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Respiratory and Neural Characterization of a Mouse Model of Pompe Disease: Insights into Gene Therapy Mediated Treatment

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Respiratory and Neural Characterization of a Mouse Model of Pompe Disease: Insights into Gene Therapy Mediated Treatment
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DERUISSEAU, LARA ROBERTS ( Author, Primary )
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2008

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Subjects / Keywords:
Diaphragm ( jstor )
Diseases ( jstor )
Gene therapy ( jstor )
Generally accepted auditing standards ( jstor )
Glycogen ( jstor )
Glycogen storage disease type II ( jstor )
Spinal cord ( jstor )
Tidal volume ( jstor )
Trucks ( jstor )
Ventilation systems ( jstor )

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University of Florida
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University of Florida
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Copyright Lara Roberts Deruisseau. Permission granted to the University of Florida to digitize, archive and distribute this item for non-profit research and educational purposes. Any reuse of this item in excess of fair use or other copyright exemptions requires permission of the copyright holder.
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8/31/2007

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RESPIRATORY AND NEURAL CHARACTE RIZATION OF A MOUSE MODEL OF POMPE DISEASE: INSIGHTS INTO GENE THERAPY MEDIATED TREATMENT By LARA ROBERTS DERUISSEAU A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLOR IDA IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY UNIVERSITY OF FLORIDA 2006

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Copyright 2006 by Lara Roberts DeRuisseau

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This dissertation is dedicated to my parents, Mark and Patricia Roberts.

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iv ACKNOWLEDGMENTS There are many colleagues, friends and family that I would like to thank for their support throughout my graduate training. Firs t, I would like to thank my mentor, Dr. Barry Byrne for the opportunity to work in his lab. His enthusiasm for both basic science and patient care has offered me a bette r appreciation for the long-term goal of experiments. The opportunities I have b een afforded through his mentorship will continue to aid my scientif ic career. Through his mentor ship I have met patients and their families, whose courage and commitment to finding a cure for Pompe disease have inspired me. I would also like to thank a ll the members of the Byrne lab including Dr. Cathryn Mah, Dr. Kerry Cresawn, Dr. Bijoy Thattaliyath, Laura Mi riel, Mimi Zarate, Denise Cloutier, Christina Pacak, Jeffrey Ke lly, Dr.Gregory Simon, Dr.Yoshihisa Sakai, Melissa Lewis, Stacy Porvasnik, Sean Germain and Sophia Wa ng for their technical support, scientific discussions and daily encouragement. I greatly appreciate the guid ance, patience, and insight of Dr. David Fuller. Without his expertise, this project would not have been possible. Nicholas Doperalski and Sandy Morrison in his lab have given of their time unconditionally and always treated me like a regular member of the Fuller lab. Dr. Fuller’s dedication to this project has been unwavering. The expertise of Dr. Paul Reier has been indispensable throughout my time at the University of Florida. His list of acc omplishments, accolades and commitments is extensive, yet he always made time for scie ntific discussions a nd career advice. Dr.

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v Reier’s support throughout this process has given me confidence in my own skills and decisions. This dissertation in Dr. Byrn e’s lab would not be possible without the dedicated effort of Dr. Roger Reep, graduate advisor for the Department of Physiological Sciences. Dr. Reep’s guidance throughout my graduate training has made it possible for me to study under the direction of Dr. Byrne. Without Dr. Reep I would not have met Drs. Byrne, Fuller and Reier who have been in strumental in my trai ning. In addition to Dr. Reep’s leadership throughout my time as a graduate student at UF, he has made me think outside of the box for this entire thesis. Finally, I would like to thank my famil y. My parents have offered endless encouragement and support throughout my unde rgraduate and graduate training. Their unconditional love has given me a life that is overflowing. I would like to thank my brother, Brad, who has always had confidence in me as a scientist, but more importantly has always been my friend. These acknowle dgements would not be complete without thanking my husband, Dr. Keith DeRuisseau. Keith’s love, emotional support and continuous faith in my abilities as a scientist have made this journey possible.

