EDL. Unfortunately, higher doses of Vivo-morpholinos are known to be toxic in vivo and

could not be tested. Our data demonstrate proof of concept evidence that a transient

knockdown of AMPD1 in wild type mice can cause muscle weakness and support the

hypothesis that an acquired deficiency of AMPD1 can result in muscle weakness.

Previous studies have demonstrated that the type II (fast) fibers are more prone to

inflammation-induced degeneration or atrophy than are type I (slow) muscle fibers 55,56

.

We also found that the proportion of slow-twitch fibers was increased in the EDL of

infiltrated mice. Our twitch parameter analysis corroborated the finding that the EDL in HT

mice does not behave like a typical fast-twitch muscle. The most telling change was

observed in the dpdt value, which describes the ascending slope of the muscle twitch. The

lower dpdt values in HT mice indicated a flattened curve (more like a slow-twitch muscle)

when compared to the sharp peak seen EDL muscles from healthy controls. The values for

38

TPT are a less reliable measure of fiber type composition, since TPT values can be

influenced by a lower peak force (Pt) and, indeed, the Pt values for the EDL and soleus

were reduced in infiltrated mice. Overall, these results support the conclusion that the

skeletal muscle tissue becomes weaker and takes on some of the characteristics of slow

muscle fibers.

Perhaps the most novel finding discussed in this chapter was that AMPD1 enzyme

activity is significantly decreased in pre-infiltrated mice. While it has been reported that

AMPD1 levels are decreased in human myositis patients, there are no data concerning

weakness occurring prior to lymphocyte infiltration in human muscle, leaving some

researchers to question whether the AMPD1 deficiency is congenital or acquired during

disease onset. Unlike humans, our mouse model has no known mutation in the AMPD1

gene, and we were able to examine the mice both before and after the onset of

inflammation. Our data show that AMPD1 levels normally increase with age, but the onset

of myositis causes a loss of AMPD1 activity, especially in type II fibers. Furthermore,

enzymatic data show that a significant decrease in AMPD activity can be detected in the

pre-infiltrated phase of the disease. The decline in AMPD enzyme activity prior to

lymphocyte infiltration strongly suggests that the overexpression of MHC class I in skeletal

muscle contributes to a non-immune-mediated pathology within the muscle fibers.

Currently, the initiating factors that drive the early expression of MHC class I

molecules and inflammation in humans are still unknown. Earlier work by our group has

revealed that in the HT mice, the surface expression of MHC class I molecules also

coincides with increased expression of markers for ER stress, lysosome formation, and

autophagy 57-60

. These previous results suggest that ER stress and the unfolded protein

39

response play a part in the non-immune-mediated degeneration muscle fibers in myositis.

We set out to gain insight into this self-sustaining pathology by examining pre-infiltrated

muscle tissue from HT mice with a gene expression array. The results of our analysis

indicated that AMPD1 was significantly down-regulated prior to the infiltration of

lymphocytes, as were its downstream metabolites. Also, the expression array analysis

revealed a global deregulation of energy production in the muscle fibers. These findings are

consistent with prior evidence of the disruption of glycolysis in human myositis patients 61

.

In summary, the results of this research support the hypothesis that decreased

AMPD1 activity contributes to muscle weakness in inflammatory myopathies. These

results further support the hypothesis that the loss of muscle strength can be due to an

acquired loss of AMPD1, rather than simply due to autoimmune muscle damage. We also

presented evidence that the energy production capacity of the skeletal muscle cells may be

diminished in inflammatory myopathies. Overall, these data support the contention that a

non-immune-mediated pathology could be responsible for the muscle weakness in these

patients and that targeting these pathways may have therapeutic value.

40

Chapter 3: Effects of D-ribose treatment on myositis mice

3.1 Effect of D-ribose treatment on disease phenotype in myositis.

As discussed in the previous chapters, the loss of AMPD1 results in a decrease in its

downstream metabolites. One of the downstream metabolites, ribose-1‟monophosphate

(ribose-1P) can potentially be used to fuel glycolysis during intense exercise. Based on case

reports from patients with a congenital loss of AMPD1, it has been suggested the related

compound D-ribose may be able to alleviate the symptoms of muscle weakness, fatigue,

and cramping in myositis 62,63

. We have therefore chosen to utilize the transgenic mouse

model of myositis in order to assess the therapeutic potential of D-ribose in treating the

symptoms of myositis. This work is also relevant because D-ribose is advertised by some

patients and vitamin manufacturers as a treatment for muscle weakness.

It has been proposed that D-ribose could ameliorate muscle weakness by serving as

an energy source in previously published case reports, but there is little research, which

specifically addresses this topic. While it is known that D-ribose is well absorbed in the

small intestine 64

and that pentose sugars can be converted into hexose sugars, most texts

focus on reverse reaction whereby glucose is converted into ribose. However, skeletal

muscle cells have unique requirements in terms of energy production, and there is evidence

in the literature to suggest that type I skeletal fibers can catabolize AMP molecules in order

to create free ribose as a potential energy source. The strongest evidence supporting the

catabolism of AMP comes from observations of an „adenylate deficit‟ following strenuous

exercise, and a spike in serum levels of hypoxanthine following strenuous exercise 65

. In

brief, it is known that the total pool of adenylate molecules (i.e., ATP+ADP+AMP) in

41

skeletal muscle tissue is significantly decreased after exercise, but recovers over time. The

drop in number of adenylates coincides with a sharp increase in serum levels of

hypoxanthine (a breakdown product of AMP) suggesting that excess AMP is broken down

into its purine base (hypoxanthine) and phospho-ribose backbone (ribose-1P) after exercise.

An excess of free ribose-1P can then be used to fuel glycolysis, as three ribose-1P

molecules can be converted into two molecules of fructose-6P and one molecule of

glyceraldehyde-3P, via the non-oxidative phase of the pentose phosphate pathway.

3.2 An acquired loss of AMPD1 correlates with a loss of downstream metabolites

A plausible metabolic pathway linking AMPD1 enzyme activity to the production of ribose

(Fig. 3.1) was constructed based on literature searches and previously generated microarray

data from myositis mice (previously discussed in Chapter 2). In order to verify that this

pathway was affected by the acquired loss of AMPD1, we quantified the relative

abundance of the metabolites inosine monophosphate (IMP) and hypoxanthine (Figure

3.2). Quadriceps muscles from 16 week old H and HT mice were dissected, flash frozen,

and then homogenized to generate skeletal muscle lysates. Inosine monophosphate is the

immediate downstream product of AMPD1, and it was observed that IMP levels in

myositis mice were 24.6% lower compared to healthy controls.

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43

44

A more severe deficit was seen in hypoxanthine, where the lysates from skeletal muscle

from myositis mice held 58.2% less hypoxanthine (7.17±0.49 M, n = 10) compared to

healthy controls (17.19±1.72 M, n = 10). Hypoxanthine is produced simultaneously with

ribose-1P at a 1:1 ratio and therefore serves as a detectable indicator for the amount of free

ribose-1P that is produced within the skeletal muscle fibers. This data is consistent with the

findings previously shown in Chapter 2. The results support the hypothesis that an acquired

AMPD1 deficiency results in a significant loss of downstream metabolites.

3.3 Treatment with D-ribose fails to improve activity in myositis mice.

Mice were treated with oral supplements of D-ribose according to the schedule depicted in

Fig. 3.1B. The withdrawal of doxycycline at 5 weeks of age marks the beginning of the

onset of myositis in HT mice, while H mice remain healthy. Treatment with D-ribose

began when mice were 10 weeks old, and continued until mice were 16 weeks old. Four

groups in total were used in the trial: untreated healthy mice (H), healthy mice given daily

oral D-ribose (H + Rib), untreated myositis mice (HT), and myositis mice treated with

daily oral D-ribose (HT + Rib).

When mice were 15 weeks of age, they were subjected to five days of behavioral

assessments. Measurements such as mouse grip strength and mouse movement are

considered behavioral assay because the results can vary due to subjective influences such

as the animal‟s willingness to participate or expectation of being tested. The H mice

showed normal grip strength, regardless of treatment status, for both the forelimbs (Fig.

3.3B) and the hindlimbs (Fig. 3.3C). In comparison, the HT myositis mice appeared weaker

45

compared to the healthy controls in forelimb strength (compare the first and third groups in

Figure 3.3B). Measurements of the hindlimb strength varied too widely to discriminate

between healthy and diseased mice (Fig. 3.3C). Treatment with ribose did not have any

effect on grip strength for myositis mice (compare the third and fourth groups of Fig. 3.3A

& B).

Body weights were recorded when the mice were 16 weeks of age, and the results

are shown in Figure 3.3A. The onset of myositis resulted in a significant decrease in body

weight when untreated H mice are compared to untreated HT myositis mice. Treatment

with D-ribose resulted in a downward trend in bodyweight, although the decrease in

weights did not reach statistical significance for either two treated groups of mice.

