Toxicity and Processing of Cellular Prion Protein in Skeletal Muscles

TOXICITY AND PROCESSING OF CELLULAR PRION PROTEIN IN SKELETAL MUSCLES

By JINGJING LIANG

Submitted in partial fulfillment of the requirements For the degree of Doctor of Philosophy

Dissertation Advisor: Dr. Qingzhong Kong

Department of Pathology CASE WESTERN RESERVE UNIVERSITY

January 2012

CASE WESTERN RESERVE UNIVERSITY SCHOOL OF GRADUATE STUDIES

We hereby approve the dissertation of

Jingjing Liang______

Candidate for the Ph.D. degree *.

(Signed) Xiongwei Zhu______(Chair of the committee) Qingzhong Kong______

Neena Singh______

Youwei Zhang______

Clive R. Hamlin______

(Date) 08/23/2011 ______

*We also certify that written approval has been obtained for any proprietary material contained therein.

TABLE OF CONTENTS

Table of Contents ...... 1 List of Figures ...... 3 Acknowledgements ...... 5 Abstract ...... 7 Chapter 1. Introduction ...... 9 Prion Protein ...... 10 General Information ...... 10 Prion Disease ...... 11 Biological Functions ...... 13 Physiological Processing ...... 14 Prion Protein in Skeletal Muscle ...... 15 p53 Pathway ...... 17 General Information ...... 17 p53 and Prion Protein ...... 19 p21 ...... 19 ADAMs ...... 20 General Information ...... 20 ADAM8 ...... 22 Tg(HQK) Transgenic Mouse Line ...... 23

Chapter 2. Microarray and Real-Time PCR Analysis Reveal Activation of p53- Regulated Pro-apoptotic Pathways in Skeletal Muscles Over-expressing PrP ...... 24 Abstracts ...... 25 Introduction ...... 26 Methods ...... 29 Results ...... 35 Discussion ...... 50

Chapter 3. p53 Pathway Plays a Critical Role in PrPC–Mediated Myopathy…..53 Abstracts ...... 54 Introduction ...... 55 Methods ...... 57 Results ...... 60 Discussion ...... 65

Chapter 4. Cellular Prion Protein Regulates Its Own Cleavage through ADAM8 in Skeletal Muscle……...…………………………………………………..67 Abstracts ...... 68 Introduction ...... 69 Methods ...... 73 Results ...... 78 Discussion ...... 89

Chapter 5. Conclusions and Further Directions ...... 93

References ...... 98

List of Figures

Figure 1.1 Schematic structure of full length PrPC.

Figure 1.2 Physiological processing of PrPC.

Figure 1.3 Schematic structure of ADAM protein family.

Figure 2.1 Clustering of gene expression data.

Figure 2.2 MEF2c protein level is down-regulated in skeletal muscles of Tg(HQK) mice treated with Dox.

Figure 2.3 Real-time PCR analysis of Atrogin-1 and MuRF1.

Figure 2.4 Real-time PCR analysis of mdm2 and p53.

Figure 2.5 Upregulation of the p53 pathway: Analysis using the Ingenuity Pathway Knowledge Base (IPKB).

Figure 3.1 Total p53 protein is up-regulated in the skeletal muscles of Tg(HQK) mice treated with Dox.

Figure 3.2 PFT-α attenuates PrP-mediated myopathy in Dox-treated Tg(HQK) mice.

Figure 3.3 PFT-α inhibits p21 up-regulation without affecting p53 protein level in skeletal muscles.

Figure 4.1 Quantitative real-time PCR analysis of ADAMs gene expression.

Figure 4.2 ADAM8 protein level correlates with PrP C1/full-length ratio in skeletal muscle tissue of Dox-induced Tg(HQK) mice.

Figure 4.3 ADAM8 protein level correlates with PrP C1/full-length ratio in skeletal muscle of Tg mice constitutively expressing PrP at different levels.

Figure 4.4 ADAM8 protein level correlates with PrP C1/full-length ratio in skeletal muscle tissue of mice constitutively expressing mouse PrP at different levels.

Figure 4.5 Over-expression of mouse PrP leads to up-regulation of ADAM8 in C2C12 cells.

Figure 4.6 The PrP C1/full length ratio is proportional to active ADAM8 protein level in C2C12 myoblast cells.

Figure 4.7 Over-expression of mouse PrPC leads to up-regulation of ADAM8 in transgenic mice.

Figure 5.1 A model of PrP-mediated myopathy.

ACKNOWLEDGEMENTS

First and foremost, I would like to gratefully acknowledge the enthusiastic guidance and

support from my thesis advisor, Dr. Qingzhong Kong. He was always there to listen and

to give advice. His extensive scientific knowledge, rigorous demands for my research,

creative teaching skills and kindness not only inspire me having a deep thought of my

research study, but also set a moral exemplar of being a real scientist. Without doubt, Dr.

Qingzhong Kong will have a lasting impact on my life.

Second, I wish to express my appreciation to Dr. Mark Smith, who was my previous

committee chair, but unfortunately passed away last December. Dr. Smith was a great person and for me a generous Professor who helped me in my tough struggling time of my Ph.D. He was always a friendly face in my thesis committee meetings, making me at ease with his sense of humor. He would ask some tough but really good questions to make me think and realize something new. Definitely, he would always be one of the best professors and mentors I have met in my memory. I also would like to acknowledge Dr.

Xiongwei Zhu, who is currently my committee chair. He provided me with many helpful suggestions, important advice and constant encouragement during the course of this work.

Besides, I would like to thank the rest of my thesis committee: Dr. Neena Singh, for his encouragement and valuable suggestions, Dr. Youwei Zhang, who gave me insightful comments, provided numerous ideas and useful discussions, and Dr. Clive Hamlin, for his consistent kind help and support. I would also like to thank our collaborators, Dr.

Stephanie Booth’s lab in Canada, for their contributions to the microarray and real-time

PCR analysis; and Dr. Gemma Casadesus, for her help on animal strength assay. I want to thank Dr. Robert Petersen for advice and encouragement.

I also need to thank every member of our group, both past and present. Here I send my gratitude to Wei Wang, Dr. Xinyi Li, Meiling Wang, Laure Farnbauch, Baiya Li, Liuting

Qing, Shenghai Huang, Xiaoqin Liu and so many other people that have already contributed to my research work.

Last, but not least, I am forever indebted to my parents, for giving me life in the first place, for educating me, for unconditional support and encouragement to pursue my interests, and my husband, Xinglong, for understanding, endless patience and encouragement when it was most required.

Toxicity and Processing of Cellular Prion Protein in Skeletal Muscles

Abstract

JINGJING LIANG

The cellular prion protein (PrP) level in muscle has been reported to be elevated in patients with inclusion-body myositis, polymyositis, dermatomyositis and neurogenic muscle atrophy, but it was not clear whether the elevated PrP accumulation in the muscles is sufficient to cause muscle diseases. We have generated a transgenic mouse line [Tg(HQK)] with muscle-specific expression of PrP under extremely tight regulation

by doxycycline, and demonstrated that overexpression of wild-type PrP in skeletal muscles is sufficient to cause a progressive primary myopathy accompanied by preferential accumulation of a truncated PrP fragment termed C1.

To dissect the molecular mechanism of PrP-mediated myopathy in the skeletal muscles of doxycycline-treated Tg(HQK) mice, we performed extensive DNA microarrays, real-time PCR and Western blot analysis and found deregulation of a large number of genes. Among many gene expression changes, prominent up-regulation of p53 and p53-regulated genes involved in cell cycle arrest and apoptosis was found to parallel the initiation and progression of the muscle pathology. I found that treatment with pifithrin-α, a chemical inhibitor of p53 that blocks p53-dependent transcriptional activation and apoptosis, largely prevented severe muscle wasting and decline of muscle

function, confirming the critical role of p53 in PrP-mediated myopathy. I also discovered that ADAM8, a metalloproteinase, was highly up-regulated in the skeletal muscle of

Tg(HQK) mice soon after doxycycline treatment and PrP expression, and the increase of

ADAM8 preceded C1 accumulation. I have demonstrated that, in myoblast cell line

C2C12 and skeletal muscle tissues of transgenic mice, the production of C1 is linearly correlated with active ADAM8 protein level and overexpression of PrP up-regulates

ADAM8. These results indicate that in skeletal muscles, ADAM8 plays a major role in

PrPC processing for C1 fragment production and overexpression of PrPC enables C1

accumulation through upregulation of ADAM8.

CHAPTER 1

Introduction

PrP Protein

General information

Prion protein (PrP), also known as CD230 (cluster of differentiation 230), is encoded by

the highly conserved single-copy PRNP gene (1,2). The human PRNP gene is located on

the short arm of chromosome 20 between the terminus of the arm and position 12, from

base pair 4,615,068 to base pair 4,630,233 (1,2). Two distinct major conformational

forms of PrP have been identified in the nervous system, one is normal cellular isoform

denoted PrPC and the other is scrapie isoform denoted PrPSc. The normal cellular prion protein (PrPC) is mainly expressed in the nervous system, such as brain and spinal cord

(3). Many other cells and tissues, such as blood, lymphocytes, muscle, heart, kidney,

digestive track and skin also express PrPC at lower levels (3). PrPC is a ubiquitous small

glycoprotein attached to the cell membrane by a glycosylphosphatidylinositol (GPI)

anchor (3,4). The NMR predicted PrPC structure includes three helical regions and two β

strands. PrP has two glycosylated sites, one on helix 2 (Asn181 for human PrP) and the

other on helix 3 (Asn197 for human PrP). A disulfide bridge creates a loop between

Cys179 of the second helix and Cys214 of the third helix (human PrP numbering) (3,5).

The octapeptide repeat segment, which binds divalent cations, extends from residue 51 to

91 (3). (Figure 1.1)

The PrP protein is famous for its central role in prion diseases (see below). According to

the protein-only hypothesis of prion propagation, PrPC can be converted into a range of self-propagating disease-related aggregated conformers named PrPSc, leading to prion

diseases (3,4). The abnormal PrPSc isoform and PrPC have distinct secondary and tertiary

Figure 1.1

Schematic structure of full length PrPC. The normal cellular prion protein has three helical regions, two beta strands, and two asparagine-linked glycosylation sites at residues 181 and 197. A disulfide bridge between residues 179 and 214 creates a loop between Cys179 of the second helix and Cys214 of the third helix.

structures but share identical primary sequence. It was reported that normal PrPC has 43%

α-helix and 3% β-sheet, while PrPSc has 30% α-helix and 43% β-sheet. Unlike protease- sensitive PrPC, PrPSc is resistant to proteolysis. The detailed mechanism by which PrPC is

converted into PrPSc remains unclear, but PrPC is known to be required for PrPSc-

mediated pathogenesis (6).

A wealth of evidence indicates that PrPC is beneficial to the cell. However, depletion of

PrPC is relatively innocuous (6). The PrP-null mice live a normal life span without

displaying obvious developmental defects and only present subtle phenotypes, such as

mild cognitive and behavioral deficits (6). Postnatal knockout of PrP expression in the

brain did not affect neuronal survival in transgenic mice either (7).

Prion Disease

Prion diseases, also named transmissible spongiform encephalopathies (TSEs), are a

group of fatal neurodegenerative diseases that affect the central nervous system of many

animals, including humans and many animals, such as cattle, sheep, cervids (deer and

elk), mink, and rodents (8,9). According to the prevailing hypothesis, TSEs are

transmitted by prions, though some scientists still believe the involvement of

Spiroplasma infection (10). Progressive brain degeneration leads to rapidly progressive

deterioration of mental and physical capacities (10,11). The degenerative brain tissue

damage caused by prion diseases is characterized by spongiform change, neuronal loss,

astrocytosis and PrPSc deposits (11). Common clinical signs in humans include

personality changes, psychiatric problem such as depression, lack of coordination and an

unsteady gait (11).

Prion diseases of humans include classic Creutzfeldt-Jakob disease, new variant

Creutzfeldt-Jakob disease, Gerstmann-Straussler-Scheinker Syndrome, fatal familial insomnia and kuru (8,9). Animal prion diseases include bovine spongiform encephalopathy of cattle (commonly known as mad cow disease), chronic wasting disease of cervids, scrapie of sheep and goat, transmissible mink encephalopathy for mink, feline spongiform encephalopathy for cats, and exotic ungulate spongiform encephalopathy of zoo animals (8,9).

Unlike other kinds of infectious diseases which are spread by microbes, the infectious

agent in prion diseases is a specific misfolded protein called scrapie prion protein (PrPSc)

(1,2). PrPSc carries the disease between individuals and cause deterioration of the brain.

Pathogenic mutations in the PRNP gene cause familial/genetic prion disease (1,2), believed to result from the enhanced tendency of the mutant PrP protein to adopt the

disease-causing conformation. However, most human cases of prion diseases are sporadic

whose cause is unknown. Rarely, prion diseases also can be transmitted by consumption

of prion-contaminated food or exposure to prion-contaminated materials (1,2,7). Since

host PrPC expression is essential for prion replication and prion pathogenesis, knocking-

out or reducing PrPC expression in the host brain is actively pursued to treat and prevent

prion diseases.

Biological Functions

PrPC is expressed at high levels throughout adult life, predominantly in neurons.

Numerous works has been done to unravel PrPC functions in neuronal systems. PrPC is

mainly localized at synapses, in cholesterol-rich micro-domains or caveolae (12,13). PrPC

was reported to play a functional role in neuronal cell adhesion, migration and

differentiation by modulating different cell-signaling pathways (14), and it interacts with

several neuronal proteins, including Bcl2, Bax, stress-inducible protein 1 as well as cell

adhesion molecules or extracellular matrix proteins, such as laminin, vitronectin and

NCAM, to mediate the neuritogenesis and neuronal differentiation in several cell models

(15-20). Besides, PrPC was also reported to show neuroprotective activity through

influencing neuronal and glial factors involved in antioxidative defense (21).

PrPC plays important roles in other tissues/cells as well. It was found to be expressed on

long-term repopulating hematopoietic stem cells and is important for their self-renewal

(22), recruits important interacting signaling molecules to influence T cells activation

through association with the lipid raft proteins reggie-1 and reggie-2 (23), and was suggested to function in embryogenesis by regulating embryonic cell adhesion (24-26).

Interestingly, accumulation of PrPSc (or PrPSc-like forms) was found in the muscles of

prion-affected humans and animals (27-32) and in transgenic mouse models of some

familial prion diseased (33), suggesting that skeletal muscles could be involved in some

prion diseases. Besides, overexpression of wild type PrPC (34-35) or expression of

pathogenic PrP mutants, leads to myopathy in transgenic mice (33,36). Recently, it was reported that PrPC promotes regeneration of adult muscle tissue through activating the

stress-activated p38 pathway to induce myogenic precursor cells to exit cell cycle (37).

Physiological Processing

PrPC has been reported to undergo physiological proteolytic cleavages (38,39,40). One of

the cleavages generates an N-terminal fragment, referred to as N1 that is released in the

extracellular medium, and a C-terminal fragment, referred to as C1 that remains tethered

to the plasma membrane, a process termed α-cleavage that is both constitutive and

protein C (PKC)-regulated (41,42) (Figure 1.2A). This cleavage occurs at the 110/111

peptide bond, which is within the 106-126 domain of PrPC that is thought to convey

intrinsic toxicity and essential for PrPC to PrPSc conversion (43-45). Thus, this cleavage

has been thought as a physiological means to deplete cells of convertable PrPC and

PrP106-126 associated toxicity (43-45). Three ADAM (A Disintegrin And

Metalloproteases) proteins were reported to contribute to this proteolytic cleavage of

PrPC in several neuronal cell lines, either directly for ADAM10 and ADAM17 (46) or

indirectly for ADAM9 (47), but recent reports suggest they do not play a significant role

in the α-cleavage of PrPC (see Chapter 4). ADAM10 also appears to be a strong candidate

for PrPC cleavage in human brain (48). Besides, PrPC was also reported to undergo an

upstream cleavage within the octapeptide repeats (around residues 90/91), yielding a

7kDa N-terminal peptide (termed N2) and a 21-kDa C-terminal fragment (referred to as

C2) (42) (Figure 1.2B). Very few works have examined the protease involved in this

pathological cleavage.