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vi TABLE OF CONTENTS page ACKNOWLEDGMENTS.................................................................................................iv LIST OF TABLES.............................................................................................................ix LIST OF FIGURES.............................................................................................................x ABBREVIATIONS..........................................................................................................xii ABSTRACT.....................................................................................................................xi ii CHAPTER 1 BACKGROUND..........................................................................................................1 History of Pompe Disease............................................................................................1 Glycogen Storage and Breakdown...............................................................................1 Biochemistry of Acid -glucosidase............................................................................2 Gene Expression of Acid -glucosidase......................................................................3 Pompe Disease..............................................................................................................4 Target Organs...............................................................................................................6 Skeletal and Cardiac Muscle.................................................................................6 Brain and Spinal Cord...........................................................................................6 Animal Models.............................................................................................................8 Treatments for Pompe Disease...................................................................................10 Enzyme Replacement Therapy............................................................................10 Gene Therapy......................................................................................................12 Summary.....................................................................................................................18 2 RESPIRATORY AND NEURAL C HARACTERIZATION OF A MOUSE MODEL OF POMPE DISEASE................................................................................19 Introduction.................................................................................................................19 Methods......................................................................................................................21 Animals................................................................................................................21 Barometric Plethysmography..............................................................................21 Hemoglobin, Hematocrit, Glucose and Sodium Blood Levels...........................23 Conscious Arterial Blood Sampling....................................................................23 Glycogen Quantification.....................................................................................24

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vii Retrograde Labeling of Phrenic Motoneurons....................................................25 Histological Glycogen Dete ction in Motoneurons..............................................26 In Vitro Diaphragmatic Contractility..................................................................26 Efferent Phrenic Nerve Recordings.....................................................................27 Statistics...............................................................................................................29 Results........................................................................................................................ .31 General Features of Gaa-/Mice..........................................................................31 Glycogen Quantification and PAS Staini ng of the Cervical Spinal Cord...........31 Ventilation...........................................................................................................36 Hemoglobin and Hematocrit...............................................................................39 Conscious Arterial Blood Sampling....................................................................39 Muscle-Specific hGAA Mice..............................................................................40 Efferent Phrenic Activity.....................................................................................43 Discussion...................................................................................................................43 The Gaa-/Mouse Model and Altered Ventilation...............................................44 Evidence for a Neural Contribution to Respiratory Deficits in Gaa-/Mice........46 Contribution of Diaphragm Musc le in Ventilation Deficits................................48 Therapeutic Implications.....................................................................................49 3 CHARACTERIZATION OF THE IN TRATHORACIC INJECTION USING ADENO-ASSOCIATED VIRUS SEROTYPE 1.......................................................51 Background.................................................................................................................51 Materials and Methods:..............................................................................................52 Animals................................................................................................................52 Virus Preparation.................................................................................................52 Survival Surgery for Intrapleural Injection.........................................................52 Tissue Harvesting................................................................................................53 Immunostaining for LacZ....................................................................................53 DNA Isolation.....................................................................................................54 Polymerase Chain Reaction (PCR).....................................................................54 LacZ Protein Activity..........................................................................................55 Fluorescent Imaging of Mouse Diaphragm.........................................................55 Fluorescent Labeling of Motoneurons.................................................................56 Co-labeling of AAV with Motoneurons..............................................................56 Statistics...............................................................................................................57 Results........................................................................................................................ .57 Intrathoracic Injection: Dilution Media...............................................................57 Intrathoracic Injection: Targeting of Mouse Diaphragm.....................................57 DNA Detection of Control Gene.........................................................................61 Protein Activity of Control Gene........................................................................62 Labeling of Motoneurons....................................................................................62 Discussion...................................................................................................................66 4 METHOD OF BAROMETRIC PLETHYSMOGRAPHY........................................68 What is the Principle of Barometric Plethysmography?.............................................68

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viii Thermal and Pressure Drifts................................................................................71 Inaccurate Measurements of Body or Chamber Temperature.............................71 The Assumption that Warmed, Saturated Air from the Airways Reaches Equilibrium Immediately Once It Returns to the Chamber.............................72 The Assumption That When Flow Is Zero During the Breath, That This Is the End of Inspiration............................................................................................72 Alterations in Frequency and Airway Resistance Can Alter the Tidal Volume Calculation.......................................................................................................73 5 CONCLUSIONS AND FUTURE DIRECTIONS.....................................................74 LIST OF REFERENCES...................................................................................................76 BIOGRAPHICAL SKETCH.............................................................................................89