We were also able to record behavioral activity using a Digi-Scan open-field

recorder, wherein the position of each mouse is tracked in three dimensions using a grid of

infrared beams. Untreated healthy mice covered a large distance (Fig. 3.4A), and showed

robust horizontal (Fig. 3.4B) and vertical movements (Fig. 3.4C). Untreated HT mice

moved very little by any measure, consistent with their advanced stage of myositis.

Additionally, treatment with daily oral D-ribose showed no benefit to HT myositis mice

treated with D-ribose when compared to untreated myositis mice.

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47

48

3.4 Histology of skeletal muscle showed no differences between treated and untreated

mice. Histological analysis was performed on sections of muscle tissue to examine the

degree of inflammation within the mice. While D-ribose was anticipated to improve the

symptoms of myositis, there was some concern that D-ribose could stimulate greater

proliferation of autoreactive lymphocytes. D-ribose is also utilized in the de novo synthesis

of all nucleotides and can act as a limiting factor for DNA synthesis and mitosis. For

hematoxylin and eosin (H&E) staining, quadriceps muscles from H and HT mice were

dissected, flash frozen, sectioned, and stained with H&E (Figures 3.5A - D). The images

shown are representative of the histology in each treatment group. The untreated H mice

(Fig. 3.5A) possessed normal muscle histology, while untreated HT myositis mice (Fig.

3.5C) showed the expected severe inflammation and muscle fiber degeneration. A

comparison between Figure 3.5A and Figure 3.5B shows no change in muscle histology in

treated and untreated H mice, respectively. Likewise, a comparison between Figure 3.5C

and Figure 3.5D shows that treatment with D-ribose had no apparent effect on the degree of

inflammation and muscle fiber degeneration.

3.5 Treatment with oral D-ribose did not improve muscle function.

Muscle wasting (accompanied by inflammation and fiber degeneration) is generally worse

in muscles that possess a large proportion of fast twitch (i.e. type II) fibers. Figure 3.6A

shows a loss in muscle mass in the EDL (which is primarily composed of type II fibers) in

HT mice compared to H controls (compare the first and third group in Fig.3.6A), but

treatment with daily oral D-ribose had no significant protective effect on muscle mass. In

order to obtain more reliable data on the muscle function of treated and untreated mice, we

49

performed force contraction analysis on freshly dissected individual muscles. Force

contraction analysis is performed on recently dissected muscles under controlled conditions

and is therefore free from the subjective bias inherent in grip strength and voluntary motion

measurements. Using force contraction analysis, we were able to accurately determine the

maximal force and specific force generated by individual EDL in 16 week old mice.

Consistent with the disease pathology of myositis, the EDL muscle from untreated HT

mice showed significant decreases in performance for both maximum force generation

(Fig. 3.6B) and for specific force generation (Fig. 3.6C). Treatment with oral D-ribose had

no significant effect on the performance of the EDL.

Force contraction analysis was also performed on freshly dissected soleus muscles

from 16 week old mice. The soleus muscle differs from the EDL in its composition, as the

soleus is composed almost entirely of slow twitch (i.e. type I) muscle fibers. Consequently,

the soleus is not as severely damaged as the disease progresses. Figure 3.7A shows that the

HT myositis mice do not experience muscle wasting in the soleus, but rather experience a

slight increase in muscle mass (compare the first and third groups in Fig. 7A). Untreated

myositis mice showed weaker soleus muscles compared to H controls, both in terms of

maximal force (Fig. 3.7B) and specific force (Fig. 3.7C), although not as severe as the

weakness seen in the EDL. However, treatment with oral D-ribose had no significant effect

on soleus muscle function.

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52

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3.6 Effects of D-ribose treatment on muscle fatigue resistance.

The EDL muscles were additionally tested for fatigue resistance after completing force

contraction analysis. Fatigue resistance is measured as a ratio between the maximal force

production and the force production remaining after 60 consecutive contractions. The data

for fatigue resistance is displayed in Figure 3.8. The EDL muscles from untreated H mice

fatigued rapidly as expected. In contrast, untreated HT myositis mice showed a varying

amount of fatigue resistance (compare the first and third columns of Fig. 3.8A). The fatigue

resistance seen in untreated HT mice has previously been observed 60

. Treatment with daily

oral D-ribose did not have a statistically significant effect in either H or HT mice, although

it did appear to increase the mean fatigue resistance in both groups (Fig. 3.8A). One

potential pitfall in the fatigue resistance protocol concerns the size of the fiber and the rate

at which metabolites like lactic acid and H+ can diffuse more quickly out of smaller

muscles during the 60 consecutive contractions. This is a particular concern here, since

Figure 3.6A shows a drop in the average muscle mass of the EDL after D-ribose treatment

of myositis mice. In order to account for this experimental artifact, we plotted the EDL

muscle fatigue resistance against EDL cross sectional area for all untreated (H and HT) and

all D-ribose treated mice (H+Rib and HT+Rib), as seen in Figure 3.8B. The linear

regressions formed by the two groups form nearly parallel lines, and it is apparent that the

y-intercept for D-ribose treated animals is larger than the y-intercept for untreated animals.

This result suggests that the small increase in the average fatigue resistance after treatment

is due to the effect of D-ribose, and is not an experimental artifact.

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3.7 Possible explanations for lack of D-ribose efficacy in the mouse model

In order to better explain the results of our trial, we performed QPCR analysis to examine

gene expression levels for each of the relevant enzymes in both treated and untreated mice.

Total RNA was isolated from frozen quadriceps tissue and used to perform quantitative

PCR (QPCR). The expression of each gene was then compared to the expression of the

housekeeping gene GAPDH. We quantified the expression of each of the enzymes shown

on the pathway depicted in Figure 3.1A, as well as other genes important for muscle

metabolism. A portion of the QPCR results are shown in Figure 3.9. The genes for both

AMPD1 and muscle creatine kinase (CKM) are known to be downregulated in humans and

mice with myositis. As mentioned previously, AMPD1 is believed to control AMP levels

in skeletal muscle, while CKM is essential for regulating levels of phosphocreatine. The

data obtained for AMPD1 (Fig. 3.9A) and CKM (Fig. 3.9B) show that both genes are

downregulated in untreated HT mice compared to untreated H mice. Treatment with daily

oral D-ribose had no significant effect on the expression of these genes when comparing

untreated and treated HT mice. QPCR analyses on the expression of 5‟-nucleotidase C2

(NT5C2), purine nucleoside phosphorylase (PNP), and phosphoglucomutase 2 (PGM2)

showed that each of these genes was expressed in the skeletal muscle, but no significant

differences in gene expression were observed between any of the treatment groups (data

not shown). However, QPCR analysis on RBKS and hexokinase 2 (HK2) revealed that the

basal expression of RBKS was only 18.1% that of HK2 expression. Like all simple sugars,

ribose must be phosphorylated in order to prevent its diffusion back into the bloodstream.

As illustrated in Figure 3.1, RBKS is required to phosphorylate D-ribose and convert it into

ribose-1P. The more abundant HK2 enzyme performs the analogous reaction for all hexose

57

sugar molecules. The low expression of RBKS therefore suggests that mouse muscle cells

may not effectively retain and metabolize ingested D-ribose. A comparison of RBKS

expression profiles in humans and mice using publicly available expression array results

through the NLM Unigene database suggests that both humans and mice have either no

expression or very low basal expression of RBKS in the skeletal muscle.

3.8 Discussion

This chapter summarizes effect of oral administration of D-ribose on symptoms of myositis

in mice. While we were able to demonstrate that there is indeed a deficiency of

hypoxanthine and other AMP breakdown products in myositis mice, we found that

treatment of these mice with daily oral doses of D-ribose had no significant effects on mice.

These results are consistent with the wider body of literature stating that D-ribose has no

effect on muscle performance in healthy patients, or patients with other metabolic diseases

66,67. We also proposed that the lack of efficacy is a consequence of very low levels of the

enzyme ribokinase, suggesting that skeletal muscle cells may not be able to retain and

utilize ingested D-ribose. Taken together, these results suggest that oral supplements of D-

ribose have negligible effects on the skeletal muscle tissues, and thus not likely to have any

beneficial effects in patients with myositis.

Myositis is currently treated as an autoimmune disease and patients are typically

treated with immunosuppressive drugs (e.g. prednisone, methotrexate) while metabolic

abnormalities are marginalized. This work was originally proposed based on a combination

of previously published findings. First, it has been observed that both patients and mice

with myositis acquire a deficiency of the muscle specific enzyme AMPD1. Second, there

58

are published case reports describing the successful treatment of muscle pain and fatigue in

patients with a congenital loss of AMPD1. Finally, an examination of the metabolites

within the muscle tissue of myositis mice revealed a deficiency of metabolites related to the

internal production of D-ribose. Taken together, these observations suggested that

supplementing D-ribose to mice with myositis could reverse the symptoms of muscle

weakness and fatigue. Despite the lack of data concerning treatment for myositis, D-ribose

has been touted as a treatment for both myositis and fibromyalgia by patients and vitamin

manufacturers. It should be noted that while D-ribose is known to show no ergogenic

benefits in healthy patients, healthy patients are not expected to have an AMPD1

deficiency. Since patients with myositis are expected to be deficient for AMPD1 enzyme

activity, treatment with D-ribose remained an untested potential therapeutic compound.