The putative functions of C1 and C2 remain to be established. It was reported that C1 but

not C2 increased p53 transcription and p53-like immunoreactivity and activity in

HEK293 cells (49).

Prion protein in skeletal muscle

Skeletal muscle expresses PrPC at significant levels (25,50). Although no major

abnormalities of neuromuscular function had been described in early works with PrP

knockout mice, more recent reports indicate that ablation of PrP gene may affect skeletal

muscles by enhancing oxidative damage (51) and diminishing tolerance for physical

exercise (52), suggesting a role for PrPC in muscle physiology. Skeletal muscle has also been reported to be associated with prion pathology, as evidenced by the accumulation of

PrPSc (or PrPSc-like forms) in the muscles of TSE-affected humans and animals

(27,29,31,32,53,54) and in transgenic mouse models of some inheritable prion diseases

(33).

Expression of PrPC has been reported to be increased in the skeletal muscles of patients

with sporadic and hereditary inclusion body myositis (55,56), polymyositis,

dermatomyositis, and neurogenic muscle atrophy (57). Moreover, a uniform pattern of

increased PrPC expression was described in a series of muscular disorder, suggesting that

PrPC may play a role in stress-response effect in neuromuscular disorders (58). The PrP was proteinase-K sensitive, ruling out PrPSc involvement. Studies with an experimental

Figure 1.2

Physiological processing of PrPC. (A) In physiological conditions, PrPC is endoproteolyzed mainly at the 110/111 peptide bond, yielding to a 17-kDa C-terminal fragment C1 tethered to the plasma membrane, named C1, and a soluble N1 product. (B) PrPC also undergoes an upstream cleavage near the octapeptide repeats (around residues 90/91), yielding a 7kDa N2 product and a 21-kDa C2 fragment.

model of chloroquine-induced neuromuscular disease indicated that PrP molecules that accumulated in diseased muscle fibers have distinct biochemical and biological properties

(59). Transgenic mice overexpressing the Prnpb allele in a body-wide fashion suffered

from a profound, gene dose-dependent necrotizing myopathy associated with demyelination of peripheral nerves and vacuolization of the CNS at advantaged ages (35).

However, transgenic mice overexpressing the Prnpa allele showed no signs of myopathy,

but that could be due to the difference of the transgene vectors rather than the Prnp alleles

(60). As will be described in chapter 2, we have generated transgenic mice with muscle-

specific expression of human PrPC under tight regulation by doxycycline and

demonstrated that overexpression of wild type PrP limited to skeletal muscles is

sufficient to cause a primary myopathy with no signs of peripheral neuropathy (34).

PrPC was also reported to be associated with muscle differentiation and regeneration.

PrPC was upregulated when myotubes differentiate from immortalized C2C12 murine

myoblasts (61). In addition, PrPC level was reported to increase during maturation of myocytes in primary cultures of skeletal muscle, due to both transcriptional and posttranslational changes (50). Interestingly, fast muscle fibers present a higher concentration of PrPC than slow fibers, suggesting a role of PrPC in skeletal muscle physiology (50). Using an in vivo skeletal muscle paradigm from wild type and PrP knockout mice, Stella et al. (2010) evaluated the role of PrPC in the myogenic process

from the response to inflammation to the full recovery of damaged muscle, and

concluded that PrPC was one of the factors that promote regeneration of adult muscle

tissues through the stress-activated p38 pathway (37). This finding indicated that PrPC

might play a significant functional role in proliferation and differentiation of the

myogenic precursor cells.

p53 pathway

General information p53 is a transcription factor encoded by the TP53 genes (62,63). It regulates cell cycle

and functions as a tumor suppressor (62,63). p53 has multiple anti-cancer mechanisms:

activating DNA repair proteins when DNA has sustained damage, arresting the cell cycle

at the G1/S transition point on DNA damage recognition, and initiating apoptosis or

programmed cell death when DNA damage proves to be irreparable (62-64). p53 exhibits

17 sequence specific DNA-binding, directly interacts with various cellular and viral proteins, and induces cell cycle arrest in response to DNA damage (65,66). In response to signals generated by a variety of genotoxic stress, such as UV irradiation or DNA damage, p53 is expressed and undergoes post-translational modification, resulting in its accumulation in the nucleus (67), which binds to many target genes and leads to cell cycle arrest and in some cases to apoptosis. Thus, the p53-dependent pathways help to maintain genomic stability by eliminating damaged cells.

Point mutations of the wild-type p53 gene are key events in the development of malignancy as the mutant protein acts as the dominant regulatory oncogene. Indeed, mutations of the p53 gene are the most common molecular changes identified in human cancer. They have been reported to be a frequent feature of breast, lung, colon, ovarian, brain, testicular and bladder cancers, melanoma, neurofibrosarcoma, and certain types of leukemia (68). In all of these cases, the mutation is found only in the tumor tissue and not in the normal tissue.

The human wild-type p53 protein is a 393 amino acid nuclear phospho-protein, present in the nucleus of all normal mammalian cells and appears to be involved in the regulation of cell proliferation. The normal protein has a very short half-life and is present in only minute amounts in normal tissue and cells. In contrast, mutant p53 proteins produced by malignant cells usually have significantly prolonged half-life.

In primary neurons, neuronal cell lines, and rodent models of cerebral ischemia focused on early signaling events in apoptosis, accumulation and transcriptional activation of p53 have been reported to occur rapidly in response to a wide variety of insults including

DNA damage, oxidative stress, metabolic compromise or excitotoxicity (69-72). Upon

activation, p53 translocates to the nucleus and initiates the transcription of many pro-

apoptotic target genes including Bax, and the BH3-only proteins PUMA and Noxa.

Besides, p53 can also directly trigger apoptosis by translocating to mitochondria to

rapidly induce cytochrome c release, which often occur in synaptic terminals (73-76).

p53 and prion protein

Several prominent neurological disorders, including stroke, Alzheimer’s disease, prion

diseases, manifest symptoms that result from degeneration and death of neurons (76).

Recent reports showed that p53 may be involved in neurodegenerative process and

contributed to neurodegenerative diseases (77-79). There have been a few reports of

apoptotic cells in the brains of animals and humans with prion diseases and have further

suggested the activation of the p53 pathway for apoptosis (80). In terms of the

relationship between p53 and PrPC, several works have focused on studying the role of

p53 in PrPC overexpression induced cell death in several neuronal cell lines. It was

proposed that endogenous cellular PrP sensitizes neurons to apoptotic stimuli through a

p53-dependent caspase-3-mediated activation controlled by Mdm2 and p38 MAPK (81),

and PrPC-induced caspase-3 activation was found to be closely linked to its endocytosis

(82). Furthermore, PrPC seems to regulate p53-dependent caspase-3-mediated neuronal cell death through up-regulation of p53 promoter transactivation and increasing p53 stability via Mdm2 expression (83). It was reported that the PrP C1 fragment, but not C2,

leads to the increase of p53-like immunoreactivity (49).

p21

p21/WAF1, also known as cyclin-dependent kinase inhibitor 1 or CDK-interacting

protein 1, is a protein that in humans is encoded by CDKN1A gene located on

chromosome 6 (84). The p21 protein binds to and inhibits the activity of cyclin-CDK2 or

cyclin-CDK4 complexes, and thus functions as a regulator of cell cycle progression at G1

(85). The expression of this gene is tightly controlled by the tumor suppressor protein p53,

thereby mediating p53-dependent cell cycle G1 phase arrest in response to a variety of

stress stimuli (85). p21 can also function as a senescent cell-derived inhibitor to mediate cellular senescence (86), and interact with proliferating cell nuclear antigen (PCNA) to regulate S phase DNA replication and DNA damage repair (87).

A Disintegrin And Metalloproteinase (ADAM)

General information

ADAM, A Disintegrin And Metalloproteinase protein, is a family of peptidases, consisting of propeptide, metalloprotease, disintegrin-like, cysteine-rich and epidermal growth factor like domains. As ADAMs are membrane proteins, they also contain transmembrane and cytoplasmic domains. The domains contained within the ADAMs family have been characterized, and linked with their functional and structural roles (88).

Besides, ADAMs contain a consensus sequence with three histidine residues that bind to the catalytically essential zinc ion. Importantly, the propeptide is removed through cleavage by a furin type protease, yielding the active enzyme. ADAMs participate in a wide variety of cell surface remodeling processes, including ectodomain shedding, regulation of growth factor availability and mediating cell-matrix interactions. For example, ADAM17 and ADAM10 participate in proteolytic cleavage of the ectodomain

Figure 1.3

Schematic structure of the ADAM protein family. The ADAM family proteins consist of propeptide, metalloprotease, disintegrin-like, cysteine-rich, epidermal growth factor like domains, transmembrane and cytoplasmic domains. Cys-rich, cysteine-rich domain; EGF-like, epidermal growth factor like domain; TM, transmembrane domain.

of Notch-1 to produce the cytoplasmic fragment of Notch-1, which transfers to the

nucleus for cell signaling (89); the ephrins EPH receptor A2 and A3 are shed by

ADAM10 , releasing soluble Eph receptors, which inhibit tumor angiogenesis in mice

(90); ADAM17 is able to release active tumor necrosis factor-α (TNF-α) and heparin- binding EGF-like growth factor (HB-EGF) from their membrane bound precursors, which can indirectly affect angiogenesis (91). Besides, dysregulation of ADAMs is also linked to cardiovascular disease, asthma, and Alzheimer’s disease (92).

Importantly, ADAM proteins were reported to be most important factors contributing to the cleavage of PrP to generate C1 and N1. Inhibition of ADAM10 and ADAM17 was reported to drastically reduce PrP C1 production in HEK293 cells (46). Besides, data from over-expression of human ADAM10 and ADAM17 suggest that ADAM10 contributes to constitutive C1 formation whereas ADAM17 mainly participates in regulated C1 formation (46,93). ADAM-9 is also implicated in the constitutive secretion

21 of C1 in HEK293 cells, TSM1 neurons and mouse fibroblasts by promoting the shedding of active ADAM10 (47). However, more recent articles indicate these ADAMs play no major role in C1 production although they do result in PrP shedding (see Chapter 4).

ADAM8

ADAM8 is a member of the ADAM family. It is a transmembrane glycoprotein, also known as MS2 and CD156. It was first described as a monocyte-specific protein originally cloned from mouse macrophages (94). Human ADAM8, mapping to the human chromosome 10q26.3, was isolated from cDNA libraries of the human macrophage cell line THP-1 and from human granulocytes (95). The name CD156 was given to indicate that ADAM8 is a leukocyte differentiation antigen that may play an important role in the immune system. ADAM8 (826 amino acids) contains a type-I transmembrane domain and a canonical HExxHxxxxxH zinc metalloproteinase motif that has been shown to be proteolytically active (96).

ADAM8 is up-regulated in the central nervous system following neurodegeneration and activation of glia cells (astrocytes and microglia), suggesting that it may have a role in neuron-glia interactions (97). It has been shown to be produced by bone marrow cultures stimulated by vitamin D (98). ADAM8 stimulates osteoclasts, suggesting a role in cell adhesion and cell fusion (98). Although ADAM8 was first reported to be monocyte- specific, it is actually expressed in a wide range of cells in culture. Studies of recombinant ADAM8 show that little or none is expressed on the cell surface; the majority is shed in the cell culture media as a 70 kDa form, similar to ADAM8 secreted into media of bone marrow cells (99).

ADAM8 knockout mice have been generated, which showed no major defects during

development or adulthood and no evident pathological phenotypes (100).

Tg(HQK) transgenic mice

Our lab has generated the Tg(HQK) transgenic mice with muscle-specific expression of

PrP under extremely tight regulation by doxycycline (Dox). We have demonstrated that

Dox-induced, dose-dependent over-expression of PrP in Tg(HQK) mice leads to a progressive primary myopathy (34). Under induction with 6g of Dox per kg of food, PrP

expression in Tg(HQK) skeletal muscle reached the peak level at 14 days post induction;

the accumulated PrPC in Tg(HQK) mice skeletal muscle was cleared within 2 weeks after

Dox withdrawal (34).

Increased variation of myofiber size and endomysial fibrosis in the absence of intra-

cytoplasmic inclusion and rimmed vacuoles are important features of PrP-mediated

myopathy in Tg(HQK) mice. At day 7 of Dox treatment, a few scattered cells with

central nuclei in skeletal muscles were visible in Tg(HQK) mice. After 4-5 weeks of Dox

induction, nearly 15% of muscle fibers exhibited one or more centrally located nuclei,

and a few showed signs of necrosis and regeneration. PrP C1/full-length ratio also increased significantly from day 7 of Dox treatment, suggesting that the accumulation of

C1 is associated with PrP-mediated myopathy (34).

CHAPTER 2

Microarray and real-time PCR analysis reveal activation of p53-regulated pro- apoptotic pathways in skeletal muscles over-expressing PrP

ABSTRACT

We have reported that doxycycline-induced over-expression of wild type prion protein

(PrP) in skeletal muscle of Tg(HQK) mice is sufficient to cause a primary myopathy with no signs of peripheral neuropathy. The preferential accumulation of the truncated PrP C1

fragment was closely correlated with these myopathic changes. In this study we use gene

expression profiling to explore the temporal program of molecular changes underlying

the PrP-mediated myopathy. We use DNA microarrays and confirmatory real-time PCR

analysis to demonstrate deregulation of a large number of genes in the course of the

progressive myopathy in the skeletal muscles of doxycycline-treated Tg(HQK) mice.

These include the down-regulation of genes coding for the myofibrillar proteins and

transcription factor MEF2c, up-regulation of genes for lysosomal proteins that is

concomitant with increased lysosomal activity in the skeletal muscles, up-regulation of

genes for p53-related pathways involved in cell cycle arrest and promotion of apoptosis

that paralleled the initiation and progression of the muscle pathology. The data shows that

several mechanistic features contribute to the myopathy observed in PrP over-expressing

mice, especially p53-related apoptotic pathways.

INTRODUCTION

Cellular prion protein (PrPC) is a ubiquitous glycosylphosphatidyl-inositol (GPI)-

anchored glycoprotein that has gained enormous attention as the central factor in prion

diseases (4). In these diseases PrPC is converted through conformational change to a pathological form (PrPSc) that self-replicates using PrPC as the substrate. The normal

functions of PrPC remain elusive despite concerted efforts. PrPC has been implicated in

CNS development, neurite outgrowth and neuronal survival, early synaptic neuronal

transmission and reorganization of neuronal circuitry within the hippocampus, regulation

of circadian rhythm, memory formation and cognition, maintenance of Ca2+ activated K+

currents in hippocampal CA1 pyramidal neurons, protection against brain injury in rat

and mouse models of ischemic stroke and in T cell development and function (101).

Over-expression of PrPC has been shown to exert a protective effect in BAX and TNF-α-

mediated cell death and conversely a pro-apoptotic function in studies of staurosporin-

induced cell death (83,102,103). It has been demonstrated that depletion of endogenous

PrP reduces susceptibility to staurosporine-induced caspase-3 and p53 activation (81).

Although PrPC is mainly expressed in neuronal systems and neurons are generally

regarded as the model of choice for studying PrPC function and physiological processing,

the expression of the protein in several other organs suggests that PrPC has a conserved role in different tissues and study of PrPC function and processing in other tissues is also of importance. One such tissue is skeletal muscles, which express significant amounts of

PrPC (104,105) related to muscle pathophysiology. On one hand, skeletal muscle has

been found to upregulate PrPC in patients with sporadic and hereditary inclusion body

myositis (55,106), as well as in patients with polymyositis, dermatomyositis and

neurogenic muscle atrophy (57); on the other hand, skeletal muscle has been shown to be

directly affected in PrPC knockout mice, characterized by enhanced oxidative damage

(107) or diminished tolerance for physical exercise (52).