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ix LIST OF TABLES Table page 2-1 Body weight values for control and Gaa/mice at 6, 12 and >21 months of age. *= Gaa-/different from control values for all age groups. =male different from female for all age groups. No age effect detected...................................................31 2-2 Baseline ventilation characteristics . *=p<0.01: different from control. =p<0.01: >21 months diffe rent from 6 months......................................................38 2-3 Six month mean response to hyperca pnia. Ten minute mean response to hypercapnia (7% CO2, balanced O2). * = Gaa-/different from control. = male different from female for each variable presented...........................................38 2-4 Body weight as a possible cova riate to respiratory volumes......................................39 2-5 Hemoglobin, hematocrit, sodium and glucose levels for control and Gaa-/mice at 12 months. *=p<0.01: different from control......................................................39 2-6 Conscious arterial blood sampling to meas ure arterial partial pressure of O2 in control and Gaa-/mice at 12 months. *=p<0.01 : different from control...............40 2-7 Bodyweights of muscle specific Gaa mi ce used in ventilation experiments...........40 2-8 Phrenic neurophysiology characteristics for control, Gaa-/and MTP mice. *=p<0.01, different from control..............................................................................42

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x LIST OF FIGURES Figure page 2-1 Glycogen quantification for unspecified spinal segments. *=Gaa-/different from control, #=24 month different fr om 6 month, glycogen measured by g glycogen/mg wet weight. n=4/group........................................................................32 2-2. Glycogen quantification for cervical spinal segments C3-C5. *=p<0.01: different from control. N=6/group..........................................................................................33 2-3 Histological staining for glyc ogen in phrenic motoneurons....................................34 2-4 Minute Ventilation : Expired CO2 ratio for control and Gaa-/mice at 6 (A), 12 (B) and >21 (C) months of age. *= Gaa-/different from control............................35 2-5 Frequency (5A, 5D, 5G), tidal volume (5B, 5E, 5H) and minute ventilation (5C, 5F, 5I) in 6 (left panel), 12 (middle pa nel) and >21 (right panel) month control and Gaa-/mice at baseline and 10 mi nutes of hypercapnia. ..................................37 2-6 Diaphragm contractile function (n=3/g roup) and minute ven tilation for control (n=8), Gaa-/(n=8) and MTP (n=6) mice. Airf low tracings from representative control, Gaa-/ and MTP mice................................................................................42 2-7 Thirty second mean phrenic inspiratory burst amplitude for control, Gaa-/and muscle specific hGaa mice with similar arterial PaCO2 values (2-8-A)..................44 3-1 Media comparision for diaphragm -Galactosidase activity from mice injected with 1 x 1011 particles of AAV1-CMV-LacZ..........................................................58 3-2 Media comparision for spinal cord -Galactosidase activit y from mice injected with 1 x 1011 particles of AAV1-CMV-LacZ..........................................................59 3-3 Intrathoracic injection with 100L of IR Dye 800 (Li-Cor Biosciences; images green)........................................................................................................................6 0 3-4 X-Gal staining of mouse diaphragm following intrathoracic injection with AAV1-CMV-LacZ (left panel) or phospha te buffered saline (right panel).............60 3-5 PCR product run on a 1.5 % agarose gel from diaphragm DNA amplified with primers specific for the LacZ gene...........................................................................61

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xi 3-6 PCR product run on a 1.5 % agarose gel from isolated spinal cord (C3-C5) DNA amplified with primers specific for LacZ.................................................................62 3-7 -galactosidase activit y of diaphragm muscle from mice injected with AAV1CMV-LacZ or phosphate buffered saline.................................................................63 3-8 -galactosidase activity of spinal cord (C3-C5) from mice injected with AAV1CMV-LacZ or phosphate buffered saline.................................................................64 3-9 Fluoro-gold identified motoneurons.........................................................................65 3-10 FITC filter used for identification of GFP positive cells. No cells detected...........66 4-1 Theoretical basis for plethysmography....................................................................69 4-2 Thermal drift............................................................................................................71

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xii ABBREVIATIONS AAV Adeno-associated virus B-gal Beta-galactosidase BSA Bovine serum albumin GAA Acid alpha-glucosidase protein GAA Acid alpha-glucosidase gene Gaa-/Acid alpha-glucosidase knockout mouse model hGAA human GAA protein hGAA human GAA gene HRP Horse radish peroxidase ITR Inverted terminal repeat M6PR Mannose 6-phosphate receptor min minute PAS Periodic Acid Schiff PBS Phosphate Buffered Solution rAAV Recombinant adeno-associated virus rhGAA Recombinant human GAA protein TRE Tetracycline response element wk week/s