However, after treating myositis mice for six weeks with daily oral supplements of

D-ribose, we observed that treated mice failed to show any improvement by any

measurement. Mice with myositis fared poorly compared to healthy littermates, and

treatment with D-ribose had no significant effect on either diseased or healthy mice.

Treatment with D-ribose did not protect against muscle wasting, or prevent an overall loss

in bodyweight. Neither did D-ribose protect against the loss of muscle function (maximal

and specific force generation) in the myositis mice. Histologically, treatment with D-ribose

did not appear to have any effect on the degree of muscle fiber damage or infiltration of

autoreactive lymphocytes.

In fact, a pattern of potential harm becomes apparent after examining bodyweight,

voluntary activity, muscle mass, and muscle function. In all of these analyses, treated mice

performed worse on average than the untreated controls. Although this drop in performance

59

was not statistically significant in any individual comparison, the consistency of this pattern

suggests that daily oral D-ribose has potentially negative consequences. The mechanism

behind this consistent drop in performance is not known. There was some prior concern

that D-ribose supplements would allow greater proliferation of infiltrating lymphocytes,

since ribose is a limiting factor in the synthesis of new DNA in dividing cells. However, the

histology results seen in Figure 3.5 did not show any significant differences in the degree of

intramuscular inflammation between treated and untreated animals. In terms of muscle

fatigue, supplementation with D-ribose had no statistically significant effect on muscle

fatigue, despite our observation that average fatigue resistance was slightly improved in D-

ribose treated animals.

In summary, the results of this research refute the hypothesis that oral supplements

of D-ribose can be used to treat the symptoms of myositis. Contrary to the initial

hypothesis, animals treated with D-ribose showed worse performance on average,

compared to controls. The failure of D-ribose in treating D-ribose does not invalidate the

hypothesis that catabolism of AMPD provides substrates for glycolysis, because ingested

D-ribose requires RBKS to be utilized, whereas the internal production of D-ribose via

AMP catabolism does not require RBKS. These results are also consistent with the broader

literature on D-ribose and its lack of observed effects. However, this research is the first to

suggest that D-ribose has insignificant effects on skeletal muscle due to low basal

expression of RBKS. Overall, these results suggest that patients with myositis will not

experience any benefits from taking D-ribose.

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Chapter 4: The effects of inflammatory cytokines on AMPD1 expression

4.1 Cytokine and chemokine milieu in myositis

A large number of cytokines and chemokines have been proposed to play a role in the

pathogenesis of the various idiopathic inflammatory myopathies. Specifically, the

molecules IL-1, IL-6, IL-15, IL-17, IFN, IFN, CCL2, CCL3, CCL4, CCL5, CXCL10,

CXC3L1, TNFSF13, and VEGF have all been observed to differ significantly between

groups of myositis patients and healthy control subjects. In addition, recent publications

have implicated various TLR receptors in the pathogenesis of myositis 38,43,68-70

.

In light of the fact that the muscle weakness takes on a symmetric pattern despite

patchy inflammation and that some patients continue to experience weakness even after

successful immunosuppression, we have hypothesized that the symptoms of muscle

weakness arise due to the effect of signaling molecules secreted by the innate immune

system, or by the muscle tissue itself. This hypothesis is supported by findings such as

prominent staining for IFN within muscle fibers from myositis patients, but not healthy

patients. Results from our earlier microarray experiment indicate that diseased HT mice

also show a strong Type I interferon signature that is absent in H mice, as shown in Table

4. Several Type I interferon targets are significantly upregulated, although the nearly all

Type I interferon target genes are more highly upregulated during the infiltration phase of

the disease. Other reports show TLR3 and TLR7 were detectable on the surface of muscle

fibers from myositis patients, but not in healthy controls 70,71

. Specifically, analyses of

skeletal muscle tissue from HT mice showed significant activation of NF- and an

61

upregulation of its downstream targets 57

. However, the cytokine(s) responsible for

inducing NF- activation in myositis mice have not been definitively identified.

62

There is a growing body of literature, which has implicated the Type I interferons response

in the pathogenesis of inflammatory myopathies. The Type 1 interferon response has been

observed to be strongest in patients with DM and PM, while patients with IBM are not

expected to show an upregulation of Type I interferon targets. The Type 1 interferon

response is mediated by both IFN and IFN. The IFN cytokine has many isoforms (the

result of multiple gene duplications), secreted by primarily by lymphocytes. In contrast, the

singular IFN cytokine is secreted by a wide range of tissues and was initially called

“fibroblast derived interferon”. Both IFN and IFN share the same receptor (a

heterodimer composed of IFNAR1 and IFNAR2). Both cytokines have been observed to

target the same genes for upregulation, although the various isoforms of IFN are known

to have variable potency. Interestingly, IFN has been described to have one unique

function in controlling the fate of osteoblasts, prompting speculation that the calcifications

seen in some myositis patients may be due to chronic IFN secretion. To date, there are

two open trials attempting to treat polymyositis and dermatomyositis by blocking Type I

interferon signaling. These two trials focus on the use of Rituximab to indirectly target

Type I interferon signaling, and the use of Medi-545 (an anti-IFN monoclonal antibody)

to directly block IFN signaling. The use of Rituximab to treat the various forms of

myositis is controversial as the results of several case reports have found inconsistent

benefits from Rituximab in patients with myositis and other autoimmune disorders 72-75

.

63

64

65

4.2 Cytokines, Chemokines and TLRs in Myositis Mouse Muscle

With the transgenic mouse model of myositis, we attempted to focus on inflammatory

cytokines or TLR receptors that are significantly different or unique in the skeletal muscle

tissue from diseased HT mice when compared to control littermate H mice. We first

utilized a gene expression microarray in order to identify mRNA transcripts within the

skeletal muscle tissue corresponding to each cytokine and its matching receptor. The

mRNA expression data is shown in Table 5. With this approach, we identified IL-1,

CCL5, CXC3L1, and TNFSF13 as significantly upregulated cytokines, and IL-15 as

significantly downregulated in HT mice. The receptors CCR5, CX3CR1, TNFRSF13 were

all found to be upregulated, while the IL15RA receptor was found to be downregulated and

the IFNAR1/2 genes were unchanged between HT and H mice. Additionally, TLR2, TLR4,

TLR6, and TLR7 were all found to be upregulated in muscle tissue from HT mice, but

barely detectable in H mice. Other cytokines and receptors, like IFN and IL1R1, showed

no significant changes in transcript levels between H and HT mice. A few cytokines, such

as CXCL10, were excluded from further experiments due to the absence of transcripts for

its receptor, CCR3, in the muscle tissue.

4.3 In vitro reporter assay to screen modulators of AMPD1 expression

In order to investigate the potential mechanisms linking cytokine signaling and the

expression of AMPD1, a luciferase reporter construct driven by the murine AMPD1

promoter, named pGL3-AMPD1, was obtained from Dr. Judith K. Davie. The construction

of the pGL3-AMPD1 plasmid has been described previously 76

, but in brief, the reporter

66

contains the AMPD1 promoter covering bases -359 to +74 controlling the expression of the

firefly luciferase gene as depicted in Figure 4.1B. The majority of the 5‟UTR of the

AMPD1 gene (stopping short of the ATG start codon located at +82) is included in this

reporter construct, and public genomic databases indicate that the upstream genomic

conserved regions of the promoter fall between base -170 and 0. Despite the relatively

compact size of the AMPD1 promoter there have been very few prior analyses of the

promoter, although there is a known canonical MEF2 binding sequence centered roughly at

base -102, which is the presumed site of action for the MEF2, MyoD and MyoG

transcription factors. The effect of inflammatory innate cytokines on the AMPD1 promoter

has not previously been analyzed. We performed transcription binding site prediction using

the Alggen-Promo algorithm with TRANSFAC v8.3 weight matrices to analyze the

promoter 77,78

. The prediction algorithm successfully identified the known MEF2 binding

site in the AMPD1 promoter, and also identified many other potential transcription factor

binding sites, including binding sites for interferon response factor 1 (IRF-1), signal

transducer and activator of transcription 4 (STAT4), and the androgen receptor (AR). The

IRF-1 and STAT4 sites are potential sites of action for Type I interferon signaling, and

their location in the AMPD1 promoter is depicted in Figure 4.1A.

The pGL3-AMPD1 reporter was then used to transfect Immorto SV40-A58T

myoblast cells followed by treatment with the candidate cytokines which were previously

singled out in Table 4. Immorto SV40-A58T myoblast cells were chosen because these

cells are clonal derivatives of a myoblast stem cell isolated from a transgenic mouse strain

which carries an inducible and reversible oncogene. These cells can therefore be cultured

much longer than primary cells, but retain the ability to fuse and mature into myotubes

67

under permissible conditions 79

. The results of our initial cytokine screening are presented

in Figure 4.2. Normal expression from the AMPD1 promoter was measured using

transfected cells without any further treatment. We were able to test a majority of the

candidate cytokines, although there was no readily available source for some such as

CXC3L1 or TNFSF13.