In a previous study, we generated the Tg(HQK) mouse line that expresses human PrPC

exclusively in the skeletal muscles under tight regulation by Dox (34). We found that

induced over-expression of PrPC in the muscles leads to a progressive primary myopathy

characterized by increased variation of myofiber size, centrally located nuclei and

endomysial fibrosis, in the absence of cytoplasmic inclusions, rimmed vacuoles, or any

evidence of a neurogenic disorder (34). While the pathogenic mechanism of the PrPC-

mediated myopathy was not determined, we found that the myopathy was accompanied

by preferential accumulation of PrP C1 fragment, which is also found in the skeletal

muscles of wild-type mouse but at a much lower level [a molar ratio of close to 1:1 over

full-length PrPC in wild type mice in contrast to a ratio of 3:1 in the Dox-induced

Tg(HQK) model] (34). A number of studies have shown expression of N-terminus truncated forms of PrPC to be associated with toxicity in animal models (108, 109). The protein Doppel, which is homologous to the C-terminus of PrP, has also been shown to be cytotoxic when ectopically expressed in neurons (110-112). In both cases, the toxicity can be abrogated by the co-expression of full length PrPC (113,114). The C1 fragment has

also been reported to potentiate staurosporine-induced toxicity via caspase-3 activation in cultured cells (49), but this toxic effect is similar to what was reported for full-length

PrPC (81-83). Thus, it is highly possible that PrPC overexpression-mediated myopathy in

skeletal muscles of Tg(HQK) mice is due to accumulation of PrP C1 fragment.

In this chapter, in order to understand the molecular mechanism of PrP-mediated

myopathy, we performed microarray analysis and real-time PCR to determine gene

regulatory networks that were triggered following overexpression of PrPC in the skeletal muscles of Tg(HQK) mice with increasing duration of Dox treatment at short time

intervals.

METHODS

Animals and Treatment

The doxycycline-inducible Tg(HQK) mice were described previously (34). The HQK

transgene contained two genes: reverse tetracycline responsive transcription activator

(rtTA) under the control of the mouse PrP promoter of the half genomic PrP clone, and

human PrP ORF regulated by the tetracycline-responsive promoter (tetO-hCMV*-1) from

the core plasmid (115). The Tg(HQK) mice were generated in the FVB background, and

Tg(HQK)/Prnp0/0 mice were obtained through breeding with the Zurich I PrP-null mice

(60) in FVB background. Line Tg(HQK) 18, referred to as Tg(HQK) for simplicity, was

used for this study.

Animal Treatment and Specimen Collection

Wild type (WT), PrP-null (KO), and Tg(HQK) mice were fed food pellets either lacking or containing 6 g doxycycline (Dox)/kg food (Bio-Serv) to induce PrPC expression.

Skeletal muscles from the quadriceps of hind legs were removed at day 0, 4, 7, 14, 30 and

60 days following administration of Dox. For immunoblot and microarray analysis, the muscle tissues were immediately frozen on dry ice, and stored at -80°C.

RNA Isolation

Total RNA was isolated from frozen skeletal muscle using the RNeasy skeletal muscle

RNA isolation kit (Qiagen) following the manufacturer’s specifications. The total RNA preparations were further treated with Turbo DNA-Free DNase (Ambion) to remove

29 residual genomic DNA contamination, and examined with Bioanalyzer 2100 (Agilent) for purity and quantity.

RNA Amplification and Labeling for Microarray Analysis

Total RNA was amplified and labeled for microarray analysis using the AminoAllyl

Message Amp II aRNA amplification kit (Ambion) following the manufacturer’s specifications. In brief, 1 µg total RNA was reverse transcribed to first-strand cDNA, followed by subsequent second-strand cDNA synthesis. In vitro transcription to synthesize amplified aRNA was performed and the resultant aRNA quantified. Ten to fifteen micrograms of aRNA was designated as reference (WT) or experimental (KO,

HQK), and then coupled to either Alexa Fluor succinimidyl ester 555 or Alexa Fluor succinimidyl ester 647 dye in 30% DMSO/coupling buffer in the dark at room temperature for 1 hour. Each sample was labeled individually with both Alexa Fluor 555 and 647 for subsequent dye-swapped hybridizations to account for intensity bias.

Uncoupled dyes were removed and labeled aRNA purified following the manufacturer’s specifications. cDNA Microarrays

A total of 16,315 cDNA expressed sequence tags from the Brain Molecular Anatomy

Project (BMAP) mouse brain library http://www.ncbi.hlm.nih.gov were spotted in duplicate onto CMT-GAPS Gamma Amino Propyl Silane coated glass slides (Corning) using the Virtek Chip Writer. Five micrograms of both reference (WT) and experimental

(KO and HQK) Alexa Four labeled aRNA were used in each competitive hybridization.

Each labeled aRNA was resuspended in 35 µl DIG Easy HybTM hybridization buffer

(Roche) containing 20 µg mouse cot1 DNA and 20 µg poly(A)-DNA to block non- specific hybridization. Three biological replicate samples from each of the reference and experimental groups were combined, heated for 5 minutes at 95°C, then cooled and maintained at 42°C. The labeled aRNA sample mixtures were added to a BMAP microarray and incubated in the dark at 42°C overnight to competitively hybridize to

reference and experimental samples. The number of slides hybridized in each experiment

corresponded to the number of biological replicates in each group of experimental

interest. Following hybridization, the slides were washed once in low stringency wash

buffer (1×SSC, 1.2% SDS) preheated to 42°C for 5 minutes, once in high stringency

wash buffer (0.1×SSC, and 0.2%SDS) for 5 minutes at room temperature, and then once

in 0.1×SSC for 5 minutes at room temperature. The slides were analyzed in two

channels using the Agilent HT microarray scanner (Agilent). Raw, background and net

intensity values were collected using Array-Pro software (Media Cybernetics). In order to

account for variation in fluorescence, LOWESS sub-grid normalization was performed

by Gene Traffic software (Iobion), and the subsequent normalized log2 ratios obtained.

The resulting ratio between reference and experimental signals for each individual gene

was used as a measure of differential gene expression using EDGE (Extraction of

Differential Gene Expression), an open source software program for the significance

analysis of DNA microarray experiments (http://www.genomine.org/edge/).

EDGE implements statistical methodology specifically designed for time course experiments (116). A significance measure is assigned to each gene via the Q value (false discovery rate) methodology (117). We selected a Q-value cutoff to display the genes that

met our significance threshold. We performed a “between class” analysis of the data

over time; the class variables, or biological groups, were the PrP over-expressing mice

[Tg(HQK)] and the PrP-KO mice.

Agilent Whole Mouse Genome Oligonucleotide Microarrays

One microgram of each Alexa Fluor 555 and 647 labeled samples as prepared above were

fragmented, reference and experimental samples together, in 250 µl fragmentation mix in

preparation for hybridization to Agilent’s Whole Mouse Genome 44 K oligonucleotide

microarrays. Following the manufacturer’s protocol, an equal volume of 2 ×

hybridization buffer was added to stop RNA fragmentation and prepare the samples for

hybridization. Four hundred fifty microliters of each mixture containing the reference and

experimental samples was then added to an individual slide hybridization assembly and

allowed to rotate at 4 rpm at 65°C for 17 hours. Slides were washed and scanned as

recommended in the protocol, then analyzed using Agilent Feature Extraction Software.

Raw, background and net intensity values were collected. A linear and LOWESS

normalization correction method was selected in order to account for variations in

fluorescence. A two-sided t-test of feature versus background, set at a p value of 0.05,

was used to obtain a list of genes whose log10 ratios were significant.

Validation of Gene Expression Using Quantitative PCR

Some of the genes that appeared to be differentially regulated based on microarray

analysis were confirmed with quantitative real-time PCR (qRT-PCR), using probe specific TaqMan gene expression assays on the Applied Biosystems 7500 Fast Real-Time

PCR system. 100 ng of total RNA previously isolated and used for microarray analysis was reverse transcribed using the High Capacity cDNA Reverse Transcription kit.

Subsequently, 1 µl from each reverse transcription reaction was assayed in a 20 µl single-

plex reaction for real-time quantification with the 7500 Fast PCR System using probes

specific to those genes of interest. Each sample was run in biological triplicate, of which

3 technical replicates were performed. GAPDH was used as the endogenous control, and

gene expression of target genes for KO and HQK samples were quantitatively measured

relative to the WT samples. Relative quantification values were determined using the 2-

ΔΔct method, and expressed as fold-change over WT.

Immunoblot Analysis

Mouse skeletal muscle tissues were homogenized in lysis buffer containing 50 mM Tris

(pH7.5), 200 mM sodium chloride, 0.5% sodium deoxicholate, and 5 mM EDTA. Protein concentrations were determined by the BCA protein assay (Pierce). After addition of

LDS sample buffer (Invitrogen) and sample reducing agents (Invitrogen), the homogenates were denatured at 100°C for 10 minutes, and the proteins were revolved on

10% NuPage Tris-Bis Gels (Invitrogen) and blotted onto nitrocellulose membranes

(Invitrogen). For MEF2c detection, the membrane was incubated with a rabbit polyclonal anti-MEF2c antibody (Cell Signaling) (1:5000 diluted in 0.5% normal goat serum

[Vector Laboratories], 1×TBS, 0.1% Tween-20) at 4°C with gentle shaking overnight.

The blots were developed with the Immobilon Western Chemiluminescent HRP substrate

(Millipore) according to the manufacturer’s instructions. Skeletal muscle actin was

probed with a rabbit polyclonal antibody (Abcam) (1:5000 diluted in 0.5% normal goat

serum, 1×TBS, 0.1% Tween-20) similarly after stripping the blots with a stripping buffer

containing 1.4% 2-mercaptoethanol, 2% SDS and 62.5 mM Tris (pH 6.8). The Western

33 blots were scanned and the protein bands were quantified with the UN-SCAN-IT gel 6.1 software (Silk Scientific).

RESULTS

Induction of PrPC Specifically in the Skeletal Muscle of Transgenic Mice Results in a

Temporally Regulated Transcription Profile

The transgenic mice [Tg(HQK)] used in this study have been described previously, in which PrPC is exclusively expressed in skeletal muscles under the strict control of doxycycline (Dox) and the induced over-expression of PrPC leads to a progressive

primary myopathy (34). To determine the temporal patterns of gene expression that accompany the induced myopathy, we carried out microarray analysis of skeletal muscles

from Tg(HQK) mice, wild-type FVB mice (WT) and PrP-knockout control mice (KO)

using a 16,315-gene cDNA array constructed in Stephanie Booth’s laboratory. Skeletal

muscles from the hind legs (quadriceps) of the mice were collected at 0, 4, 7, 14, 30, and

60 days following administration of Dox. Three animals were taken at each time point for

each of the three mouse lines [Tg(HQK), WT, and KO]. Temporally regulated genes in

the quadriceps of Tg(HQK) and KO, in comparison to WT, were identified using EDGE

(extraction and analysis of differential gene expression), a significance method for

analyzing time course microarray data (118,119). A Q value cut-off of 0.05, and a fold

change of 3 for at least one time-point, was the criteria used for the selection of

differentially expressed genes. In the muscles of Dox-treated Tg(HQK) mice, 1499

differentially expressed genes were identified in comparison with similarly treated, age-

matched WT mice; a cluster plot of al differentially expressed genes based on similarities

in their expression profiles is shown in Figure 2.1A. In contrast, only 13 genes showed

significant differential expression in the muscles of KO mice in comparison with

similarly treated, age-matched WT mice. To verify the expression of genes identified on

our cDNA array, and to sample a more complete set of genes covering the whole mouse

genome, we purchased additional microarrays from Agilent Technologies. These arrays

consisted of 44,000 oligonucleotide probes representing the whole mouse genome. We

re-examined the day 14 samples since the majority of the 1499 temporally deregulated

genes showed differential expression at this time point. A two-sided t-test of feature versus back-ground, set at a p value of 0.05, was used to obtain a list of genes whose log10 ratios were significant. This list was in good agreement with the data from the

Booth lab cDNA array, confirming the deregulation of almost two-thirds of genes

originally identified by the cDNA array, in addition a set of genes which were not

represented by probes on Booth lab cDNA arrays were identified. In total, 1265 selected

genes were annotated in the Ingenuity Pathway Analysis (IPA) database and are provided

as Additional file 1 (up-regulated) and Additional file 2 (down-regulated). A summary of

the most common biological functions and toxicity-related pathways associated with

these genes is shown in Figure 2.1B. Gene ontology analysis revealed that up-regulated genes were particularly enriched for gene involved in development, cell cycle regulation, programmed cell death, lipid metabolism and ion homeostasis (Table 1). Down-regulated gene ontology categories were enriched for genes involved in cellular energy metabolism, particularly carboxylic acid metabolism, protein metabolism and muscle developmental processes (Table 2).

PrPC Over-expression Regulates Multiple Targets with Established Roles in Myopathy

Many of the gene expression changes identified in the Tg(HQK) muscle are consistent

with the observed progressive atrophy, which is characterized by a decrease in myofiber

size and total muscle mass accompanied by a concomitant accumulation of lysosomes.

Table 1 List of genes belonging to some of the most significantly up-regulated Gene

Ontology Categories

Description Gene Name cell development ARF6, KIF5C, PRM2, NEB, SOX9, NDN, RUNX1, FCER1G, ENAH, PRKDC, GADD45G, PURB, METRN, BIRC5, TRADD, LGALS1, EPHB1, TRIM35, GPX1, STMN3, E2F2, NEFL, DEDD, RHOA, JMY, MAL, DCX, CASP14, UNC5B, BNIP1, CD28, GDNF, ITGB1BP3, ALS2CR2, NFKB1, TIMP1, CARD10, SEMA6A, DAB1, CHRNA1, UCHL1, TNFRSF12A, HSPA1A, MYOG, AKT1S1, PIP5K1C, TRIAP1, PMAIP1, MT3, SOCS2, GADD45B, ABI2, TNNT2, GSK3B, SGPP1, RPS6KB1, HIPK2, IGFBP3, PERP, PPP1R13B, CDK5R1, HOOK1, EDA2R, CTF1, EHMT2, ITGA3, SOX10, HIPK3, E2F1, BCL2L13, PURA, YBX2, IBRDC2, APP, BOK, TNP1, FAF1, PHLDA1, CAMK1D, CSPG4, DOCK1, FARP2, DIABLO, GDF11, ZFP91, PEG3, PTPRC, BAK1, RHOT1, NRAS, CDKN1A, NAB2, DAP3 cell cycle ANLN, CDC27, PRM2, TIMELESS, RB1, TACC3, SMARCB1, GADD45G, CDC14A, BIRC5, INCENP, CHES1, UHRF2, PDGFB, CGREF1, MIS12, E2F2, CDK6, PSMD2, JMY, CITED2, SUV39H2, PPP2R3A, CD28, ALS2CR2, PLK2, MERTK, CLASP1, CRKL, PRC1, TRIAP1, GADD45B, CPEB1, FOS, GSK3B, HIPK2, EHMT2, SPAG5, RANBP1, E2F8, E2F1, PLEKHO1, GAK, CCRK, PURA, HDAC5, RASSF4, APP, DHX16, E2F3, THPO, MKI67, BIN1, PTPRC, RGS2, ABL1, ANAPC1, NRAS, CDKN1A, JUNB, MDM2, ITGAE programmed cell death ARF6, SOX9, FCER1G, PRKDC, PURB, GADD45G, BIRC5, TRADD, TRIM35, GPX1, E2F2, NEFL, RHOA, DEDD, JMY, MAL, CASP14, GDNF, BNIP1, CD28, UNC5B, ALS2CR2, NFKB1, CARD10, SEMA6A, TNFRSF12A, HSPA1A, AKT1S1, PMAIP1, TRIAP1, GADD45B, SGPP1, GSK3B, IGFBP3, HIPK2, PERP, CDK5R1, PPP1R13B, EDA2R, HIPK3, E2F1, BCL2L13, PURA, IBRDC2, APP, BOK, FAF1, PHLDA1, CAMK1D, DOCK1, DIABLO, ZFP91, PEG3, PTPRC, BAK1, RHOT1, NRAS, CDKN1A, DAP3 cellular lipid metabolic process ISYNA1, PRKAA1, LCAT, NEB, SULT2B1, PIP5K1C, B4GALNT1, VLDLR, FDPS, SERINC2, SGPP1, LDLR, ADIPOR2, RDH11, SYK, CDS2, SNCA, PRKAG2, MYO5A, ELOVL6, HEXB, CDS1, CD81, BMPR1B, ST6GALNAC6, SOAT1, FADS3, PIP5K1A, CHKB, PIGO, ELOVL3, UGCG, AYTL2, SLC37A4, AGPAT3, PBX1, AGPAT2, SYNJ1, INSIG1, PIGK, HMGCS2, PRKAB2, ACBD3, CYB5R3, PISD, SERINC1, MTMR7, HEXA cellular ion homeostasis CHRNA1, APP, ATOX1, CHRNG, APLP2, SV2A, CHRNB4, MT3, NDN, RYR3, PRND, ATP2A2, PTPRC, BAK1, SLC37A4, MT4, SYPL2, ANXA7, PRNP, SLC39A5, HEXB