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xiii Abstract of Dissertation Pres ented to the Graduate School of the University of Florida in Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy RESPIRATORY AND NEURAL CHARACTE RIZATION OF A MOUSE MODEL OF POMPE DISEASE: INSIGHTS INTO GENE THERAPY MEDIATED TREATMENT By Lara Roberts DeRuisseau August 2006 Chair: Barry J. Byrne Major Department: Veterinary Medicine Pompe disease is an inborn metabo lic error characterized by glycogen accumulation in all tissues, particular ly striated muscle, due to acid-glucosidase (GAA) deficiency. Respiratory dysf unction is a hallmark feature of this disorder and muscle weakness is viewed as the underlying cause, although a potential ne ural contribution has not been explored. We thus examined be havioral and neurophys iological aspects of breathing in an animal model of Pompe disease, the Gaa-/mouse, and in a second transgenic line (MTP) expressing GAA only in skeletal muscle. Glycogen content was significantly elevated in Gaa-/mouse cervical spinal cor d, including identified phrenic motoneurons. Ventilation was assessed via barometric plethysmography and was attenuated during both quiet breathi ng and hypercapnic challenge in Gaa-/mice (6 to >21 months of age) vs. wild-type controls. To verify hypove ntilation, conscious arterial blood samples were collected which demonstrated reduced arterial partial pressure of O2 in Gaa-/mice vs. controls. We confirmed th at MTP mice had normal diaphragm

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xiv contractility; however MTP mice had ventilation similar to the Gaa-/mice during quiet breathing. Neurophysiological r ecordings demonstrated that efferent phrenic nerve inspiratory burst amplitudes were substantially lower in anesthetized Gaa-/and MTP mice vs. controls under standa rdized conditions. We conclude that neural output to the diaphragm is deficient in Gaa-/mice, and therapies targeting muscle alone may be ineffective in Pompe disease. Proof of concept studies we re initiated to treat the respiratory impairments of Gaa-/mice using adeno-associated viral vectors for gene therapy. Intrathoracic injection of AAV serotype 1 containing a control gene tran sfected the diaphragm, but did not reach therapeutic levels in the motone urons at the dose utilized in this study. In order to treat both the muscular and neural components of Pompe disease, higher doses of AAV1 may be necessary with the relatively non-i nvasive intrathoracic injection.

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1 CHAPTER 1 BACKGROUND History of Pompe Disease In 1932 J.C. Pompe discovered glycogen accu mulation in vacuoles of all tissues examined in a 7-month old female who had died suddenly of what was thought to be pneumonia (Pompe 1932; translated in Melvin 20 00). He was the first to correlate this clinical phenotype with vacuolar accumulation of glycogen. The disease was subsequently termed Pompe disease, but is also known as Glycogen Storage Disease Type II, acid maltase deficiency and glycogenosis type II. There were additional reports in the 1930Â’s and 1940Â’s of excess glycogen in tissues of patients with hypotonia and cardiac hypertrophy that resulted in infantile fatalities (Clement and Godman 1950; Di Sant'Agnese, Andersen et al. 1950; Zellweger , Dark et al. 1955). Although pathology associated with glycogen metabolism was characterized by Cori in 1954 (Cori and Schulman 1954), the biochemical basis for Po mpe disease was not elucidated until 1963. Two discoveries were made that year: 1.) th e lysosome was identified as a cell organelle (de Duve 1963) and 2.) the enzyme acidglucosidase was characterized and found to be absent in patients with Pompe disease (H ers 1963). These two breakthroughs laid the groundwork for Pompe disease as well as many other (>40) lysosomal storage diseases. Glycogen Storage and Breakdown Because maintenance of blood glucose levels must be tightly regulated within the body, the storage of glucose in the form of glyc ogen is important for cells to have glucose available in times of need. Glycogen is formed in the cytoplasm of all cells via

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2 glycogenesis, but is most abundant in muscle (1% wet weight) and liver (7% wet weight) (Groff 1999). Tissues are able to utilize glycogen by conver ting it to glucose through the process of glycogenolysis. Skeletal muscle uses glycogen for ener gy during exercise and times of stress, while the liver utilizes glycogen to have glucose available during hypoglycemia, and in response to epinephrine or glucagon release. Stored glycogen is also present in the nervous system to defend against hypoglycemia and hypoxia (Chen 1995). When excess amounts of stored glycogen have accumulated in the cytoplasm, they are tagged to be transported to the lysosome for degradation (Geddes and Stratton 1977). It is not understood what signal sends glyc ogen to the lysosome, but this organelle processes and degrades complex substances su ch as proteins, phospholipids, nucleic acids and carbohydrates (de Duve 1963). For this reason, it is hypothes ized that the cell recognizes that sufficient or excess glycogen is present in the cytoplasm. Once in the lysosome, glycogen is hydrolyzed by acid -glucosidase (GAA) to form glucose (Hers 1963). This glucose is then tran sported via GLUT transporters to be utilized by the cell. GLUT 2 is known to transport glucose in liver, GLUT 3 in neurons and GLUT 4 in muscle (Groff 1999). Biochemistry of Acid -glucosidase Acid -glucosidase (GAA) is a lysosomal enzyme that hydrolyses glycogen, maltose and isomaltose at -1,4 (Lejeune, Thines-Sempoux et al. 1963) and -1,6 (Jeffrey, Brown et al. 1970) gl ycosidic linkages. It is act ive in an acidic environment (4.0-5.0 pH), but cleaves glycoge n most efficiently at a pH of 4.4 (Koster and Slee 1977; Murray, Brown et al. 1978). GAA is first tran slated as a 110 kDa product, and undergoes post-translational modification in the endoplasmic reticulum (Oude Elferink, Strijland et