68

The available cytokines were used in varying doses in order to reflect the maximum

physiological concentration based on prior investigations of cytokine levels in the serum of

myositis patients 38,80

. Agonists of the TLR receptors were also tested. Specifically, the

TLR2 agonist PAM3SK4, the TLR4 agonist lipopolysaccharide (LPS), and the TLR7

agonist ORN-06 (an ssRNA oligomer) were utilized. The results from this cytokine screen

indicated that the most potent inhibitors of the AMPD1 in vitro was IFN (reduced to

18.7% of normal activity) and IL-1b (reduced to 54.4% of normal activity), while the most

potent stimulator was IL-15 (increased 288.1% over normal activity). Among the TLR

ligands, the TLR2 agonist PAM3SK4 showed the strongest inhibitory activity (reduced to

54.7% of normal activity).

4.4 Generation of an AMPD1 reporter stable cell line

In order to aid in the future identification of cytokines and compounds that might modulate

AMPD1 activity (and thus impact muscle function), we created a stably transfected

AMPD1 reporter cell line. The stable plasmid construct was composed of the pGL4.5

vector and the same AMPD1 promoter found in the pGL3-AMPD1 plasmid. The resulting

pGL4.15-AMPD1 plasmid was transfected into Immorto SV40-A58T myoblasts and

selected for using Growth Media supplemented with hygromycin-. The resulting stably

transfected cell line was observed to retain the characteristic bipolar/tripolar appearance of

myocytes and the ability to fuse into myotubes after six days in culture with Differentiation

Media. The stably transfected cell line was then verified to respond to cytokine treatments

in the same pattern as previously observed in transiently transfected cells (Fig. 4.3).

69

70

Stably transfected cells showed were found to respond to IFN, IL-1, and IL-15 in a

pattern that was very similar to the results seen in transiently transfected myoblasts

(Compare Fig. 4.3 and 4.2). Specifically, both IFN, IL-1were able to inhibit the

expression of luciferase under the AMPD1 promoter while IL-15 was able to upregulate

the expression of luciferase under the AMPD1 promoter. However, the stably transfected

cells required a greater amount of time (6 days) to show significant differences in

expression compared to transiently transfected cells (3 days). The discrepancy in time may

be due to the fact that stably transfected cells likely possess a single copy of the reporter

construct, while transiently transfected cells may possess multiple copies of the reporter

plasmid.

4.5 Expression of IFNAAR and IL15RA in mice

Based on the results obtained from the in vitro reporter assay, we chose to verify the

presence of receptor proteins for both type I interferons and IL-15 in the skeletal muscle of

both H and HT mice. Both IFN and IFN bind to the heterodimeric receptor

IFNAR1/IFNAR2, while IL-15 binds with high affinity to the IL15RA surface receptor.

Both mice and humans have been reported to possess multiple splice isoforms of the

IL15RA receptor, although only the full length is believed to be functional. The results of

our Western blot analysis are shown in Figure 4.4. The vinculin is used a muscle specific

loading control. Both IL15RA and IFNAR1 were detected in lysates from skeletal muscle

in both H and HT mice. Densitometry was then performed on the Western blot results to

determine whether the observed changes in protein levels were significant. In the case of

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IFNAR1, there was a trend in the average receptor protein level in HT mice compared to

healthy mice, although the difference was not statistically significant. However, it was

observed that the protein levels for IL15RA were significantly reduced in HT animals to

only 78.1% of the levels seen in healthy mice.

4.6 Effect of Type I interferon expression in vivo in mice.

When we combine the results from our in vitro reporter assay, our Western blot analysis,

and previously published reports it is possible to propose a mechanism linking type I

interferons to refractory muscle weakness in myositis patients. While IFN is principally

secreted from lymphocytes, IFN can be secreted from endothelial cells and muscle fibers.

Both of these cytokines act on the same receptor, which is present in mouse muscle tissue.

If Type I interferons can downregulate AMPD1 expression in vivo as well as in vitro, then

the weakness associated with the loss of AMPD1 could be the direct result of Type I

interferons secreted from inflamed muscle fibers even in the absence of infiltrating

lymphocytes.

In order to test this potential mechanistic link between Type I interferons and an

acquired AMPD1 deficiency, were chose to transiently overexpress IFN in the tibialis

anterior (TA) and measure any resulting changes in the function of the extensor digitorum

longus (EDL) muscle, which is adjacent to and in contact with the TA muscle. We were

able to obtain a murine IFN mammalian expression plasmid from Dr. Ron Jubin, along

with an empty control vector. Our in vivo electroporation protocol was successfully tested

by using a GFP expression plasmid in mice, as shown in Figure 4.5. Using this protocol,

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we were able to express GFP in muscle fibers of the TA, but not fibers of the adjacent

EDL.

73

74

75

Histological analysis revealed that that electroporation protocol itself induced a significant

amount of muscle fiber regeneration (visible as central nuclei within individual muscle

fibers) as well as a small amount of inflammation (visible as clusters of lymphocyte nuclei

between muscle fibers), as shown by Figure 4.6A. The same level of regeneration and

inflammation was seen in the TA for injection of both the empty vector DNA and the GFP

expression plasmid. The EDL adjacent to the site of injection was observed to be unharmed

by the electroporation protocol (Fig. 4.6B) as there was no visible inflammation and little to

no muscle fiber regeneration.

In order to test the hypothesis that Type I interferons secreted from muscle fibers

can induce muscle weakness and a loss of AMPD1, we next injected and electroporated the

IFN expression plasmid into the left TA of healthy mice, and the empty control vector

into the contralateral TA. Twelve days after the injection, the muscle function of the

adjacent EDL muscle was assessed by in vitro force contraction, while the TA muscle

tissue was preserved for histology and QPCR analysis.

Histology revealed that the forced expression of IFN resulted in severe

inflammation within the TA near the injection site (Fog. 4.7A) as well as significant

inflammation in the adjacent EDL muscle (Fig. 4.7B). The in vitro force contraction

analysis of the EDL adjacent to the electroporated TA revealed that the expression of IFN

from the TA skeletal muscle was able to induce weakness in the adjacent EDL (Fig. 4.8). In

contrast, the injection of either GFP or an empty vector plasmid into the contralateral TA

had no significant effects on the muscle function of its adjacent EDL.

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In order to verify that IFN was indeed secreted and active within the muscle tissue, we

isolated mRNA from the TA muscle and performed QPCR analysis on several Type I

interferon target genes. Simultaneously, we measured the mRNA levels for AMPD1 to

confirm whether or not IFN secretion from the muscle fibers influenced AMPD1

expression. The two target genes MXB and IFIT2 were chosen since both are known

targets of IFN and both were found to be upregulated in diseased HT mice. The results of

the QPCR analysis are shown in Figure 4.9. The expression of MXB and IFIT2 were

upregulated 7.64 and 7.98-fold, respectively, over baseline levels seen in the contralateral

TA. However, there was no observed drop in AMPD1 expression in the TA following the

expression of IFN. This evidence suggests that Type I interferons may be able to induce

significant muscle weakness and inflammation (both within muscle groups and in adjacent

muscle groups). However, further experiments will be needed to confirm whether or not the

weakness induced by Type I interferons in vivo is dependent upon changes in AMPD1

enzyme activity.

4.7 Conclusions and discussion

This chapter has described the work that has been accomplished to date on identifying and

verifying inflammatory cytokines that have the potential to induce weakness in mice even

in the absence of autoreactive lymphocytes. Among all of the cytokines and TLRs

previously associated with myositis in humans, only a subset of these cytokines were also

found to be significantly expressed in the skeletal muscle tissue of mice with myositis. We

77

chose to focus on these cytokines in order to test our hypothesis that the muscle fibers

themselves were capable of secreting factors capable of inducing muscle weakness.

78

79

Using an AMPD1 reporter construct, we were able to identify two cytokines that had a

profound effect on the expression of AMPD1; IFN and IL-15. These two cytokines

induced opposite effects in vitro, as IFN suppressed AMPD1 expression while IL-15

increased the expression of AMPD1. Surprisingly, the various TLR agonists that were

tested in vitro showed comparatively little ability to influence the expression of AMPD1.

Previous literature reports had suggested possible mechanisms whereby necrotic muscle

fibers induced greater inflammation via HMGB1-TLR4 or ssRNA-TLR7 signaling 69

.

While these interactions may still be capable of inducing inflammation, the focus of this

research was to identify a mechanism that could explain the acquired loss of AMPD1 and

coincident weakness that occurs even prior to the infiltration of lymphocytes.