Table 2 List of genes belonging to some of the most significantly down-regulated Gene Ontology Categories

Description Gene Name

carboxylic acid metabolic process NR3C1, TARSL2, AHCY, SHMT2, PTGES3, LYPLA1, IRG1, MCCC1, PRKAG1, MAT2B, CYP39A1, MCFD2, IDH2, GLUL, IARS, PYCR2, FBP2, PLP1, ABAT, ADIPOR1, LYPLA2, BCKDHA, CROT, GPD2, CAV1, MTHFD1, SH3GLB1, ACADSB

protein metabolic process PPP3CB, BZW2, PPP2CB, AGA, CDC16, CHEK2, HERC2, UBE2B, PRMT7, NCKAP1, EIF4A2, CCT2, TLK2, KLK8, PDPK1, CSE1L, OMA1, UBQLN2, SLC30A9, PRKAR2A, HAT1, CAPZA2, CLK1, CPA3, LCK, CAMK2G, CAV1, MTM1, PSMB9, HUWE1, UBE2D1, PRKRIR, FKBP8, FKBP4, ARAF, RPS6KC1, PPP2R2A, VWF, CCT6A, GART, EPHA7, EIF3S6, WWP1, DVL1, IARS, ASPH, HTRA2, RNF6, RNF8, UBE2A, MCPT4, HS3ST5, CUL3, PCTK3, EEF2, UBE2G1, MMP13, UQCRC2, PRPF4B, AP3M1, BRCC3, SH3GLB1, RPL36, TARSL2, CDKL2, CAMK1, USP15, ULK2, BACE2, HECTD1, DNAJC12, ITGB4BP, CRY1, RAD21, FBXL5, DMD, MGRN1, RCHY1, IPO11, VBP1, USP1, VPS35, YME1L1, RPL22, COPB1, LGTN, GLMN, RSL1D1, RPL4, SUV420H2, ETF1, MAP2K5, USP38, EGLN1, TBCE, CUZD1, FURIN, PAIP1, CDC25B, EIF4G2, IFNAR1, TRIM23, CAV2, PSMD6, PIGY, LAP3

muscle development CSRP3, DMD, MYL2, TSC1, MYH7, CAV2, MTM1, CAV1, ACTC1, DVL1, ACTG2, MYH6, CACNA1S, MEF2C

Specific changes included a significant down-regulation of genes coding for the

myofibrillar proteinsMYH2, MYH6, MYH7, MYL2, MYL3, and an increase in

expression of the transcription regulator MDFI (MyoD Family Inhibitor) that acts as a

negative regulator of myogenic proteins, and induction of MyoG (myogenin), a muscle-

specific transcription factor that can induce myogenesis in a variety of cell types in tissue

culture. The MEF2c (Myocyte Enhancer Factor 2C) gene was also down-regulated in

Dox-induced Tg(HQK) muscles. Immunoblot analysis showed that there was statistically significant reduction of MEF2c protein level in the skeletal muscle from day14 of Dox

Figure 2.1

Clustering of gene expression data. (A) Measurements of relative gene expression for 6 time points (after 0, 4, 7, 14, 30, 60 days of Dox Treatment) in Tg(HQK) and PrP knockout mice (KO). Mice were treated with 6 g Dox/kg food, and three animals were taken at each time point as indicated. Total RNA was extracted from skeletal muscles (quadriceps) from the hind legs and subjected to microarray analysis, yielding expression profiles of genes with normalized expression ratios. Red and purple represent relative over-expression and under-expression, respectively, and the color intensity represents the magnitude of digression. (B) Bioinformatic analysis (Ingenuity Pathways Analysis) to determine the top biological functions and associated p values of the selected genes is shown. The top four categories are listed for diseases and disorders, molecular and cellular functions, physiological system and development and function, and pathways associated with toxicity.

Figure 2.2

MEF2c protein level is down-regulated in the skeletal muscles of Tg(HQK) mice treated with Dox. Tg(HQK) mice were treated with 6 g Dox/kg food for 0-60 days as indicated, and three animals were taken at each time point. Skeletal muscle (quadriceps) from the hind legs was subjected to immunoblot analysis in three blots. Fifteen micrograms of total proteins were loaded for each sample. Skeletal muscle (quadriceps) sample from an untreated WT FVB mouse serves as the control to normalize data from the triplicate blots. (A) A representative immunoblot probed with anti-MEF2c antibody followed by probing with an anti-actin antibody after stripping. (B) Plot of the MEF2c protein levels over increasing duration of Dox treatment. The MEF2c protein level for each sample was normalized against the actin level in each blot and expressed as the ratio against the normalized MEF2c protein level in the untreated WT FVB mouse on the same blot. The error bars denote standard errors calculated from the three blots. The bars with asterisk(s) indicate a statistically significant difference when compared to the 0 day Tg(HQK) samples. *p<0.05; **p<0.001.

treatment, and the reduction reached 50% after 30 days of Dox treatment (Figure 2.2).

MEF2c has been studied extensively in muscle cells. It is a key regulatory transcriptional

factor that plays an essential role in the transcription control of muscle development as

well as remodeling of adult muscles in response to physiological and pathological signals

(120,121). It has been reported that MEF2c directly activates the expression of a muscle

specific protein kinase Srpk3 and Srpk3-null mice exhibit widespread centronuclear

myopathy via an unknown mechanism (122). We speculate that the down-regulated

MEF2c gene expression might play a role in the progressive central nucleus localization

observed in the skeletal muscles of Dox-treated Tg(HQK) mice (34) through a reduction

of the Srpk3 activity.

A number of lysosomal peptidases were up-regulated including CTSS, CTSD, CTSZ and

DPEP2, coincident with an observed accumulation of lysosomes in Tg(HQK) mice over-

expressing PrPC (34). The gene CTSL, which codes for a lysosomal cysteine proteinase, is commonly used as a universal marker for muscle atrophy but was not represented on our arrays (123). qRT-PCR revealed expression of this gene was induced transiently following PrPC induction, peaking at 7 days following the onset of Dox treatment and

returning to the baseline by 60 days post-induction. The genes encoding lysosomal

proteins HEXA, HEXB and LAMP1 were also up-regulated at late time points.

Previous studies have shown that the development of muscle atrophy in a number of

models of systemic wasting states (fasting, cancer cachexia, renal failure , diabetes) an in

disuse atrophy induced by denervation or spinal cord isolation follows a common

program of transcriptional changes (124,125). One of the main features of this program is

a general increase in expression of genes involved in proteolysis including both

lysosomal proteases, and an ATP-dependent process requiring ubiquitin and the proteasome. The degradation of PrPC and PrPSc is also believed to involve the proteasome

(126), and compromised/inhibited proteasome activity was proposed to lead to

accumulation of cytosolic PrPC that is neurotoxic (126); but the latter notion has been

challenged (127,128). Following induction of PrPC we observed that the expression levels

of genes involved in proteasomal protein degradation were for the most part unchanged.

Out of the 44 unique proteasome related genes represented on the microarrays, only three

(PSMD2, PSMD4, PSMD7) were up-regulated and four (PSMD6, PSMD12, PSMD13 and PSMD14) were down-regulated.

A further feature reported in a number of different models of diseases resulting in muscle atrophy is the substantial up-regulation of two E3 ubiquitin ligases, atrogin-1/MAFbx

(FBXO32) and MuRF1 (TRIM63) (129,130), which are generally induced early during the atrophy process. Upon fasting, the rise in atrogin-1 expression precedes the loss of muscle mass, conversely, deletion of either Atrogin-1 or MuRF1 has been shown to significantly alleviate muscle atrophy (130). Our microarray data did not reveal a significant increase in Atrogin-1 expression in the Tg(HQK) atrophy model and no probe for MuRF-1 was present on either of our array platforms. qRT-PCR was used to determine the expression levels of these two genes, and a small, less than 3-fold increase in the expression of both MuRF1 and Atrogin-1 was detected following induction of PrPC

(Figure 2.3A and 2.3B); this is much lower than the 10-40 fold increase generally found

in other models of muscle atrophy. In a recent study FOXO protein (a key activator of

atrophy) as well as the fall in PGC-1 alpha and beta (transcriptional corepressors of

Figure 2.3

Real-time PCR analysis of Atrogin-1 and MuRF1. qRT-PCR analysis of Atrogin-1 (A) and MuRF 1 (B) gene expression in RNA samples from Tg(HQK) mice relative to similarly treated WT control mice. Measurements of relative gene expression for 4 time points (over 4-60 days) in mice following treatment with 6 g Dox/kg food beginning on day 0. Total RNA was extracted from skeletal muscles (quadriceps) from the hind legs and subjected qRT-PCR analysis. Results represent the mean ± s.e.m. of triplicate measurements performed. *p<0.01; **p<0.001.

Atrogin expression) were identified in numerous types of muscle wasting (131,132). A 3- fold decrease of FOXO1 and no change in expression of PGC-1 alpha and beta were detected in Dox-treated Tg(HQK) mice. These data suggest only minor involvement of

the ubiquitin-protease proteolysis pathway in the observed muscle atrophy and a program

of transcriptional changes that is not reminiscent of systemic wasting states.

Activation of Multiple Significant Signaling Pathways Following PrPC Induction in

Skeletal Muscle of Tg(HQK)

The dramatic transcriptional response to PrPC over-expression in the muscles of Tg(HQK)

mice lacks key features of the common transcriptional program indicative of several

reported forms of muscle atrophy. This includes striking de-regulation of over 400 genes

involved in cell death and regulation of the cell cycle, which suggests a toxic effect of the

over-expressed PrP. Using the Ingenuity Pathway Analysis (IPA) tool, we identified

many pathways invoked in response to PrPC over-expression, among which the p53

signaling pathway scored highly with a p value of 1.27 ×10-7. Other molecular pathways

Table 3 List of genes belonging to some of the most significantly de-regulated pathways that have been implicated in toxicity-associated biological processes as resulted by Ingenuity Pathway Analysis

Toxicity-Associated P-value Genes Process P53 signaling 1.27 X 10-7 BBC3, BIRC5, C12ORF5, CDKN1A, CHEK2, E2F1, GADD45B, GADD45G, GSK3B, HIPK2, PERP, PIK3R5, PMAIP1, PPP1R13B, PRKDC, RB1, TP63, TP53INP1

G1/S transition of the cell 1.53 X 10-7 ABL1, CCNE2, CDK6, CDKN1A, cycle E2F1, E2F2, E2F3, E2F6, GSK3B, HDAC5, RB1, RBL2, SIN3A

Mitochondrial dysfunction 1.04 X 10-5 AIFM1, APP, BACE2, COX6B2, CYB5R3, GPD2, GSR, HTRA2, NDUFAF1, NDUFB5, OGDH, PRDX3, SDHA, SDHB, SNCA, SOD2, UQCRC1, UQCRC2, UQCRFS1

-5 Oxidative Stress 4.11 X 10 FOS, GPX1, GSR, GSTA5, GSTM2, GSTM1, NFKB1, NFKB2, PRDX3, SOD2, STAT3

that scored significantly were the related G1/S transition of the cell cycle (p=1.53 × 10-7),

mitochondrial dysfunction (p = 1.04 ×10-5) and oxidative stress response (p = 4.11 ×

10-5) (Table 3).

The involvement of the p53 signaling pathway was of particular interest as mounting

evidence suggests that over-expression of PrPC sensitizes cells to apoptotic stimuli

through a p53-dependent pathway (49, 81-83). The p53 gene itself did not meet our

selection criteria (a change of 3-fold or more in at least one time point) as significantly

deregulated in the microarray analysis; however qRT-PCR showed it to be marginally

Figure 2.4

Real-time PCR analysis of mdm2 and p53. qRT-PCR analysis of mdm2 (grey) and p53 (black) gene expression in RNA samples from Tg(HQK) PrP over-expressing mice relative to similarly treated WT control mice. Measurements of relative gene expression for 5 time points (over 4-60 days) in mice following treatment with 6 g Dox/kg food. Total RNA was extracted from skeletal muscles (quadriceps) from the hind legs and subjected qRT-PCR analysis. Results represent the mean ± s.e.m. of triplicate measurements performed. **p<0.01; ***p<0.001.

up-regulated from day 7 following the onset of PrPC induction. This transient over-

expression was low, approximately 1.5-2.5 fold, but statistically significant in all

Tg(HQK) mice tested (Figure 2.4).

Deregulation of Genes Involved in p53-Dependent G1 Cell Cycle Arrest and Apoptosis

Systematic examination of the genes differentially expressed following PrPC over

expression revealed over 60 genes that were annotated, or cited in PubMed, as being p53

responsive genes. We used the IPA tool to build a network of potential regulatory

interactions between the products of these genes; the resulting network is shown in

Figure 2.5. The genes making up this network are primarily involved in the regulation of

the cell cycle and cell death. A number of these are transcription factors including the

proinflammatory regulator NF-κB which has been shown to be activated in degenerating

muscle of Duchenne muscular dystrophy patients and dystrophin-defficient mouse

models (133-135). Two products of up-regulated genes induced in Tg(HQK) muscle,

CDNK1A (cyclin-dependent kinase inhibitor , p21) and GADD45B (growth arrest and

DNA-damage inducible, beta), stand out as crucial to the initiation of cell cycle mediated

by activated p53. p53 tightly controls the expression of CDNK1A, which mediates the

p53-dependent cell cycle arrest at the G1 phase by binding to and inhibiting the activity

of cyclin-CDK2 or cyclin-CDK4 complexes in response to a variety of stress stimuli.

Expression of CDNK1A was confirmed by qRT-PCR to be increased by more than 20-

fold over that in control WT mice at 30 days post induction. The up-regulation of

GADD45A, closely related in function to GADD45B, was also confirmed by qRT-PCR.

These genes are often coordinately expressed and can function cooperatively to inhibit

Figure 2.5

Upregulation of the p53 pathway: Analysis using the Ingenuity Pathway Knowledge Base (IPKB). This figure illustrates potential functional relationships of TP53 responsive genes de-regulated in the muscles of Dox-treated Tg(HQK) mice. Direct (solid lines) and indirect (dashed lines) interactions reported for these genes (grey shading) in the IPKB database. Color shading corresponds to the type of de-regulation, red for up-regulated genes, and green for down-regulated genes. The shape of the node indicates the major function of the protein (see key), and a line denotes binding of the products of the two genes while a line with an arrow denotes ‘acts on’.

cell growth and induce apoptosis. Other up-regulated genes known to play a role in cell- cycle arrest are RB1,which binds to E2F transcription factors to prevent transcription of genes required for the G1 to S phase transition, and CGREF1, which is produced in response to stress and serves as a negative regulator of the cell cycle (136). Taken together these gene expression changes indicate p53-dependent G1 cell cycle arrest was

C induced in Tg(HQK) muscle following induction of PrP expression.