We were able to verify that receptors for IFN and IL-15 were present within the

mouse skeletal muscle tissue, both before and after the onset of disease, although levels for

the IL15RA receptor are significantly reduced in HT animals. The observed drop in protein

levels for IL15RA is particularly novel, and it allows us to propose a novel pathogenic

mechanism in onset of muscle weakness in myositis. Though much of the IL-15 literature

focuses on its proliferative effect on lymphocytes, IL-15 signaling has recently been

described to have significant effects on the development of type II muscle fibers 40

. IL-15

signaling may have been previously been overlooked by other myositis researchers since

serum levels for IL-15 were not found to be abnormal for DM, PM or IBM patients

(although juvenile cases of DM showed some significant loss of IL-15) 38

. Specifically, we

80

propose that a loss of IL-15 signaling is required to maintain the expression of proteins

characteristic of type II muscle fibers, such as AMPD1.

In the case of Type I interferon signaling, we were able to test the proposed

mechanistic link between IFN, AMPD1 expression, and muscle weakness in vivo. We

were able to demonstrate successful in vivo electroporation with a GFP expression vector,

and were further able to verify successful production of IFN from a separate plasmid by

observing an increase in the expression of Type I interferon target genes. When healthy

mice are forced to overexpress IFN from the muscle fibers, both the TA muscle and the

adjacent EDL muscle experienced significant lymphocyte infiltration and weakness. These

results confirmed our hypothesis that the secretion of IFN is capable of affecting nearby

muscles, although there were no signs that the weakness spread in a systemic or

symmetrical pattern within twelve days of the initial electroporation. This result also lends

strong support to the hypothesis that Type I interferon signaling is a significant source of

pathology in myositis patients. Unfortunately, the forced expression of IFN from the

muscle fibers did not appear to have any effect on the transcription of AMPD1 within the

TA muscle. Due to practical constraints, it was not possible to either directly measure

muscle strength in the TA, or to quantitatively measure AMPD1 enzyme activity in the

EDL. Until such experiments can be completed, we cannot definitively conclude whether

or not the weakness induced by Type I interferons is dependent upon changes in AMPD1

levels.

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Chapter 5: Conclusions and future directions

This dissertation research can be divided into three separate aims. The first aim established

an important mechanism of persistent muscle weakness in the transgenic mouse model of

idiopathic inflammatory myositis. The second and third aims were derived from the results

of the first aim, but address different aspects of research into myositis. The second aim

focused on an attempt to treat the disease symptoms with a pilot preclinical trial, while the

third aim was an investigation into the potential connection between persistent muscle

weakness and cytokines of the innate immune system. In the course of this research, we

were able to make several novel observations that are relevant to the design of future

therapies for myositis.

The first aim of this research was to test the hypothesis that an acquired deficiency

of the enzyme AMPD1 is a mechanistic explanation of the persistent muscle weakness in

myositis. In order to address this hypothesis that the loss of AMPD1 we utilized the

transgenic mouse model of myositis. Within this mouse model we were able to show that

healthy mice express high levels of AMPD1, and that the onset of myositis coincides with a

drop in AMPD1 mRNA and protein levels. Furthermore, the loss of AMPD1 appeared to

be a specific symptom of myositis, as mouse models of other myopathies did not

experience a similar loss of this enzyme. Importantly, we observed that AMPD1 activity

was significantly decreased in HT mice prior to the appearance of lymphocytes, and that

this drop in AMPD1 in activity coincided with a loss of strength in the EDL muscle. In

follow up experiments, we were able to achieve a modest knockdown of the protein in

healthy animals with the intravenous administration of small antisense Vivo-morpholinos.

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The modest knockdown of the protein was able to induce significant weakness in the soleus

muscle. We were able to corroborate the results of our knockdown experiment using mice

heterozygous for the AMPD1 gene. Overall, these results provide strong evidence in favor

of the hypothesis that myositis results in an acquired deficiency of AMPD1, and that this

acquired deficiency coincides with muscle weakness independent of autoimmune

inflammation.

The second aim of this research was to attempt to treat the symptoms of muscle

weakness in myositis mice by supplementing diseased mice with metabolites downstream

of AMPD1. Based on the results from the first aim and prior publication, the administration

oral D-ribose was proposed as a possible therapeutic compound for the treatment of

myositis. While there is ample evidence that healthy subjects do not experience any

benefits from ingesting D-ribose, there are several case reports which describe how patients

with a congenital loss of AMPD1 benefited from ingesting D-ribose. Since we

demonstrated that mice with myositis also possess a deficiency of AMPD1 activity, we

sought to evaluate the therapeutic value of oral supplements of D-ribose in the mouse

model of myositis.

However, the results of our trial indicated that treatment with daily doses of oral D-

ribose did not have any significant effect on the performance of myositis mice. In fact, a

disturbing trend appeared where mice treated with D-ribose were, on average, slightly

worse off than untreated mice in terms of voluntary movement, grip strength, and muscle

strength. The only potential benefit of D-ribose administration was observed when fatigue

resistance was measured in the EDL. Myositis mice treated with D-ribose showed a small,

though not statistically significant, improvement in fatigue resistance compared to

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untreated mice. The lack of effect seen for oral D-ribose was attributed to the low

expression of RKBS in the skeletal muscle. This enzyme, which is required for any cell to

prevent ribose from simply diffusing back into the bloodstream, was found to be expressed

at very low levels in mouse skeletal muscle. The low level of RBKS mRNA suggests that

despite the efficient uptake of D-ribose in the lower intestine it is unlikely that skeletal

muscle tissues are able to retain and use any significant amount of D-ribose. These findings

do not directly contradict the hypothesis that the breakdown of AMP releases ribose-1P for

conversion into glucose, since RBKS is not required for the retention of internally

generated ribose-1P. Nevertheless, these findings do cast doubt on the therapeutic benefit

of D-ribose in the treatment of myositis, or any other muscle disorder. Still, these results

are important to the broader field of myositis research since D-ribose is currently marketed

by vitamin manufacturers as a therapeutic agent for the treatment of inflammatory myositis

and chronic fatigue syndrome.

The third aim of this dissertation research was to identify a soluble factor of the

innate immune system that might modulate the expression of AMPD1. As discussed

previously, one of the features of myositis in humans is symmetrical muscle weakness

despite intramuscular inflammation appearing patchy and localized. Additionally it is

known that muscle weakness does not correlate well with lymphocytes infiltration. We

demonstrated previously that weakness and loss of AMPD1 can occur prior to the

appearance of lymphocytes in the muscle tissue. These combined results suggest that some

soluble factor secreted directly from the muscle fibers is capable of inducing muscle

weakness, even in the absence of lymphocytes. In order to investigate this hypothesis, we

set out to identify inflammatory cytokines or other soluble molecules that could account for

84

the early loss of AMPD1 and muscle weakness seen in the skeletal muscle of myositis

mice.

Although there are many cytokines (and a pair of TLRs) that have been implicated

in the pathogenesis of myositis, only a small subset of those described cytokines were

found to be present at the mRNA level in muscle tissues from both H and HT mice. Using

and AMPD1 reporter assay, we tested the effect of each available cytokine for its ability to

affect AMPD1 expression. The results of the in vitro screening assay identified IFN and

IL-1 as the two most potent inhibitors of AMPD1 expression, while IL-15 was found to

significantly increase AMPD1 expression. Follow up experiments demonstrated that

receptors for both IFN and IL-15 were present in the mouse skeletal muscle. Surprisingly,

we found that protein levels for the IL-15 receptor were significantly reduced only in HT

mice. This observation allows us to propose a novel mechanism of weakness, whereby the

loss of IL-15 signaling in myositis results in decreased expression of AMPD1 and muscle

weakness. Currently, little is known about how IL-15 signaling controls the expression of

muscle specific genes like AMPD1.

Based on the results of our in vitro assay and the numerous prior articles that have

associated the Type I interferon response to polymyositis and dermatomyositis, we

proposed that the expression of IFN from skeletal muscle cells would be able to suppress

the transcription of AMPD1 mRNA and induce weakness in neighboring muscle tissues.

Using an IFN expression plasmid and an in vivo intramuscular electroporation protocol,

we were able to test our hypothesis in healthy mice.

85

When IFN was overexpressed in vivo from the TA skeletal muscle, we observed a

significant loss of force in the adjacent EDL muscle, but we did not detect any change in

the levels of AMPD1 within the TA. We could not definitively conclude from this data

whether or not Type I interferons induce muscle weakness via suppression of AMPD1

activity. While this result was in some ways disappointing, it was also a significant result

that will hopefully provide the rationale for more targeted suppression of inflammation in

myositis patients without resorting to the use non-specific treatments such as methotrexate

or cyclosporine.

Future Directions

During the course of these experiments we generated a stably transfected cell line carrying

the luciferase gene under the control of the AMPD1 promoter. With this cell line, we are

currently collaborating with Dr. James Inglese of the NIH Chemical Genomics Center to

perform high throughput screening using this reporter cell line. Small compounds that are

able to stimulate the expression of AMPD1 are expected to hold therapeutic potential for

myositis, and would likely be amenable to preclinical trials using the transgenic mouse

model of myositis.

Apart from drug screening, the role of IL-15 in myositis remains an unexplored

topic. As discussed previously, the role of IL-15 signaling in skeletal muscle was found to

have significant influence over the development of type II (fast) muscle fibers. We found

that IL-15 was found to significantly increase AMPD1 expression in vitro, and the IL-15

receptor appeared to be reduced in myositis mice compared to healthy controls. A loss IL-

86

15 signaling and the maintenance of type II fibers could possibly explain why type II fibers

are more severely affected in mice and patients with myositis. Future experiments will

focus more on exploring IL-15 signaling both in mouse and in human muscle tissues, with

the intent of determining if treatments such as exogenous IL-15 or IL15RA agonists might

hold therapeutic potential for patients with myositis.

87

Chapter 6: Materials and Methods

Animals

Transgenic mouse model of myositis: The generation and genotyping of the HT myositis

mouse model was described previously 50

. All mice were handled according the local

IACUC guidelines. Single-transgenic animals (designated “H mice”) do not develop

myositis. Double-transgenic animals (“HT mice”) spontaneously develop myositis after

doxycycline withdrawal. For controls, age-matched single-transgenic littermates were used.

Only female mice were utilized for our experiments and data analysis. At 5 weeks of age,

doxycycline was withdrawn from the water supply. Mice 15 weeks of age or older were

designated infiltrated mice, and those 8 to 13 weeks old, pre-infiltrated. Muscle tissue was

dissected and flash-frozen in isopentane (Sigma-Aldrich) pre-chilled with liquid nitrogen or

fixed in 10% neutral buffered formalin (Fisher Scientific) for further analysis.

Myositis mice treated with oral D-ribose: All mice were handled according the local

IACUC guidelines. Animals were housed at room temperature with a 12-12 hour light-dark

cycle. Genotyping was carried out at 3-4 weeks as previously described 50

. Briefly, the

mouse model of myositis utilizes double transgenic mouse, where the T gene (an MCK

promoter driven tet-Off transcription factor) forces the expression of the H gene (a TRE

driven H-2Kb MHC class I molecule). Double transgenic animals are labeled „HT‟ and

spontaneously develop myositis after doxycycline withdrawal. Single transgenic animals

are labeled as „H‟ mice and do not develop myositis even in the absence of doxycycline.

For controls, single-transgenic age- and sex-matched littermates were used. For all animals,

88

doxycycline was withdrawn from the water supply at 5 weeks of age, and treatment with

D-ribose (4 g/kg bodyweight) via oral intake. D-ribose (Sigma Aldrich R9629) was

dissolved in double distilled water and prepared fresh weekly for mice. The bodyweight of

each mouse was measured weekly. Symptoms of myositis were readily apparent by 15

weeks of age. Since the disease progresses at different rates in male mice versus female

mice, only female mice were utilized for this trial. At the end of the experiments, the mice

were killed by CO2 inhalation followed by cervical dislocation according to IACUC

guidelines. Muscle tissue was immediately dissected and flash-frozen in isopentane pre-

chilled by liquid nitrogen.

AMPD1 KO mice: Mice heterozygous for the engineered AMPD1tm1a

mutant allele

(KOMP #CSD23933) were obtained from the KOMP repository at UC Davis. These mice

carry a single copy of a null mutation for the AMPD1 gene. The engineered mutation

inserts a splice acceptor, a -galactosidase gene, and a synthetic poly-A terminator

sequence following exon 2 of AMPD1, resulting in a severely truncated and non-function

protein product. When heterozygous AMPD1tm1a

mice are crossed the resulting litters are

composed of 50% wild type and 50% AMPD1tm1a

heterozygous offspring. It is assumed

that AMPD1-/-

embryos are non-viable. It is possible to convert the AMPD1tm1a

allele into a

conditionally knockout allele with the Flp and Cre recombinase enzymes.

Functional and behavioral activities

89

Rotarod test: The latency-to-fall from the Rotarod apparatus was assessed as described

previously 81

. In brief, mice were trained on the Rotarod (Ugo Basile, Italy) for 2 days

before data collection was begun. Each trial consisted of placing the mice on the rod at 10

rpm for 60 sec (stabilizing period), followed by acceleration from 10 rpm to 40 rpm. Each

trial was done twice a day for 3 consecutive days, with a minimum 2-hr interval between

the data collection times. The latency-to-fall was recorded, and average values were

calculated from all six scores.

Grip strength test: Grip strength for both fore- and hindlimbs was assessed using a grip

strength meter consisting of a horizontal forelimb mesh and an angled hind limb mesh

(Columbus Instruments, Columbus, OH) according to a previously published protocol 82

.

The animals were acclimatized on fore- and hindlimb meshes for 3 consecutive days prior

to actual data collection, and for 90 sec before actual data collection. Force was measured

according to the amount of horizontal force that was required to break the mouse‟s grip

from the mesh surface. Five successful hind-limb and forelimb strength measurements

were recorded within 2 min. The maximum values of each day over a 5-day period were

averaged and normalized to body weight and expressed as KGF/kg unit.

Behavioral activity: Voluntary activity in an open field was measured using an open field

Digi-Scan apparatus (Omnitech Electronics, Columbus, OH) as described previously 82

. All

mice were acclimatized to the scanning chamber one week prior to actual data collection.

The data were collected every 10 min over a 1-hr period each day for 4 consecutive days.

The results were calculated as mean ± standard error for all recordings. The recorded

90

quantitative measures of activity were horizontal activity, vertical activity, total distance,

and rest time.

Force contractions on isolated skeletal muscle

Force contraction experiments were conducted on the EDL and soleus muscles of the right

hindlimb of mice. Each mouse was anesthetized with an intraperitoneal injection

containing ketamine (100mg/kg) and xylazine (10mg/kg). The muscles were isolated, and

6-0 silk sutures were tied securely to the distal and proximal tendons. Each muscle was

then carefully removed from the mouse and placed vertically in a bath containing buffered

mammalian Ringer‟s solution (137mM NaCl, 24mM NaHCO3, 11mM glucose, 5mM

KCL, 2mM CaCl2, 1mM MgSO4, 1mM NaH2PO4 and 0.025mM turbocurarine chloride)

maintained at 25˚C and bubbled with 95% O2-5% CO2 to stabilize the pH at 7.4. The distal

tendon of the muscle was tied securely to the lever arm of a servomotor/force transducer

(model 305B, Aurora Scientific) and the proximal tendon to a stationary post in the bath.

After removal of the muscle, the mouse was euthanized by gassing with CO2 according to

IACUC guidelines. The muscle was stimulated between two stainless steel plate electrodes.

The voltage of single 0.2-msec square stimulation pulses was increased until supramaximal

stimulation of the muscle was achieved, and the muscle length was then adjusted to the

length that resulted in maximal twitch force (i.e., optimal length for force generation). With

the muscle held at optimal length, the force developed during trains of stimulation pulses

was recorded, and stimulation frequency was increased till the maximal isometric tetanic

force was achieved. For the EDL muscle, 300-ms trains of pulses were used, and a stimulus

frequency of ~220Hz was typically needed to achieve the maximum isometric force.

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Maximal isometric force for the soleus muscle was achieved at a frequency of 120Hz with

1000-ms trains. The muscle length was measured with calipers, and the optimal fiber

length was calculated by multiplying the optimal muscle length by a constant of 0.45, an

established fiber length/muscle length ratio for EDL muscle and 0.71 for the soleus muscle.

The muscle mass was weighed after removal of the muscle from the bath. The muscle-

specific force, a measure of the intrinsic force generation of the muscle, was calculated

according the following equation: specific force = maximal isometric force/ (muscle mass *

(density of muscle tissue * fiber length)-1

). The muscle tissue density was 1.056 kg/L.

Measurement of skeletal muscle AMPD enzymatic activity

Enzyme isolation: Frozen skeletal muscle samples were homogenized with 50 strokes in an

ice-cold hand-held glass homogenizer containing 150µL of 50mM imidazole, pH 6.8, with

450mM KCl, 5µg/mL leupeptin, and 1mM DTT. Homogenates were clarified by

centrifugation at 14,000 x g for 10 min at 4° C. Supernatants were recovered and dialyzed

overnight at 4° C against 1L of 50mM imidazole, pH 6.5, with 150mM KCl and 1mM

DTT. Dialysates were recovered and clarified by centrifugation at 14,000 x g for 10 min at

4° C. Total soluble protein in the clarified extract was quantified by the Lowry method.