Following cell cycle arrest, cells either recover or undergo p53-mediated apoptosis due to

transcriptional activation of a number of pro-apoptotic genes. Key transducers of

apoptosis include PMAIP1 (phorbol-12-myristate-13-acetate-induced protein 1 or Noxa)

(137,138) and BBC3 (BCL2 binding component 3 or PUMA) (139,140). Both were significantly up-regulated based on our microarray analysis. PMAIP1 induces the expression of other death effectors including BAK1 (141,142) that was also significantly induced in Dox-treated Tg(HQK) muscles. Deregulation of other apoptosis effector genes includes induction of the pro-death genes BOK1 and the down-regulation of MCL1, a pro-survival BCL2 homologue. Numerous studies have identified the pro-apoptotic regulator BAX to be a major mediator of p53 induced apoptosis (143). BAX was not identified as up-regulated by our microarray analysis because of the high cut-off value (>

= 3-fold), but qRT-PCR revealed a modest up-regulation of the BAX gene (1.5-3.0 fold) over time following PrP over-expression. Similar to p53, TP73L (p63) can mediate apoptosis and was also found to be induced in atrophic muscles of Tg(HQK) mice. Less is known about the regulatory pathways triggered by p63 and its transcriptional targets have not been fully characterized (144-147). Moreover, both the p53 apoptosis effector gene PERP and the p53-inducible ubiquitin ligase p53RFP (RNF144B) were significantly

induced in the Tg(HQK) muscles as well. RERP is a potential marker of p53 driven

apoptosis since it has been found to be induced in p53-driven apoptotic cells but not in

p53-dependent G1 arrest cells and p53RFP has also been shown to be involved in

switching a cell from p53-mediated growth arrest to apoptosis (148,149).

These data indicate that not only do muscle cells of Dox-treated Tg(HQK) mice undergo p53-dependent cell cycle arrest, but at least in some instances they go on to undergo apoptosis, which strongly suggests that p53-regulated pro-apoptotic pathways play an important role in PrP-mediated myopathy.

ADAM8, a member of the ADAMs family, was also upregulated, which will be studied in detail in Chapter 4.

DISCUSSION

Our lab has previously described the generation of the Tg(HQK) transgenic mice, in

which Dox-induced over-expression of PrPC specifically in skeletal muscles cause a

primary myopathy that is correlated with accumulation of the N-terminal truncated PrP

C1 fragment (34). The aim of this study was to determine the molecular basis for the PrP- mediated myopathy by microarray analysis. The ultimate goals are to fully understand the detailed molecular pathways of PrP-mediated myopathy, so that we can better understand the role of PrP in both normal and diseased muscles and provide some clues on the pathogenic mechanism of prion diseases. Utilizing two DNA microarrays, we identified more than 1000 genes that were temporally deregulated in a specific and highly consistent manner following induction of PrPC over-expression in the muscles of

Tg(HQK) mice and the subsequent development of myopathy. The transcriptional

profiles in the muscles of Dox-treated Tg(HQK) mice strongly implicate toxicity-induced pro-apoptotic pathways in PrP-mediated myopathy, and they are quite different from the changes previously described in systemic, disuse, and denervation muscle atrophy.

Interestingly, the transcription factor MEF2c was found to be down-regulated at both the mRNA and protein levels in PrPC-mediated myopathy. MEF2c is expressed specifically

in muscle and brain, where it is a target for signaling pathways involving calcium (150).

MEF2c regulates the expression of a majority of muscle-specific genes, and interacts with members of the MyoD family of proteins to activate muscle differentiation (120).

Calcium signaling was one of the pathways significantly induced in Dox-treated Tg(HQK)

mouse muscles as evidenced by a very small p value of 8.75 × 10-9. The PrPC protein has

itself been shown to play a role in Ca2+ homeostasis (151-153) and it is possible that

over-expression of PrPC results in perturbations in Ca2+ signaling, which in turn

modulates the activity and/or expression of MEF2c. As calcium regulation has also been

shown to be altered during prion-induced neurodegeneration, this finding potentially links the molecular changes occurring in Tg(HQK) myopathy to be pathobiology of prion diseases.

The most striking finding is the deregulation of the genes involved in the regulation of the cell cycle and cell death. However, we did not have any functional data supporting that there is indeed cell cycle arrest or cell death of muscle cells in Dox-induced Tg(HQK)

mice. Thus, it is necessary to perform flow cytometry assay to examine cell cycle

transition and TUNEL staining to monitor cell death. Furthermore we find that there is

the strong and statistically highly significant induction of a p53-regulated pro-apoptotic

network in Tg(HQK) mouse muscles following induction of PrPC. Central to this network

are induction of p53 protein expression and strong induction of genes responsible for

arresting the cell cycle, as well as a number of p53-regulated pro-apoptotic (up-regulated)

and antiapoptotic (down-regulated) genes. p53 is a critical tumor suppressor and

transcription factor, and it has been linked to cell death in the central nervous system in a

number of disorders including most notably neurodegenerative disorders such as

Alzheimer's disease and prion diseases (70,154,155). The expression of p53 protein has

been found to rapidly increase in neurons in response to a range of insults including DNA

damage, oxidative stress, metabolic compromise, and cellular calcium overload. Over-

expression of PrPC has been shown to enhance staurosporine-induced toxicity and

activation of caspase-3 in the HEK293 kidney cell line (156) and increase sensitivity to

apoptotic stimuli via p53-dependent pathways in TSM1 neuronal cell line (49).

Conversely neurons devoid of PrPC expression were reported to display lower

responsiveness to staurosporine, also via p53-dependent pathways (83).

One of the main pro-apoptotic effectors of p53 is BAX, which plays a major role in

regulating neuronal death in the brain in response to a number of stimuli (157,158). The

role of BAX in prion-induced neurodegeneration is not well understood; both BAX-

dependent and BAX-independent mechanisms appear to underlie the action of neurotoxic

forms of prion proteins (159). However, in the muscle of Dox-treated Tg(HQK) mice,

only a marginal increase in BAX expression was observed whereas significant over-

expression of other p53 regulated pro-apoptotic proteins, including BAK1, BBC3 and

PMAIP1, and MCL1, were detected, suggesting that PrPC-mediated myopathy observed in this model may depend on Bax-independent pathways that involve BAK1, BBC3,

PMAIP1, and MCL1.

CHAPTER 3

p53 Plays a Critical Role in PrPC–Mediated Myopathy

ABSTRACT

We have reported that Dox-induced over-expression of wild type prion protein (PrP) in skeletal muscle of Tg(HQK) mice is sufficient to cause a primary myopathy with no signs of peripheral neuropathy. The preferential accumulation of the truncated PrP C1 fragment was closely correlated with these myopathic changes. We have also reported up-

regulation of genes for p53-related pathways involved in cell cycle arrest and promotion

of apoptosis in skeletal muscles of Dox-induced Tg(HQK) mice. Here we show that

prominent up-regulation of p53 protein paralleled the initiation and progression of the

muscle pathology of Tg(HQK) mice, and treating Dox-induced Tg(HQK) mice with

pifithrin-α (PFT-α), a proven specific inhibitor of p53, largely blocked PrPC-mediated

myopathy in Dox-induced Tg(HQK) mice via inhibiting p53 downstream signaling

pathways. These results demonstrate that p53 and p53 pathway plays a critical role in

PrPC-mediated myopathy in Tg(HQK) mice.

INTRODUCTION

The p53 tumor suppressor protein is a potent transcription factor that is activated in

response to DNA-damaging agents, including ultraviolet (UV) radiation (160-163).

Following activation, p53 regulates a change in the balance of gene expression leading to

growth arrest or apoptosis, and these effects are thought to prevent the proliferation of

genetically damaged cells (63,164). p53 has been found to have multiple anti-cancer

mechanisms: activating DNA repair proteins when DNA has sustained damage; arresting

the cell cycle at the G1/S transition point on DNA damage recognition; and initiating

apoptosis or programmed cell death when DNA damage proves to be irreparable (62-64).

Recently, several reports showed that p53 may be involved in neurodegenerative process and contributed to neurodegenerative diseases (77-79). There have been a few reports of apoptotic cells in the brains of animals and humans with prion diseases and have further suggested the activation of the p53 pathway for apoptosis (80). In terms of the relationship between p53 and PrPC, several reports focused on studying the role of p53 in

PrPC overexpression induced cell death in several neuronal cell lines. It was established

that PrPC overexpression in transfected or inducible TSM1 cells led to increased cell

death (81), and depletion of endogenous PrP reduces susceptibility to staurosporine-

induced caspase 3 and p53 activation (83). Besides, over-expression of PrPC had a protective effect in BAX and TNFα-mediated cell death and conversely a pro-apoptotic

function in staurosporin-induced cell death (83,102,103).

We previously generated the Tg(HQK) mice that express human PrPC exclusively in the skeletal muscles under tight regulation by Dox (34). Induced over-expression of PrPC in

the muscles of Tg(HQK) mice led to a progressive primary myopathy characterized by

increased variation of myofiber size, centrally located nuclei and endomysial fibrosis, in

the absence of cytoplasmic inclusions, rimmed vacuoles, or any evidence of a neurogenic

disorder (34). While the pathogenic mechanism of the PrPC-mediated myopathy was not

determined, an interesting observation was that the myopathy was accompanied by

preferential accumulation of PrP C1 fragment (34). A number of studies have shown the expression of N-terminus truncated forms of PrPC is associated with toxicity in animal

models (108,109). In addition, it has been showed that overexpression of PrP C1

fragment potentiates staurosporine-induced caspase-3 activation through a p53-dependent

mechanism, indicating that C1 positively controls p53 transcription and mRNA levels

and increases p53-like immunoreactivity and activity (49). Thus, it is highly possible that

PrPC-mediated myopathy in Tg(HQK) mice is at least partially a result of C1

accumulation in the skeletal muscles.

We have shown in Chapter 2 that the PrP-mediated myopathy in Tg(HQK) mice was accompanied by activation of p53 signaling pathway and deregulation of several genes involved in p53-dependent G1 cell cycle arrest and apoptosis, which strongly suggests

that p53-regulated pro-apoptotic pathways play an important role in PrP-mediated

myopathy. Here we further evaluate the role of p53 pathway in Dox-induced Tg(HQK) myopathy by examining changes in total p53 protein level in skeletal muscles during Dox treatment as well as the effect of a p53 chemical inhibitor on the progression of myopathy in the Tg(HQK) mice.

METHODS

Animals

The doxycycline-inducible Tg(HQK) mice were described previously (34). The HQK

transgene contained two genes: reverse tetracycline responsive transcription activator

(rtTA) under the control of the mouse PrP promoter of the half genomic PrP clone, and human PrP ORF regulated by the tetracycline-responsive promoter (tetO-hCMV*-1) from

the core plasmid (115). The Tg(HQK) mice were generated in the FVB background, and

Tg(HQK)/Prnp0/0 mice were obtained through breeding with the Zurich I PrP-null mice

(60) in FVB background. Line Tg(HQK) 18, referred to as Tg(HQK) for simplicity, was

used for this study. Wild type (Wt) FVB-NJ mice (Jackson Laboratory) were used as

controls.

Animal Treatment and Specimen Collection

Age- and weight-matched female Tg(HQK) and Wt mice (15 weeks old, 22~24 g) were

treated as follows. For Dox treatment, the animals were fed food pellets either lacking or containing 6 g Dox/kg food (Bio-Serv) to induce PrPC expression. For pifithrin-α (PFT-α)

treatment to inhibit p53 apoptotic signaling, the animals were given water either lacking

or containing 2 µg/ml PFT-α (Sigma-Aldrich, St. Louis, MO). Body weight was

measured weekly and grip strength assay were performed biweekly. For each time point

and each treatment, 7 mice were sacrificed, hind limb skeletal muscles (quadriceps)

dissected and muscle mass measured. For each mouse, left hind limb (quadriceps) and

left forelimb skeletal muscles were collected for histopathology analysis; right hind limb

and right forelimb skeletal muscles were collected for protein analysis by

immunoblotting. For immunoblot analysis, the skeletal muscles from the quadriceps of

hind limbs were removed at day 0, 4, 7, 14, 30 and 60 days following administration of

Dox and the muscle tissues were immediately frozen on dry ice, and stored at -80°C.

Immunoblot Analysis

Mouse skeletal muscle tissues were homogenized in lysis buffer containing 50 mM Tris

(pH7.5), 200 mM sodium chloride, 0.5% sodium deoxicholate, and 5 mM EDTA. Protein concentrations were determined by the BCA protein assay (Pierce). After addition of

LDS sample buffer (Invitrogen) and sample reducing agents (Invitrogen), the homogenates were denatured at 100°C for 10 minutes, and the proteins were resolved on

10% NuPage Tris-Bis Gels (Invitrogen) and blotted onto nitrocellulose membranes

(Invitrogen). For p53 protein detection, the membrane was incubated with a monoclonal anti-p53 antibody that detects total p53 proteins (Cell Signaling) (1:2000 diluted in 5% milk, 1×TBS, 0.1% Tween-20) at 4°C with gentle shaking overnight. For p21 detection, the membrane was incubated with a rabbit polyclonal anti-p21 antibody (Cell Signaling)

(1:2000 diluted in 5% milk, 1×TBS, 0.1% Tween-20) at 4°C with gentle shaking overnight. The blots were developed with the Immobilon Western Chemiluminescent

HRP substrate (Millipore) according to the manufacturer’s instructions. Skeletal muscle actin was similarly probed with a rabbit polyclonal antibody (Abcam) (1:5000 diluted in

0.5% normal goat serum, 1×TBS, 0.1% Tween-20) after stripping with a stripping buffer

containing 1.4% 2-mercaptoethanol, 2% SDS and 62.5 mM Tris (pH 6.8). The Western blots were exposed to X-ray films, which were scanned and bands quantified with the

UN-SCAN-IT gel 6.1 software (Silk Scientific).

Grip Strength Test

Grip strength was tested using a commercially available strength meter apparatus (San

Diego Instruments, San Diego, CA) that measures the gripping strength of mice. The system has two grids, one horizontal to measure forelimb strength and one at 45○ angle to measure hind limb strength, both connected to a force sensor. For forelimb measurement, mice were gently lowered over the top of the grid so that only its front paws could grip the grid and then pulled back steadily until the grip is released down the entire length of the grid; for hind limb measurement, the mouse were gently held by the scruff of the neck, placed on the angled grid and slided from bottom to top until the grip is released completely. Maximal strength (in kilograms) was recorded 5 times per animal.

RESULTS

Up-regulation of Total p53 Protein Following PrPC Induction in Skeletal Muscles of

Tg(HQK) Mice

We have described in Chapter 2 qRT-PCR analysis of mRNA levels for p53 and MDM2 genes in skeletal muscles of Dox-induced Tg(HQK) mice. Our results showed that there was 1.5 -2.5 fold increase in p53 mRNA level at day 7 of Dox treatment. However, regulation of p53 is known to take place mostly at the level of translation (132). In accordance with this, immunoblot analysis of skeletal muscle (quadriceps) of Dox-treated

Tg(HQK) mice showed that the p53 protein level started to increase at day 7 and reached over 3-fold at days 30-60 when compared with age-matched Wt controls (Figure 3.1A-B).

This result confirms that the p53 protein is up-regulated in the skeletal muscles of Dox- treated Tg(HQK) mice.

Inhibition of p53 Apoptotic Signaling Reduces PrP-mediated Myopathy in Tg(HQK)

Mice

To evaluate whether p53 plays a functional role in PrP-mediated myopathy in Tg(HQK) mice, we fed age-matched female Tg(HQK) and Wt mice with Dox-laced food pellets and drinking water containing 2µg/ml PFT-α. PFT-α is a specific chemical inhibitor of p53 that has been reported to reversibly block p53-dependent transcriptional activation and apoptosis and protect against the lethal genotoxic stress associated with anticancer treatment without promoting tumor formation (165). We measured the body weight weekly and the mass of hind limb quadriceps biweekly. The results showed that Dox and

PFT-a treatments had no detectable effect on Wt mice. For Tg(HQK) mice, Dox

Figure 3.1

Total p53 protein level is up-regulated in the skeletal muscles of Tg(HQK) mice treated with Dox. Tg(HQK) mice were treated with 6 g Dox/kg food for 0-60 days as indicated, and three animals were taken at each time point. Skeletal muscles (quadriceps) from the hind legs were subjected to immunoblot analysis in three blots. Twenty micrograms of total proteins was loaded for each sample. Skeletal muscles (quadriceps) samples from an untreated Wt FVB mouse serves as the control to normalize data from the triplicate blots. (A) A representative immunoblot probed with anti-p53 antibody followed by probing with an anti-actin antibody after stripping. (B) Plot of the total p53 protein level over increasing duration of Dox treatment. The p53 protein level for each sample was normalized against the actin level in each blot and expressed as the ratio against the normalized total p53 protein level in the untreated Wt FVB mouse on the same blot. The error bars denote standard errors calculated from the three blots. The bars with asterisk(s) indicate a statistically significant difference with compared to the 0 day Tg(HQK) samples. *p<0.05; **p<0.01; ***p<0.001. treatment alone led to significant reduction in both body weight and skeletal muscle mass starting from day 14; in contrast, with Dox and PFT-α treatment, the decrease in body

weight and skeletal muscle mass were largely prevented, and the difference between

PFT-α treated and untreated animals becomes statistically significant from day 14 (Figure

3.2A-B).

Figure 3.2

PFT-α attenuates PrP-mediated myopathy in Dox-treated Tg(HQK) mice. Age and weight-matched female Tg(HQK) and Wt mice were treated with 6 g Dox/kg food and water that was with or without 2 µg/ml PFT-α. Body weight was measured weekly; grip strength of both front and hind limbs as well as muscle mass (from hind limb quadriceps) were measured biweekly. Seven animals were taken at each time point with each treatment. (A) Plot of body weight over increasing duration of Dox treatment. (B) Plot of the mass of quadriceps over increasing duration of Dox treatment. (C) Plot of highest front limb grip strength over increasing duration of Dox treatment. (D) Plot of highest hind limb grip strength over increasing duration of Dox treatment. *p<0.05; **p<0.01.

We also performed grip strength assays on both front and hind limbs bi-weekly to evaluate the effects of PFT-α treatment on muscle strength. For Wt mice, Dox and PFT-α double treatments had no significant effect. For Tg(HQK) mice, Dox treatment alone resulted in modest but significant reduction in the strength of both front and hind limbs at day 14, which got worse at day 28; in contrast, with Dox and PFT-α double treatments,

Figure 3.3

PFT-α inhibits p21 up-regulation without affecting p53 protein level in skeletal muscles. Age and weight-matched female Tg(HQK) and Wt mice were fed with Dox-laced food pellets and drinking water containing 2µg/ml PFT-α. Seven animals were taken at each time point with each treatment. Skeletal muscles (quadriceps) from the hind legs were collected, homogenized, and subjected to SDS-PAGE and probed with anti-p53 and anti-p21 antibodies. (A,B) Representative immunoblots probed with p53 or p21antibodies followed by probing with anti-actin antibody after stripping. (C,D) Plot of p53 or p21 protein level in skeletal muscles upon Dox and PFT-α treatment. The p53 and p21 protein level for each sample was normalized against the actin level in each blot and expressed as the ratio against the normalized total p53 or p21 protein level in the untreated Wt FVB mouse on the same blot. The error bars denote standard errors calculated from the seven samples of each treatment. The bars with asterisk(s) indicate a statistically significant difference with compared to the 0 day Tg(HQK) samples.*p<0.05; **p<0.01 the decrease in muscle strength for both front and hind limbs were significantly reduced

(Figure 3.2C-D). These results indicate that PFT-α largely prevented PrP-mediated myopathy in Tg(HQK) mice.

PFT-α Inhibits p21 Up-regulation without Affectting p53 Protein Level in Skeletal

Muscles

To further study the molecular mechanism underlying the effects of PFT-α on PrP- mediated myopathy, we examined in the skeletal muscle protein levels of both p53 and p21, the latter is a p53-responsive cell cycle regulator (Fig. 3.3). For Wt control mice, total p53 protein level and p21 protein level both appear to decrease slightly with Dox treatment, and additional PFT-α treatment made no difference (Fig.3.3). For Tg(HQK) mice, Dox treatment led to significant increase of both total p53 and p21 protein levels; in contrast to Wt mice, additional PFT-α treatment did not affect the rise in total p53 protein level but largely blocked the increase of p21 (Fig. 3.3).

DISCUSSION

PrP level in muscle has been demonstrated to be elevated in patients with several sporadic and hereditary neuromuscular diseases (55,106), but it is not clear whether the elevated level of PrPC in skeletal muscle is a cause or result of muscle diseases in the affected patients and whether the elevated PrPC accumulation in the muscles is sufficient to cause muscle diseases (34). Our lab generated the Dox-inducible Tg(HQK) mice and showed that over-expression of wild type PrPC in skeletal muscles alone is sufficient to cause a progressive primary myopathy (34). As described in Chapter 2, we have also shown by microarray and qRT-PCR analysis that p53-regulated pro-apoptotic pathways were activated the skeletal muscles of Dox-treated Tg(HQK) mice.

In this Chapter, we showed that total p53 protein level was significantly elevated in the muscles of Dox-treated Tg(HQK) mice from day 7 of Dox-treatment, correlating well with the observed initiation and progression of myopathic changes. To evaluate whether p53 plays a functional role in PrP-mediated myopathy, we utilized PFT-α, which was reported to inhibit p53-dependent transcriptional activities and p53-mediated apoptosis

(165) and widely used to inhibit p53 activity in vitro and in vivo. Through assaying for body weight, muscle mass, and grip strength, we showed that the progression of PrP- mediated myopathy in Dox-induced Tg(HQK) mice was significantly attenuated by PFT-

α treatment. We also found that PFT-α administration did not affect the up-regulation of total p53 protein level due to PrP overexpression, but greatly reduced the rise in p21 protein level. PFT-α inhibition of p53 activity was reported to act downstream of p53 by modulating nuclear import/export of p53 or decreasing the stability of nuclear p53 (165), and p21 is one of the most important effectors of p53 (86). Therefore, our results suggest

that p53 plays a direct and critical role in PrP-mediated myopathy, which can be largely

blocked by PFT-α treatment through inhibiting p21 and maybe other p53-regulated effectors.

PFT-α has been reported to inhibit p53-dependent lacZ transcription activity and p53- mediated apoptosis (165), and has been widely used as chemical inhibitor to inhibit p53 activity in vitro and in vivo. However, no inhibitor is 100% specific, especially in vivo.

This is true for PFT-α. In fact, in JB6 cells, PFTα was shown to be an activator of the p53-induced apoptosis (166). Thus, examination of Tg(HQK)/p53-/- mice is needed.

In summary, these findings further our understandings on PrP-mediated pathogenic

processes in skeletal muscles and maybe other tissues. To my best knowledge, this is also

the first in vivo evidence that directly links p53 to a wild type PrP-mediated disease. A

recent report showed that over-expression of the C1 fragment increased cell death and

caspase-3 activity through a p53-dependent mechanism in cell culture models (49).

Combined with our earlier findings that the C1 fragment was preferentially accumulated

in the muscles of Dox-treated Tg(HQK) mice, we hypothesize that PrPC overexpression

somehow leads to C1 accumulation, which in turn activates the p53 pathways, thereby

resulting in the observed primary myopathy.

CHAPTER 4

Cellular Prion Protein Regulates Its Own Cleavage through ADAM8 in Skeletal Muscle

ABSTRACT

The ubiquitously expressed cellular prion protein (PrPC) is not only the central factor in prion diseases but also implicated in physiological processes and in other diseases such as

Alzheimer’s disease and cancer. PrPC is subjected to the physiological α-cleavage at a region critical for both PrP toxicity and the conversion of PrPC to its pathogenic prion form (PrPSc), generating the C1 and N1 fragments. The C1 fragment can activate caspase

3 while the N1 fragment is neuroprotective. A recent article indicates that ADAM10,

ADAM17 and ADAM9 may not play a prominent role in the α-cleavage of PrPC as previously thought, raising questions on the identity of the responsible protease(s). We have previously reported that muscular accumulation of the C1 fragment is correlated with a progressive myopathy due to over-expression of wild type PrPC in skeletal muscles in an inducible transgenic mouse model, but nothing is known about the protease(s) that generates the C1 fragment in skeletal muscles. Here we show that, in myoblast cell line

C2C12 and skeletal muscle tissues of transgenic mice, the PrP C1/full length ratio is linearly correlated with active ADAM8 protein level, indicating that ADAM8 plays a major role in the α-cleavage of PrPC in muscle cells. In addition, we found that overexpression of PrPC led to upregulation of ADAM8, suggesting that PrPC may regulate its own α-cleavage through modulating ADAM8 activity.

INTRODUCTION

The cellular prion protein (PrPC) is a ubiquitous glycosylphophatidylinositol (GPI)-

anchored glycoprotein that is highly expressed in the nervous system (25). PrP is the

central factor in prion diseases, a group of fatal neurodegenerative disorders characterized

by severe neuronal dysfunction and loss, spongiosis and accumulation of pathogenic

prion protein (PrPSc) that is converted from PrPC (4). PrP is also implicated in

Alzheimer’s disease (167-169) and cancer (170). The physiological roles of PrPC remain

elusive, but many normal functions have been proposed for PrPC (170-173). These

include signal transduction (14), resistance to oxidative stress, anti-apoptosis,

neuroprotection (21), cell adhesion, neurogenesis, axonal growth, neurite outgrowth and

neuritogenesis, neuronal differentiation, hematopoietic stem cell self renewal,

lymphocyte activation, and metal ion trafficking (174,175).

PrP is differentially cleaved in normal and prion-affected brains. In the brains of

Creutzfeldt-Jakob disease subjects (38,42) and prion-affected animals (176,177) and in

prion-infected cells (178), PrP is cleaved around the end of the octapeptide repeats

(termed β-cleavage) to generate the C2 and N2 fragments (38,179-181). β-cleavage

preserves the cytotoxic and fibrillogenic PrP106-126 core that is also critical for the

conversion of PrPC to PrPSc (43,45,182,183). β-cleavage of PrP is mediated by reactive oxygen species in CHO cells (184) and human neuroblastoma SH-SY5Y cells (185). The identity of cellular proteases involved in C2 production remains unclear. Calpain was shown to be critical for C2 production in scrapie-infected mouse brain cells (178), and

the contributions of cysteine proteases seem to depend on the cell models (186-188).

Both C2 and N2 fragments appear to be biologically inert (49,189). Nevertheless, the β-

cleavage of PrPC was reported to be critical for the anti-oxidative and neuroprotective

effect of PrPC (185,190). PrPC can also be cleaved and shedded directly by ADAM10 and

indirectly by ADAM9 (via ADAM10) at a site near the GPI anchor (191).

In addition, PrPC undergoes a physiological endoproteolytic cleavage at the 110/111 or

111/112 peptide bond (termed α-cleavage) (42,181), yielding the C-terminal C1 fragment tethered to the plasma membrane (38,41,42,179,192) and releasing the corresponding N- terminal N1 fragment (46,47,193). The α-cleavage of PrPC is stimulated by protein kinase

C agonists (193) and takes place mostly in a late compartment of the secretory pathway

(194). The α-cleavage disrupts the PrP106-126 region critical for both PrP toxicity and

PrPC to PrPSc conversion. In addition, both products of PrPC α-cleavage are biologically

active: the N1 fragment is neuroprotective in vitro and in vivo by modulating the p53 pathway (189) while the C1 fragment potentiates staurosporine-induced caspase-3 activation in the HEK293 cell line (49).

ADAMs (A Disintegrin And Metalloproteinase) is a family of transmembrane peptidases with a unique multidomain organization, including a prodomain, a proteolytic domain

(metalloprotease) that sheds ectodomains of membrane-anchored cell surface proteins and cleaves extracellular matrix proteins (ECMs), adhesive domains (including a disintegrin domain that binds to integrin and a cysteine-rich domain that binds to heparin sulfate proteoglycans) that interact with ECMs, an EGF-like domain, a transmembrane domain, and a cytoplasmic tail that modulates the sheddase activity (195,196). The

substrates for the ADAM sheddases include Notch, growth factors (such as EGF),

cytokines (such as TNF-α, TRANCE) and their receptors (such as TNF receptors I and II,

NGF receptor, IL-1 receptor and IL-6 receptor), implicating a critical role for ADAMs in

extracellular signalling events (195-197). ADAMs can also cleave adhering molecules

(such as cadherins) and ECMs (such as fibronectin and laminin), thereby promoting cell

migration and releasing ECM-bound growth factors for signaling (195,196).

Three ADAMs have been implicated in the α-cleavage of PrPC. In HEK293 cells,

ADAM10 appears to participate in the constitutive formation of C1 (46,193) while

ADAM17 seems responsible for protein kinase C-dependent formation of C1 (46,198).

ADAM9 was also reported to indirectly participate in C1 formation by modulating

ADAM10 activity in HEK293 cells, mouse fibroblasts and TSM1 neurons (46,47). One

article associates high levels of C1 with the presence of active ADAM10 in the human

brain, but other ADAMs were not examined (48). However, a recent article showed that

overexpression of ADAMs 9, 10, and 17 and depletion of ADAMs 9 and 10 failed to

change the levels of C1 in HEK cell lysates (191), which calls into question the

prominent roles of ADAMs 10, 17, and 9 in the α-cleavage of PrPC and suggests the

involvement of an unidentified alternative protease. In addition, there has been no report

studying the role of ADAMs in the physiological processing of PrPC using animal models.

PrPC is expressed at significant levels (104,105) and implicated in physiological and pathological processes in skeletal muscles.

On one hand, skeletal muscles in PrP-null mice exhibited enhanced oxidative damage

(107) and diminished tolerance for physical exercise (52). In addition, fast muscle fibers,

which during exercise undergo very active oxidative phosphorylation and produce more

reactive oxygen species, present a higher level of PrPC than slow fibers (50). This

evidence suggests a protective role for PrPC. In addition, PrPC is upregulated when

primary or immortalized myoblasts differentiate into myotubes (50,61), and it promotes

regeneration of adult muscle tissues through the stress-activated p38 pathway (37). These

data associate PrPC with muscle differentiation and regeneration.

On the other hand, skeletal muscles showed elevated levels of PrP in patients with sporadic and hereditary inclusion body myositis (55,106), polymyositis, dermatomyositis and neurogenic muscle atrophy (57). In addition, transgenic (Tg) mice constitutively overexpressing wild type PrPs from hamster, sheep or mice developed myopathy in aged animals (35). We also found that induced over-expression of wild type human PrP in the skeletal muscles of Tg(HQK) mice led to a primary myopathy that is correlated with preferential accumulation of C1 (34) and accompanied by activation of the p53- dependent apoptosis pathway (199), suggesting the involvement of C1 and p53 in PrP-

mediated myopathy. However, the detailed pathogenic mechanism of the muscle diseases

induced by over-expressed wild type PrPC is still unclear. What protease(s) performs the

α-cleavage of PrPC and how it is regulated in the skeletal muscles are also unknown.

Here we present in vitro and in vivo evidence to show that, in a myoblast cell line and

skeletal muscle tissues of Tg mouse models, ADAM8 appears to play a major role in the

α-cleavage of PrPC and overexpression of PrPC leads to elevated active ADAM8 levels,

which points to a feedback loop where PrPC regulates its own α-cleavage through

upregulation of ADAM8.

METHODS

Transgenic Mice

The Tg(HQK) mice that show skeletal muscle-specific, doxycycline (Dox)-inducible

expression of human PrPC were described previously (34). The Tg43, Tg4, Tg21 and

Tg17 mice constitutively expressing human PrP were created essentially as described for

the Tg40 mice (200). The Tga20 mice overexpressing mouse PrP at ~8-fold were kindly provided by Charles Weissmann at Scripps Florida. The wild type FVB and C57BL6 mice were from the Jackson Laboratory.

Animal Treatment and Specimen Collection

For Tg(HQK) and wild type FVB (Wt) mice, eight-week-old females were fed food pellets either lacking or containing 6g Dox/kg food (Bio-Serv, Frenchtown, NJ) to induce

PrPC expression, and skeletal muscles from the quadriceps of hind legs were removed

after 0-60 days of Dox treatment. For Tg43, Tg4, Tg21 and Tg17 mice, skeletal muscles

from quadriceps of hind legs of 2-month old female mice were directly removed. The collected muscle tissues were immediately frozen on dry ice and stored at -80oC before

analysis.