AMP deaminase assay: Soluble protein (20 μg samples) were assayed in 25mM imidazole,

pH 6.5, containing 1mg/ml bovine serum albumin (BSA), 145mM KCl, and 20mM AMP

(1000 µl total volume). Tubes were pre-warmed for 5 min at 37° C, and the assay was

initiated by the addition of substrate (AMP). Aliquots of 50 μl were removed at 5 and 10

min, frozen in a dry ice/ethanol bath, and stored at -20° C. Substrate (AMP) and product

(IMP) were separated on a Whatman Partisil 10-SAX anion-exchange column and

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developed with a linear gradient (100% to 80% buffer A over 8 min) of 5mM ammonium

dihydrogen phosphate at pH 3.4 (buffer A) and 750mM ammonium dihydrogen phosphate

at pH 4.0 (buffer B), at a flow rate of 2ml/min. Column eluate was recorded at 254 nm, and

peaks were quantified based on the correspondence of retention times to external standards

of known concentrations. Data were recorded as mU/mg soluble protein

Quantification of Metabolites

Quantification of Hypoxanthine: Frozen skeletal muscle (quadriceps) was homogenized in

a liquid N2-chilled ceramic mortar and pestle, and then lysed by repeated freeze-thaw

cycles (x3) in RIPA buffer (Sigma). The concentration of soluble protein was determined,

and then cleared lysates were stored at -80°C. The concentration of hypoxanthine in 50 ml

of cleared lysates was then determined with an Amplex Red Xanthine / Xanthine Oxidase

Kit (Invitrogen) according to the manufacturer‟s protocol. The levels of hypoxanthine for

each sample were then normalized using the protein concentration of each sample.

Quantification of fumarate: Frozen skeletal muscle (quadriceps) was homogenized in a

liquid N2-chilled ceramic mortar and pestle, and then lysed by repeated freeze-thaw cycles

(x3) in RIPA buffer (Sigma). The concentration of soluble protein was determined, and

then cleared lysates were stored at -80°C. The concentration of fumarate in the cleared

lysates was determined with a Fumarate Assay Kit (BioVision) using 50 l of lysate

according to the manufacturer‟s protocol. The levels of fumarate were then normalized

93

using the protein concentration of each sample and plotted relative to fumarate levels in

control mice.

Quantification of Inosine Monophosphate: Intramuscular levels of IMP were determined

by triple-quad mass spectrometry. Briefly, frozen skeletal muscle (quadriceps) was

homogenized in a liquid N2-chilled ceramic mortar and pestle, and then resuspended in

250l of 50% methanol solution to precipitate protein. The IMP internal standard was also

resuspended in 50% methanol. Quantities for IMP were acquired in triplicate runs on a

Xevo TQ mass spectrometer (Waters). Serial dilutions of the IMP internal standard (0.1-

1275pmoles/l) were prepared and a standard curve was made with and acquired three

times. Two transitions were selected to quantify the concentration in each sample. Target

Lynx software (Waters) was used to quantify the concentration of IMP the in the samples,

and the quantity of each sample was normalized to the protein concentration of each

sample, as determined by a Bradford protein concentration assay.

Quantitative PCR analysis

For QPCR, RNA was isolated from frozen skeletal muscle tissue by grinding frozen tissue

in a liquid N2 chilled ceramic mortar and pestle (Coors), then homogenizing tissue in Trizol

(Invitrogen) using a Kontes pestle to lyse the tissue. After isolation according to the

manufacturer‟s instructions, RNA was quantified on a Nanodrop N1000

spectrophotometer. A total of 800 ng of RNA was used to produce cDNA using a Promega

Reverse Transcription System kit. For QPCR analysis of human AMPD1 expression, we

94

designed primers for AMPD1 and GAPDH using the Primer3 v0.4.0 software. The

sequence of the primers used for human genes were; AMPD1 forward TGCTG CACAA

GTCTT CAAGC, AMPD1 reverse AGGTA ATTGT CGCCC AGAAA; GAPDH forward

ACAGT CAGCC GCATC TTCTT, and GAPDH reverse AGTTA AAAGC AGCCC

TGGTG. Primers were also designed for mouse genes AMPD1, ADSL, ADSSL1, NT5C2,

PNP, CKM, HK2, RBKS, MX2, IFIT2, ENO3 and GAPDH. The sequence of the primers

used for murine genes were; AMPD1 forward TATCA GCATG CAGAG CCTCG CTTA;

AMPD1 reverse TGTGG CAGGA AATTC TTGGA TCGG, ADSSL1 forward AGACT

CTCCC AGGAT GGAAC, ADSSL1 reverse GTTGC TGGCA ATCCT TAGAA, ADSL

forward TACTT CAGCC CCATC CACTC, ADSL reverse TCACT GTAAC CGGGT

TCTCC, NT5C2 forward CCCAT TCAGC TACCT CTTCA, NT5C2 reverse ATGGC

AGTGT GTGAT CTCCT; PNP, forward GGCTT CTGCA ACACA CTGAA, PNP

reverse TTCAG CAATC CAAAC ACCAG; CKM forward GATTC TCACT CGCCT

TCGTC, CKM reverse GCCCT TTTCC AGCTT CTTCT; HK2 forward AGAAC

CGTGG ACTGG ACAAC, HK2 reverse GCCAG ATCTC TCACC GTCTC; RBKS

forward AAGAA GGCAG CCAGT GTCAT, RBKS reverse GAGCT GGGTT GAACA

AGGTT; ENO3 forward GGTCC CTCTC TACCG ACACA, ENO3 reverse CTTCC

CATAC TTGGC CTTGA; GAPDH forward CGTCC CGTAG ACAAA ATGGT,

GAPDH reverse GAATT TGCCG TGAGT GGAGT. All primers were verified to produce

single, specific amplicons of the correct size before being used for QPCR. All QPCR

reactions were prepared according to the manufacturer‟s protocol using a hot-start SYBR

green premade mix (NEB F-410), and measured on an ABI HT7900 thermal cycler. The

product NEB F-410 has since been discontinued, but the product S4438-100RXN (Sigma-

95

Aldrich). All QPCR programs utilized an initial 15 min hot start at 95º C, followed by a

three-step amplification method of 95º C for 30 sec, 55º C for 30 sec, and 72º C for 30 sec.,

repeated for 40 cycles. Relative gene expression was calculated using the ΔΔCt method,

using GAPDH as the internal reference gene.

Fiber typing by immunohistochemical staining

Fiber typing was performed using primary antibodies recognizing slow-twitch fibers (VP-

M667) and fast-twitch fibers (VP-M665) (Vector Laboratories). Antibodies were diluted

1/50 and 1/25, respectively, and incubated overnight at 4°C, then labeled with Alexafluor

488 (A-21121) for one hour before washing and mounting in Vectashield with DAPI (H-

1200). Images were captured on a Zeiss LSM510 confocal microscope using ZEN 2009

software.

Inhibition of AMPD1 translation

Knockdown of AMPD1 protein levels was accomplished by systemic (IV) injection of

custom Vivo-morpholino oligos (GeneTools Inc., Philomath, OR). The AMPD1 Vivo-

morpholino (abbreviated moAMPD1) 5‟-GAC CTG TTA GTT TGA ATA GAG GCAT -

3‟ and standard Vivo-morpholino control 5‟- CCT CTT ACC TCA GTT ACA ATT TATA

-3‟ were designed by Gene Tools Inc. Vivo-morpholino uptake was enhanced by exercise

on a horizontal treadmill (30 min at 12 m/min) for 2 days prior to injection. Vivo-

Morpholinos were diluted into 100 l of 0.9% saline and injected at 10 mg/kg into the

lateral tail vein of the mice. Injections were performed twice weekly for 2 weeks, for a total

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of four injections. Mice were 14 weeks old when injections began, and 16 weeks old at the

time of sacrifice.

Stable Isotope Labeling in Mice (SILAM)

The development of mice uniformly labeled with heavy 13

C lysine was described

previously by Kruger et al 83

. For quantification of proteins in the myositis mouse model,

we utilized quadriceps muscle lysates from 16 week old labeled female mice. Quadriceps

muscle lysates were also taken from 16 week old female H and untreated HT mice. Total

proteins were extracted from each muscle with RIPA buffer (50 mM Tris-HCl, pH 8.0,

with 150 mM sodium chloride, 1.0% Igepal CA-630 [NP-40], 0.5% sodium deoxycholate,

and 0.1% sodium dodecyl sulfate) with protease inhibitors (Roche protease inhibitor

cocktail 100X). Aliquots of the protein extracts from the muscles of labeled C57BL/6 and

either unlabelled H or unlabelled HT mice were each mixed 1:1 with protein extract from

the muscle of a SILAC-labeled mouse. Labeled and unlabeled protein mixtures were

further resolved by SDS-PAGE. The gel was stained with Bio-Safe Coomassie (Bio-Rad,

Hercules, CA), and each lane was cut into 30-35 serial slices. Proteins in each gel slice

were in-gel digested with trypsin. The resulting peptides from each band were injected via

an autosampler (6 μL) and loaded onto a Symmetry C18 trap column (5 μm, 300 μm i.d. x

23 mm, Waters) for 10 min at a flow rate of 10 μL/min and eluted with 0.1% formic acid.