Cell Culture and Transfection

Proliferating mouse C2C12 myoblasts were maintained in growth medium (Dulbecco’s modified Eagle’s medium [Invitrogen, Carlsbad, CA], supplemented with 10% fetal bovine serum [Atlanta Biologicals], 100 µg/ml penicillin [Invitrogen, Carlsbad, CA], 100 units/ml streptomycin [Invitrogen, Carlsbad, CA]) in 5% CO2 in a humid incubator at

37°C. To minimize spontaneous differentiation, cells were always kept in subconfluent

(<70%) conditions. Transfection was performed using Effectene (Qiagen, Valencia, CA)

according to the manufacturers’ instructions.

PrP Expression in C2C12 Cells

Mouse PrP (MoPrP) ORF sequence was amplified by PCR from mouse genomic DNA with primers ENS-PRPO-F (GAGAATTCGCGGCCGCGGTCATYATGGCGAACCTT

GG,Y=C+T) and PRPO-Bam-R (CGGGATCCTCATCCCACKATCAGG-AAG, K=T+G)

and cloned into pCEP4 vector (Invitrogen, Carlsbad, CA) to obtain pCEP-MoPrP. C2C12

cells were transfected with pCEP-MoPrP and selected with 300 µg/µl hygromycin B

(Invitrogen, Carlsbad, CA) to obtain several stable cell clones expressing MoPrP.

RNAi Knockdown of ADAM8 in C2C12 Cells

The Invitrogen siRNA design algorithm (BLOCK-IT RNAi Designer) was utilized to

design two 64-mer miR RNAi sequences targeting the ORF region of mouse ADAM8:

NM-1241 (TGCTGTCTCCATGCTCCACAAACAGGGTTTTGGCCACTGACTGACC

CTGTTTGGAGCATGGAGA) and NM-1741 (TGCTGCAATGTTGCTGCCTGTGCCA

AGTTTTGGCCACTGACTGACGTTTGTGGAGCATGGAGAC). These 64-mer

sequences were cloned into the pcDNA 6.2-GW/EmGFP-miR vector (Invitrogen,

Carlsbad, CA) to obtain the ADAM8 miR RNAi constructs, which were then transfected into the MoPrP-expressing C2C12 clones. The transfected cells were selected with

blasticidin S (Invitrogen, Carlsbad CA) and multiple cell clones were obtained. The

ADAM8 protein levels in the RNAi clones were then examined by Western blot (see

below).

Western Blot Analysis

Mouse skeletal muscle tissues were homogenized in a tissue lysis buffer containing 50

mM Tris/HCl (pH 7.5), 200 mM sodium chloride, 0.5% sodium deoxycholate, and 5 mM

EDTA supplemented with a protease inhibitor mixture (Roche, Indianapolis, IN). C2C12

cells grown in 35-mm dishes were washed once with phosphate-buffered saline (PBS)

and lysed with 500 µl of a cell lysis buffer containing 10 mM Tris/HCl (pH 7.5), 150 mM

NaCl, 0.5% Triton X-100, 0.5% deoxycholate and 5 mM EDTA supplemented with a

protease inhibitor mixture (Roche, Indianapolis, IN). Total protein concentrations in the

tissue homogenate and cell lysate were determined by the BCA protein assay (Pierce,

Rockford, IL). For C1 and full-length PrP detection, the samples were first deglycosylated with 10,000 units/ml PNGase F (New England Biolabs, Ipswich, MA) following the manufacturer’s instructions except for incubation at 37 overnight. Then

20 µg of total proteins for each sample were separated by SDS-PAGE℃ on a 10-20%

Criterion Tris-Triton Precast gel (Biorad, Hercules, CA), transferred onto PVDF

membranes for 90 min under 360 mA, incubated with the 8H4 antibody (for human

PrP176-186 and mouse PrP175-185) (201) (1:5000 diluted in 0.5% normal goat serum,

1× TBS and 0.05% Tween) at 4 with gentle shaking overnight, followed by incubation

with sheep anti-mouse IgG (Amersham,℃ Buckinghamshire, UK). For ADAM8 protein

detection, 25 µg of total proteins were separated by SDS-polyacrylamide gel

electrophoresis on a 10% Criterion Tris-HCl Precast gel (Biorad, Hercules, CA),

transferred onto a PVDF membrane for 90 min at 380 mA, incubated with an anti-

ADAM8 C-terminal antibody for active ADAM8 protein (Santa Cruz Biotechnology,

Santa Cruz, CA) (1:1000 diluted in 1% non-fat milk, 1× TBS and 0.05% Tween),

followed by incubation with a donkey anti-rabbit IgG (Amersham, Buckinghamshire,

UK). After stripping the blots with a stripping buffer containing 1.4% 2- mercaptomethanol, 2% SDS and 62.5 mM Tris (pH 6.8), actin was probed with a rabbit polyclonal anti-skeletal muscle actin antibody (Abcam, Cambridge, MA) (1:5000 diluted in 1% milk, 1×TBS and 0.05% Tween) for skeletal muscle samples or with a rabbit polyclonal anti-β-actin antibody (Cell Signaling, Boston, MA) (1:2500 diluted in 1%

milk, 1×TBS and 0.05% Tween) for C2C12 cell samples. The blots were developed with

ECL Western Blotting Detection Reagents (Amersham, Buckinghamshire, UK)

according to the manufacturer’s instructions. X-ray films were exposed to the blots,

developed, scanned and the bands quantified with the UN-SCAN-IT gel 6.1 software

(Silk Scientific, Orem, Utah).

Quantitative Real-Time PCR

Total RNA was isolated from frozen skeletal muscle using the RNeasy skeletal muscle

RNA isolation kit (Qiagen, Valencia, CA) following the manufacturer's specifications.

The total RNA preparations were further treated with Turbo DNA-Free DNase (Ambion,

Austin, TX) to remove residual genomic DNA contamination, and examined with a

Bioanalyzer 2100 (Agilent Technologies, Santa Clara CA) for purity and quantity.

Quantitative real-time PCR of ADAM gene expression was performed using probe

specific TaqMan gene expression assays on the Applied Biosystems 7500 Fast Real-Time

PCR System. 100 ng of total RNA was reverse transcribed using the High Capacity

cDNA Reverse Transcription kit (Applied Biosystems, Foster City, CA). Subsequently, 1

μl from each reverse transcription reaction was assayed in a 20 μl single-plex reaction for

real-time quantification with the 7500 Fast PCR System using probes specific to those

76 genes of interest. Each sample was run in biological triplicate, of which 3 technical replicates were performed. GAPDH was used as the endogenous control, and the expression of target genes in Tg(HQK) mouse samples was quantitatively measured relative to the wild type FVB mouse samples. Relative quantification values were determined using the 2-ΔΔct method, and expressed as fold-change over the wild type FVB samples.

RESULTS

ADAM8 mRNA Level Is Upregulated in Skeletal Muscle of Doxycycline-Treated

Tg(HQK) Mice

To investigate the protease(s) responsible for the α-cleavage of PrPC in skeletal muscles,

Tg(HQK) mice with strictly doxycycline (Dox)-dependent, skeletal muscle-specific expression of human PrP (34) were fed with Dox-laced food for 7, 14, 30, or 60 days; three animals were taken at each time point. Total RNA samples were isolated from skeletal muscle tissues (quadriceps) and subjected to quantitative real time PCR analysis for the three ADAMs already implicated in the α-cleavage of PrPC (ADAM10, ADAM17 and ADAM9) as well as three other ADAMs (ADAM8, ADAM12 and ADAM23). In

Dox-treated wild type FVB mice, no significant changes were found for any of the six

ADAMs; in Dox-treated Tg(HQK) mice, ADAM8 mRNA level was significantly upregulated whereas the other five ADAMs were largely unchanged (Figure 4.1). This data indicate that, instead of the three previously implicated ADAMs (ADAM10,

ADAM9 and ADAM17), ADAM8 may be the candidate protease for C1 production in skeletal muscles.

Active ADAM8 Protein Level Correlates With PrP C1 Production in the Skeletal

Muscles of Inducible and Constitutive Transgenic Mice

Another time course experiment was performed to examine the relationship between active ADAM8 protein level and PrP C1/full length ratio, the latter serves as a marker for

C1 production. Young female adult Tg(HQK) mice were fed with Dox-laced food for 0, 2,

Figure 4.1

Quantitative real-time PCR analysis of ADAMs gene expression. Hind leg skeletal muscles (quadriceps) were taken from Tg(HQK) mice treated with 6 g Dox/kg food for 7, 14, 30, and 60 days. Three animals were used for each time point. Total RNA was extracted and subjected to quantitative real-time PCR analysis for ADAM8, ADAM9, ADAM10, ADAM12, ADAM17 and ADAM23 gene expression. Wild type FVB mice were similarly treated and analyzed. No significant changes were observed in the wild type FVB mice over the time course. Results are relative to the average expression of the corresponding ADAM genes in the skeletal muscles of similarly treated wild-type FVB control mice and represent the mean ± SEM of triplicate measurements performed.

4, 7, 14, 30, and 60 days; 3 animals were sacrificed at each time point and hind leg

(quadriceps) muscles were collected and homogenized. The muscle homogenates were

either treated with PNGase F to remove N-linked glycans and immunoblotted with 8H4

(a monoclonal antibody against the human PrP 176-186 epitope) or were directly subjected to Western blot with a rabbit polyclonal antibody that detects active ADAM8 protein.

Figure 4.2

ADAM8 protein level correlates with PrP C1/full-length ratio in skeletal muscle tissue of Dox- induced Tg(HQK) mice. Tg(HQK) mice were treated with 6 g Dox/kg food for 0-60 days as indicated and three animals were taken at each time point. Skeletal muscles (quadriceps) from the hind legs of these animals were homogenized and subjected to SDS-PAGE and immunoblot analysis in three Western blots. The homogenates were either treated (for PrP probing) or not treated (for ADAM8 probing) with PNGase F before SDS-PAGE. (A) A representative immunoblot probed with anti-PrP antibody 8H4 followed by probing with an anti-actin antibody after stripping. (B) A representative immunoblot probed with an antibody against active ADAM8 followed by probing with an anti-actin antibody after stripping. (C) & (D) Diagrams of PrP C1/full-length ratios (C) and active ADAM8 protein level (D) over increasing duration of Dox treatment. The active ADAM8 protein level for each sample was normalized against the actin level for the sample in each blot and expressed as the ratio against the normalized ADAM8 protein level in the untreated wild type FVB mouse on the same blot. The error bars denote standard deviations calculated from the three blots. The bars with asterisk(s) indicate a statistically significant difference when compared to the day 0 Tg(HQK) samples for ADAM8 or to the Wt FVB samples for C1/full-length ratio; * p<0.05, ** p<0.01. (E) Plot of PrP C1/full length ratio against active ADAM8 protein level over the time course of Dox treatment.

The PrP C1/full length ratio in the skeletal muscle rose dramatically from day 4 to day 7

and stayed at a plateau of ~3.0 from day 7 of Dox treatment (Figure 4.2A & 4.2C), confirming our previous report (34). The active ADAM8 protein level in the skeletal muscle showed statistically significant increase from day 4 and kept rising with time

(Figure 4.2B & 4.2D). Plotting the PrP C1/full-length ratio against active ADAM8 protein level reveals that, before the active ADAM8 level reached 1.5, there was an apparently linear correlation between active ADAM8 level and the PrP C1/full-length ratio; but after active ADAM8 level reached 1.5 at day 7, the PrP C1/full-length ratio stayed at ~3.0 despite further increase of the active ADAM8 level from day 14 to day 60

(Figure 4.2E).

In Tg(HQK) mice treated with Dox, both PrP and ADAM8 expressions were in a dynamic state (rising with time) and the PrP levels were extremely high from day 14, which complicate data interpretation. To address this issue, the relationship between active ADAM8 protein level and C1 production in the skeletal muscles was assessed in 4 transgenic (Tg) mouse lines, Tg43, Tg4, Tg17 and Tg21, which constitutively express human PrP at different levels in the skeletal muscles (Figure 4.3A). Skeletal muscles

(quadriceps) from 3 young female adult animals for each transgenic line were examined by Western blot for active ADAM8 level and PrP C1/full length ratio as described above for Tg(HQK) mice; skeletal muscle from a wild type FVB mouse was used as control. Tg lines with higher active ADAM8 protein levels in skeletal muscles also showed higher

PrP C1/full length ratios in skeletal muscles, and the two exhibited an excellent linear correlation with a R2 value of 0.93 (Figure 4.3).

Figure 4.3

ADAM8 protein level correlates with PrP C1/full-length ratio in skeletal muscle tissue of Tg mice constitutively expressing human PrP at different levels. Skeletal muscles (quadriceps) from the hind legs of wild type FVB mice (Wt) and four different Tg mice lines (Tg43, Tg4, Tg17 and Tg21) constitutively expressing human PrP at different levels were collected. Muscle homogenates were either treated (for PrP probing) or not treated (for ADAM8 probing) with PNGase F, subjected to SDS-PAGE, and probed with an anti-PrP antibody (8H4) or an antibody against active ADAM8. (A) A representative immunoblot probed with anti-PrP antibody 8H4 followed by probing with an anti-actin antibody after stripping. (B) A representative immunoblot probed with an antibody against active ADAM8 followed by probing with an anti-actin antibody after stripping. (C) & (D) Diagrams of active ADAM8 protein level (C) and PrP C1/full-length ratios (D) in the skeletal muscles of the four Tg mouse lines. The active ADAM8 protein level for each sample was normalized against the actin level and expressed as the ratio against the normalized ADAM8 protein level in the untreated sex and age matched wild type FVB mouse on the same blot. The error bars denote standard errors calculated from the three animals for each Tg line. The bars with asterisk(s) indicate statistical significance when compared to the Tg43 mouse samples; * p<0.05, ** p<0.01. (E) Plot of PrP C1/full length ratios against active ADAM8 protein levels in the four Tg lines. Linear regression analysis revealed an R2 value of 0.93, highly indicative of a linear correlation between active ADAM8 protein level and the PrP C1/full length ratio.

Figure 4.4

ADAM8 protein level correlates with PrP C1/full-length ratio in skeletal muscle tissue of mice constitutively expressing mouse PrP at different levels. Skeletal muscles (quadriceps) from the hind legs of wild type C57BL6 mice (BL6) and the Tga20 line constitutively expressing mouse PrP at high levels were collected. Muscle homogenates were either treated (for PrP probing) or not treated (for ADAM8 probing) with PNGase F, and subjected to SDS-PAGE and immunoblotting with an anti-PrP antibody (8H4) or an antibody against active ADAM8. (A) A representative immunoblot probed with anti-PrP antibody 8H4 followed by probing with an anti-actin antibody after stripping. (B) A representative immunoblot probed with an antibody against active ADAM8 followed by probing with an anti-actin antibody after stripping. (C) Plot of PrP C1/full length ratios against relative active ADAM8 protein levels. The error bars denote standard errors calculated from the three animals for each mouse line. The bars with an asterisk indicate statistical significance (p<0.05) when compared to the wild type C57BL6 (BL6) samples.

Active ADAM8 levels and PrP C1/full length ratios were also examined in the skeletal muscles of wild type C57BL6 mice and Tga20 mice that overexpress mouse PrP. When compared with those of C57BL6 mice, the active ADAM8 protein level and PrP C1/full length ratio in the Tga20 mice were both significantly higher (2.3-fold and 1.8-fold, respectively) (Figure 4.4), confirming that active ADAM8 protein level is positively correlated with C1/full length ratio irrespective of the species origin of the PrP.