The sample was subsequently separated on a C18 reversed-phase column (3.5 μm, 75 μm x

15 cm, LC Packings) at a flow rate of 250 nL/min using an Eksigent nano-hplc system

(Dublin, CA). The mobile phases consisted of water with 0.1% formic acid (A) and 90%

acetonitrile (B). A 65-min linear gradient from 5 to 60% B was employed. Eluted peptides

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were introduced into the mass spectrometer via a 10-μm silica tip (New Objective Inc.,

Ringoes, NJ) adapted to a nano-electrospray source (ThermoFisher Scientific). The spray

voltage was set at 1.2 kV and the heated capillary at 200° C. The LTQ-Orbitrap-XL

(Thermo Fisher Scientific) was operated in data-dependent mode with dynamic exclusion,

in which one cycle of experiments consisted of a full MS in the Orbitrap (300-2000 m/z,

resolution 30,000) survey scan and five subsequent MS/MS scans in the LTQ of the most

intense peaks, using collision-induced dissociation with the collision gas (helium) and

normalized collision energy value set at 35%.

Only proteins identified as having ≥2 peptides in H or HT group were considered for

further analysis. The mean ratios were calculated independently of the unlabeled-to-labeled

ratios obtained for each of the comparisons and were further compared to determine

statistically significant (p<0.05) proteomic modulations.

Database search and SILAC ratio measurement: Protein identification and quantification

was performed using Integrated Proteomics Pipeline (IP2) version 1.01 software developed

by Integrated Proteomics Applications, Inc. (http://www.integratedproteomics.com/). Mass

spectral data were uploaded into IP2 software. Files from each lane were searched against

the forward and reverse Uniprot human database (UniProt release 15.4, June 2009, 22697

forward entries) for tryptic peptides, allowing one missed cleavage and possible

modification of oxidized methionine (15.99492 Da) and heavy Lys (6.0204 Da). IP2 uses

the Sequest 2010 (06_10_13_1836) search engine. Mass tolerance was set at ± 30 ppm for

MS and ± 1.5 Da for MS/MS. Data were filtered based on the following Xcorr values

98

(Xcorr ≥ 1.9 for z=1, ≥ 2.5 for z=2, and ≥ 3.5 for z=3). These criteria provided a 3% false

discovery rate. Only proteins that were identified by at least two unique peptides were

retained for further quantitative analysis. Census software version 1.77, built into the IP2

platform, was used to determine the ratios of unlabeled to labeled peptide using an

extracted chromatogram approach. The distribution of ratios was plotted, and correction

factors were applied to adjust for errors in sample mixing. Data were checked for validity

by using regression correlations >0.95 for each peptide pair. The relative change in protein

quantity between H and HT mice was calculated as a ratio by the formula Fold change =

(HT quantity / B6 quantity) / (H quantitiy / B6 quantity), where the same labeled B6

protein lysate same was used as a reference for both H and HT sample lysates.

Western blotting

Western blots were performed as described previously 84

. Briefly, 30 g of protein was

loaded into a 4-12% PA gel and run at 180V for 1.5 h, then transferred onto nitrocellulose

at 80 mA overnight at 4º C. Membranes were blocked in 5% BSA in PBS. Primary and

secondary antibodies were incubated at room temperature in 0.5% BSA in PBS for 2 h.

Primary antibodies specific for AMPD1 (1:2000 dilution, Imgenex IMG-6068A), IFNAR1

(1:500 dilution, Abcam ab45172), IL15RA (1:500 dilution, Abcam ab86486), and vinculin

(1:5000 dilution, Abcam ab18058) were utilized. Secondary goat-anti-rabbit (1:3000

dilution, BioRad #172-1019) and goat anti-mouse (1:3000 dilution, BioRad #172-1011)

secondary antibodies were used to visualize bands on X-ray film (Amersham 28906835)

99

Densitometry analysis was carried out on a BioRad GS-800 Calibrated Densitometer

running the Quantity One software package.

Cell culture of Immorto SV40-A58T myoblasts

The Immorto SV40-A58T myoblast cell line was derived from muscle satellite cells. The

isolation and maintence of these cells has been described in detail previously 79

. In brief,

these myoblasts contain a temperature sensitive variant of the SV40 large T-antigen

oncogene. Expression of the oncogene can be induced by the addition of IFN to the media.

In order to promote myoblast proliferation, cells were maintained in 33°C and received

fresh Growth Media (DMEM high glucose, 20% FBS-Gold, 2% chick embryo extract, 2%

L-glutamine, 1% penicillin/streptomycin) twice weekly. All flasks were treated for

myoblast cell cultures by coating with filtered 0.04% porcine gelatin, then allowing the

gelatin to set at room temperature for 30 min before aspirating. Fresh gamma-interferon

was added proliferating myoblasts at 1U/ml. When cells reached 75% confluency or were

observed to be forming myotubes, culture flasks were washed, trypsinized, and reseeded at

21,000 cells per 75cm2 culture flask. For cell culture experiments, myoblasts could be

induced to form myotubes after reaching 95% confluency, replacing Growth Media with

Differentiation Media (DMEM high glucose, 10% horse serum, 1% penicillin

/streptomycin), and maintaining cells at 37° C degrees 5% CO2. Myoblasts differentiate

and fuse into myotubes between 5 to seven days.

Transfection of Immorto SV40-A58T myoblasts

100

The transfection of Immorto SV40-A58T myoblasts is similar to the manufacturer‟s

standard protocol with some minor modifications as described below. 24 hours prior to

transfection, H2Kb cells were plated into a gelatin treated 12-well plate at 2.5x105 cells/ml.

Cells were then incubated at in a 33º C, 10% CO2 incubator overnight. Cells must reach

80-100% confluency before proceeding with transfection. Plasmid DNA and

Lipofectamine in were diluted separately in blank media, with 1.6 g of DNA dissolved in

100 l of OPTIMEM, while 4 l of Lipofectamine 2000 was dissolved in 100 l of

OPTIMEM. Plasmid DNA and Lipofectamine 2000 were allowed to incubate for 5 min,

then mixed together and allowed to incubate together for an additional 30 min. Plated cells

were quickly washed twice with PBS (w/o Ca2+

or Mg2+

), then all liquid was aspirated off.

A total of 200 l of the transfection mixture was then pipette onto the cells. Next, a volume

of 800 l of blank DMEM-high glucose was then added to the wells, and the cells were

returned to the 33° C, 10% CO2 incubator for no more than 5 hours. After the 5 hour

incubation, the transfection media was aspirated off of the cells, and replaced with 1 ml of

Differentiation Media, then kept in a 37° C, 5% CO2 incubator. Cells were harvested 48

hours post transfection.

Luciferase activity assay

Cells transfected with luciferase reporter plasmid were harvested in Luciferase Lysis Buffer

(25mM Gly-gly, 15mM MgSO4, 2mM EGTA, 1mM DTT, 1% (v/v) Triton X-100, pH to

7.8) and cleared lysates were stored at -80 C until tested. Luciferase activity was be

measured using the Luciferase Assay System (Promega) according to the manufacturer‟s

101

protocol. The data was measured and recorded on a Centro LB 960 Luminometer (Berthold

Technologies). All luciferase activity was normalized using the total protein content of the

lysis sample, as measured by a detergent compatible BCA protein concentration assay

(BioRad).

Intramuscular electroporation of plasmid DNA

For the electroporation of plasmid DNA into the TA of mice, 12 week old female mice

were first anaesthetized with an intraperitoneal injection containing ketamine (100mg/kg)

and xylazine (10mg/kg). Once anaesthetized, the hindlegs of the mice were restrained and

Nair hair removal cream was applied to both hindlegs of the mouse above the TA using a

Q-tip, and allowed to act for 1 min. The fur and remaining Nair cream were then gently

removed with cotton swabs. Plasmid DNA (previously prepared from bacteria using a

Qiagen EndoFree Plasmid Maxiprep Kit) was resuspended to a concentration of 1 g/l in

a 0.9% sterile saline solution, and a total of 30 l was injected into intramuscularly into the

TA using a 27 G needle. The injected volume was allowed to diffuse through the muscle

for 1 min before proceeding to the electroporation. The electroporation consisted of three

pairs of pulses, generated by a ECM 830 Square Wave apparatus (BTX Harvard

Instruments) attached to a 2-needle electrode (BTX 45-0168 and 45-0121). Pulse pairs

were 80V, delivered of 20 msec, with a 980 msec pause between pulses. The 2-needle

electrode was repositioned between each pair of pulses, so that a current was applied

directly across, proximal from, and distal from the injection site. The electroporation

process was performed on both hindlegs, with the right leg receiving empty vector DNA,

102

and the left leg receiving a protein expression plasmid (expressing either GFP or murine

IFN). Electroporated mice were observed carefully until each recovered from anesthesia

and soft gel food was placed in the cage to allow mice to feed without having to bear

weight on the hindlegs. All mice were assessed by in vitro force contraction and sacrificed

12 days post electroporation.

Statistical analysis

Where appropriate, statistical significance was calculated using either Student‟s t-test, one

way ANOVA tests, or two way ANOVA tests for independent samples with Prism v4

software (GraphPad Software). Within each figure, p values are indicated as follows:

p<0.05 is indicated by (*), p<0.01 by (**), and p<0.001 by (***). Graphed data are

presented as means ± standard error. For box and whisker plots, each box marks the 25th

,

median, and 75th

percentile for all data points, while the whiskers above and below the box

mark the maximum and minimum recorded values, respectively.

103

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