Figure 4.5

Over-expression of mouse PrP leads to up-regulation of ADAM8 in C2C12 cells. C2C12 cells were transfected with a vector expressing mouse PrP and selected with hygromycin to establish seven stable cell clones (numbered 1-7). The original C2C12 cells (CO) and blank vector-transfected C2C12 cells (VC) serve as controls. Lysates from these clones were directly subjected to Western blotting and the experiments repeated three times. (A) A representative immunoblot probed with 8H4 followed by probing with an anti-actin antibody after stripping. (B) A representative immunoblot probed with an antibody against active ADAM8 followed by probing with an anti-actin antibody after stripping. (C) Diagram of total PrP levels and active ADAM8 levels in the C2C12 clones. The total PrP and active ADAM8 protein levels for each sample were normalized against the actin level and expressed as the ratio against the normalized protein levels in the #1 clone. The error bars denote standard deviations calculated from the three blots. The bars with asterisk(s) indicate statistical significance when compared to the #1 clone; * p<0.05; ** p<0.01.

Active ADAM8 Protein Level Shows Linear Correlation with PrP C1/Full Length

Ratio in the Myoblast Cell Line C2C12

C2C12, a murine myoblast cell line, was utilized to examine the role of ADAM8 in C1 production in pure muscle cells. C2C12 cells normally express very little PrP, but PrP

Figure 4.6.

The PrP C1/full length ratio is proportional to active ADAM8 protein level in C2C12 myoblast cells. Two stable C2C12 clones expressing mouse PrP (MoPrP-6 and MoPrP-7) were transfected with miR RNAi sequence targeting the ORF region of ADAM8 and several stable cell lines were obtained. Lysates from these stable cell lines were either treated (for PrP probing) or not treated (for ADAM8 probing) with PNGase F, then subjected to electrophoresis and immunoblotting with an anti-PrP antibody (8H4) or an antibody against active ADAM8. The immunoblotting was repeated three times. (A) Representative immunoblots probed with anti-PrP antibody 8H4 or with an antibody against active ADAM8 followed by probing with an anti-actin antibody after stripping. (B) Plot of PrP C1/full-length ratios against active ADAM8 protein levels in ADAM8-knockdown C2C12 cell lines. The active ADAM8 protein level for each C2C12 cell line was normalized against the actin level and expressed as the ratio against the normalized active ADAM8 protein level in the original MoPrP-6 control cell line (CO) on the same blot. Data from the average of three duplicate blots are plotted. CO, the original MoPrP6/MoPrP7 cell line; VC, blank vector-transfected MoPrP6/MoPrP7 cell line; KD, knockdown. expression is activated during myoblast differentiation (37,50). To study the α-cleavage of PrPC in the C2C12 cell line, C2C12 cells were first transfected with pCEP4 carrying full-length mouse PrP (MoPrP) ORF and many C2C12 clones expressing MoPrP at various levels were established (Figure 4.5A). Two of such C2C12 clones (MoPrP-6 and

Figure 4.7

Over-expression of human PrPC leads to up-regulation of ADAM8 in transgenic mice. Western blots for PrP and ADAM8 in homogenates of skeletal muscles from Dox-treated Tg(HQK) mice (shown in FIGURE 1A-B), Tg lines constitutively expressing human PrP at various levels (shown in FIGURE 2A-B), and C57BL6 (BL6) and Tga20 mice (shown in FIGURE 4A-B) were quantified after scanning the X-ray films. The active ADAM8 protein level and total PrP protein level for each sample were normalized against the actin level and expressed as the ratio against the corresponding normalized protein level in the untreated sex and age matched wild type FVB mouse (for panels A & B) or wild type C57BL6 mice (for panel C) on the same blot. (A) Diagram of total PrP levels and active ADAM8 levels in Tg(HQK) mice over the time course of Dox treatment. The statistical difference was calculated based on comparison with day 0 Tg(HQK) samples for ADAM8 or with day 2 Tg(HQK) samples for total PrP. (B) Diagram of the total PrP levels and active ADAM8 levels in the four constitutive Tg mouse lines. The statistical difference was calculated based on comparison with Tg43 samples. (C) Diagram of the total PrP levels and active ADAM8 levels in C57BL6 and Tga20 mice. The statistical difference was calculated based on comparison with C57BL6 (BL6) samples. For all panels, the error bars denote standard deviations calculated from the three animals for each Tg line; *P<0.05, ** p<0.01.

MoPrP-7) that express comparable levels of MoPrP were selected for ADAM8

knockdown (KD) with miR RNAi sequences cloned in the pcDNA vector. Three stable

ADAM8-KD cell lines derived from MoPrP-6 and four stable ADAM8-KD cell lines

derived from MoPrP-7 were obtained. Cell lysates from these stable ADAM8-KD cell lines were immunoblotted for PrP or active ADAM8, and the PrP C1/full length ratios

were plotted against the levels of active ADAM protein in these cell lines (Figure 4.6).

The result shows that decreasing active ADAM8 protein level in the ADAM8-KD cells is accompanied by a proportional drop in the PrP C1/full length ratio; linear regression analysis reveals an excellent linear correlation between the active ADAM8 level and the

C1/full-length ratio with an R2 value of 0.94 (Figure 4.6).

Together with the data from the Dox-treated Tg(HQK) mice and the constitutive Tg mouse lines, this result strongly argues for a primary role for ADAM8 in the α-cleavage of PrPC for C1 production in skeletal muscle cells.

ADAM8 is upregulated by PrP

Further examination of Western blots of total PrP and active ADAM8 levels in the skeletal muscles of Dox-treated Tg(HQK) mice (Figure 4.2) revealed that, starting from

day 4 when total PrP level reached 4 times that of wild type mice, the active ADAM8

protein level rose significantly (Figure 4.7A). Quantification of Western blots for the four

Tg lines constitutively expressing human PrP (Figure 4.3) and the Tga20 mice overexpressing mouse PrP (Figure 4.4) also indicates that PrP over-expression resulted in elevated active ADAM8 protein levels (Figure 4.7B-C). Similarly, when compared with the C2C12 clone #1, the four C2C12 clones (#4, #5, #6 and #7) that expressed MoPrP

87 at >4-fold levels showed >2 fold active ADAM8 protein levels as well (Figure 4.5).

Together these in vivo and in vitro data demonstrate that overexpression of PrP leads to elevated ADAM8 activities in muscle cells. Since ADAM8 mRNA was also upregulated in the muscles of Dox-treated Tg(HQK) mice (Figure 4.1), PrP overexpression-induced upregulation of ADAM8 is likely to be at the transcriptional level.

DISCUSSION

The α-cleavage of PrPC disrupts the protein in a region critical for both toxicity and prion

conversion and generates the biologically active C1 and N1 fragments. Our current

understanding of the α-cleavage of PrPC is derived almost exclusively from in vitro

studies using cultured HEK and mouse fibroblasts or neuronal cells. However, the earlier

reports implicating the direct involvement of ADAM10 and ADAM17 and an indirect

role for ADAM9 (46,47,198) were challenged by a recent article (191), which leaves a

question mark on the identity of the protease(s) primarily responsible for the α-cleavage of PrPC.

We have investigated the protease likely responsible for the α-cleavage of PrPC in

skeletal muscles using Tg mouse models and the C2C12 myoblast cell line. We found

that in the skeletal muscles of Dox-treated Tg(HQK) mice, among 6 ADAMs examined

by quantitative real-time PCR, only the ADAM8 mRNA was significantly upregulated

(Figure 4.1). Western blot analysis confirmed a significant increase of active ADAM8

protein level starting from day 4 of Dox treatment, which preceded the dramatic

preferential accumulation of C1 that began at day 7 of Dox treatment (Figure 4.2).

Further examinations revealed that the active ADAM8 protein level is linearly correlated

with the PrP C1/full-length ratio in the skeletal muscles of not only the Dox-treated

Tg(HQK) mice (up to day 7) but also the Tg mouse lines constitutively expressing human

PrP (Figure 4.3). When compared with the wild type C57BL6 mice, a higher level of

active ADAM8 was also associated with an increased PrP C1/full-length ratio in the

Tga20 mice expressing mouse PrP (Figure 4.4), indicating that the positive correlation

between active ADAM8 level and C1/full-length ratio is not specific to human PrP.

Moreover, a linear correlation between active ADAM8 protein level and PrP C1/full-

length ratio was also observed in C2C12 myoblast clones expressing the same level of

mouse PrP but with different levels of active ADAM8 due to RNAi knockdown (Figure

4.6). Together, these in vivo and in vitro results strongly argue for a primary role for

ADAM8 in C1 production in skeletal muscles. ADAM10, the proposed protease for C1

production in the brain (48), was not upregulated in the skeletal muscles of Dox-treated

Tg(HQK) mice (Figure 4.1). We found that the ADAM10 protein level in skeletal muscle

is only about 2% of the brain level in wild type FVB mice (data not shown). These data

indicate that ADAM10 is unlikely to play a major role in C1 production in the skeletal

muscles. Nevertheless, our data cannot exclude potential involvement of other ADAMs

or other proteases in the α-cleavage of PrPC for C1 production in the skeletal muscles. It

remains to be investigated whether ADAM8 also participates in the α-cleavage of PrPC in

brain and other tissues.

We also discovered that over-expression of wild type PrP upregulates ADAM8 in the

skeletal muscles of Dox-treated Tg(HQK) mice (Figure 4.7A) and Tg lines constitutively expressing different levels of human or mouse PrP (Figure 4.7B-C), and in C2C12 cells expressing different levels of mouse PrP (Figure 4.5). Given that the ADAM8 mRNA level was also significantly increased in the skeletal muscles of Dox-treated Tg(HQK) mice (Figure 4.1), it is very likely that PrPC regulates ADAM8 at the transcriptional level.

The molecular mechanism underlying the over-expressed PrP-induced ADAM8 expression is unclear. PrPC was recently reported to promote regeneration of adult muscle

tissue through affecting the generation of TNF-α (37,202), a cytokine actively involved in

myogenesis and muscle repair (203-207). In addition, the expression of ADAM8 was

induced by TNF-α in primary cerebellar neurons and in motoneuron-like NSC19 cells in

a dose dependent manner (97). Moreover, TNF-α regulates the shedding of TNF-α

receptor 1 through elevating ADAM8 activity in the mouse brain and primary neurons

and microglia (208). Furthermore, antibody-mediated PrPC ligation was reported to cause

TNF-α shedding in cultured neurons (209). It is conceivable that, in the muscles, the over-expressed PrPC somehow activates TNF-α to induce ADAM8 transcription, and the

increased ADAM8 activity results in augmented α-cleavage of PrPC.

It is not clear why the PrP C1/full length ratio remained unchanged from day 7 to day 60

despite the continued rise in the active ADAM8 level in the skeletal muscle of Dox-

treated Tg(HQK) mice (Figure 4.2). The very high levels of PrP at these later time points

could counter the increase of ADAM8 activity to maintain a stable PrP C1/full-length

ratio. Alternatively, an unknown inhibitor(s) of ADAM8 may be activated at very high

PrP levels.

Our findings that ADAM8 can perform α-cleavage of PrPC and that PrPC can regulate

ADAM8 expression have significant implications. It points to a self-regulatory loop where overexpressed PrPC modulates its own α-cleavage through upregulating ADAM8, leading to accumulation of cytotoxic C1 when PrPC is overexpressed. This self-regulatory

loop may not only explain the myopathy incurred by over-expressed wild type PrPC in

aged Tg mice (35) and Dox-induced Tg(HQK) mice (34,199), but also provide a plausible mechanism for PrP toxicity observed in human HEK293 cells, mouse neurons and erythroleukemia cells (81,83,156,210,211). When PrP is over-expressed for an

91 extended period of time, the upregulation of ADAM8 will lead to accumulation of the cytotoxic and more stable C1 fragment (192) at a higher molar ratio than full-length PrPC.

The concurrently released neuroprotective N1 fragment has a shorter half-life (189) and may provide only transient and possibly insufficient protection from the toxicity of the accumulated C1. In addition, the elevated ADAM8 activity may induce further effects stemming from the enhanced cleavage of its many other substrates (196), some of which might also modulate the cytotoxic effect of overexpressed PrP.

CHAPTER 5

Conclusions and Future Directions

Conclusions

We performed DNA microarrays and confirmatory real-time PCR analysis to

demonstrate deregulation of a large number of genes in the course of the progressive

myopathy in skeletal muscles of Dox-treated Tg(HQK) mice. This includes up-

regulation of genes for p53-related pathways involved in cell cycle arrest and promotion

of apoptosis as well as a member of the ADAMs protein family, ADAM8, that paralleled

the initiation and progression of the muscle pathology. Treatment of Dox-induced

Tg(HQK) mice with PFT-α, a p53 specific inhibitor, led to a significant attenuation of

PrP-mediated myopathy, confirming a critical role for p53-regulated pathways.

By employing murine C2C12 myoblast cell line and several transgenic mice lines expressing human or murine PrP at various levels, we found that over-expression of PrPC

leads to up-regulation of ADAM8, and active ADAM8 protein level linearly correlates

with PrP C1/full-length ratio both in vitro and in vivo. These findings demonstrate that

PrPC regulate ADAM8 expression and ADAM8 is primarily responsible for physiological

processing of PrPC to generate the C1 fragment in skeletal muscle.

We propose the following working model to depict the PrP-ADAM8 relationship and the

mechanism of PrP-mediated myopathy (Figure 5.1). In Tg(HQK) mice, Dox-induced

over-expression of PrPC in the muscles leads to up-regulation of ADAM8, which in turn increases the production of PrP C1 fragment. Accumulation of PrP C1 fragment activates p53, thereby activating p53-regulated pro-apoptotic networks and resulting in myopathic changes. Additionally, PrPC over-expression also results in down-regulation of MEF2c,

which may be partially responsible for the progressive central nucleus localization.

Figure 5.1 A Model for PrP-mediated Myopathy.

Further Directions

The following experiments are worth pursuing in the future.

1. Is C1 responsible for p53 activation and ADAM8 required for PrP-mediated myopathy?

We have shown statistically highly significant induction of a p53-regulated pro-apoptotic network and p53 protein in the skeletal muscles of Tg(HQK) mice following induction of

PrPC. We have also found that blocking of the activity of p53 pathways significantly attenuated the progression of PrPC-mediated myopathy. However, what activates the p53 pathways remains uncertain.

We propose that the C1 fragment is the culprit. To evaluate this hypothesis, we will

generate Dox-inducible transgenic mouse line with muscle specific expression of a

mutant PrP that cannot be cleaved by ADAMs, preventing C1 production. We will

examine whether induced over-expression of full-length mutant PrP can still induce activation of p53 pathways and myopathy. We will also breed Tg(HQK) mice with the

ADAM8-KO mice (see below) and demonstrate that, in the absence of ADAM8, no C1 will be produced and high levels of full-length PrPC will not cause myopathy.

2. Further prove that ADAM8 is required for C1 production in skeletal muscles

We demonstrated that knockdown of ADAM8 in C2C12 cell line led to decreased C1

production in vitro and high levels of ADAM8 protein in skeletal muscles of transgenic

mice were correlated with higher PrP C1/full-length ratios in vivo. To prove beyond any

doubt that ADAM8 is directly and primarily responsible for C1 production in the skeletal

muscles, we will use recombinant ADAM8 to show that it can generate C1 from

recombinant PrP substrates or from PrP on cell surface. We will also use the ADAM8-

KO mouse model (10), which has been obtained from Dr. Carl L. Blobel at Weill

Medical College of Cornell University, to show that there is minimal or no C1 production

in the skeletal muscles of the ADAM8-KO mice.

3. Further dissect the signaling pathways in PrP-mediated myopathy

PrPC is implicated in the morphogenesis of the skeletal muscle to promote regeneration of

muscle tissue in adult mice by modulating signaling pathways central to the myogenic

process, in particular the p38 pathway (37,202). PrPC is also shown to regulate TNF-α

activity through TACE in skeletal muscle of adult mice (37,202), while TNF-α could

96 induce ADAM8 expression in primary cerebellar neurons and in motor neuron-like

NSC19 cells in a dose dependent manner (97). Recently, a study further dissected the neuroprotective feedback loop involving TNF-α, TNF-R1 and ADAM8, and showed that high levels of TNF-α induce TNF-R1 to activate intracellular pathways to stimulate transcription of ADAM8 gene (208). These data suggest that TNF-α may also be involved in PrP self-regulation of its own cleavage through ADAM8. We will treat PrP- expressing C2C12 cells with small molecular TNF-α inhibitors to see whether ADAM8 protein levels and C1 production are affected.

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