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3 al. 1984). It is then transported from the cis to trans golgi (Pohlmann, Waheed et al. 1982), after a mannose-6-phosphate group is exposed (Tsuji and Suzuki 1987). The GAA-mannose-6-phosphate complex then ex its the golgi via a clathrin coated vesicle (Gonzalez-Noriega, Gr ubb et al. 1980). The vesicle is transported to a prelysosome, or endosome that has a low pH, and the clathrin coating is removed. This low pH dissociates the mannose-6-phosphat e group from GAA and allows mannose-6phosphate re-uptake to occur in the cell membra ne and the golgi for it to be recycled. GAA then fuses with a primary lysosome where it is processed to form a 95 kDa intermediate and finally two mature forms th at are 76 and 70 kDa. It is these mature forms that are capable of cleaving the -1,4 and -1,6 glycosidic bonds for the degradation of glycogen (Kornfeld 1986). Although 80-90% of GAA is transported to the lysosome, 10-20% is actually secreted from the cell following removal from the trans golgi (Willingham, Pastan et al. 1981). This secreted GAA can then bind to mannose-6-phosphate receptors on adjacent cell membranes, where they are transported to pre-lysosomes and truncated to the 76 kDa and 70 kDa mature forms. The secretion cap ability of GAA is the basis for both enzyme replacement therapy and gene therapy, wh ereby cells deficient in GAA can undergo uptake of GAA through the mannose-6-phosphate receptor on the cell membranes. Gene Expression of Acid -glucosidase The promoter for acid -glucosidase (GAA) has typical housekeeping characteristics, as it has a high GC cont ent and does not contain TATA and CCAAT motifs. Despite the housekeepi ng characteristics of the pr omoter, it appears to have tissue specific regula tion (Raben, Nichols et al. 1996; Ohtsuka, Ishibashi et al. 1999; Ponce, Witte et al. 1999; Yan, Raben et al . 2001; Yan, Raben et al. 2002; Yan, Raben et

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4 al. 2002) via the Notch pathway. The Notch intracellu lar domain is translocated to the nucleus, where it associates with a DNA bindi ng protein that can activate transcription (Ohtsuka, Ishibashi et al. 1999). The probability that transcription will occur is increased with the presence of Hes-1 (Yan, Raben et al. 2002; Yan, Raben et al. 2002), a known downstream target of the Notch pathway. It is understood that GAA is under the regulation of the Notch/Hes-1 pathway, but th is pathway has been elucidated only in hepatoma (Yan, Raben et al. 2002) and fibroblas t (Yan, Raben et al. 2002) cell types. Hes-1 subsequently binds to a 25 bp el ement on the GAA gene that is 1711 bp downstream of the start of e xon 1 (Yan, Raben et al. 2001). When Hes-1 binds to this element, it acts as a silencer in hepatoma cells, but an enhancer of GAA in fibroblasts (Yan, Raben et al. 2002; Yan, Raben et al. 2002 ). In addition, in neuronal cells it has been shown that both Hes-1 and Hes-5 are necessary for the inhi bition of neurogenesis during development (Ohtsuka, Ishibashi et al . 1999). It can be hypothesized that Hes-1 would increase the transcription of GAA (thus making the 25 bp element an enhancer) in neuronal cells, since GAA is highest in neur onal tissue compared to all other tissue during murine development (Ponce, Witte et al. 1999). Pompe Disease Pompe disease is a recessive disorder that results in little to no acid -glucosidase (GAA) activity, eventually lead ing to lysosomal glycogen a ccumulation. Its prevalence has been debated, but ranges from 1/40,000 to 1/300,000 births worldwide (Lin, Hwang et al. 1987; Bashan, Potashni k et al. 1988; Ausems, Verbie st et al. 1999; Poorthuis, Wevers et al. 1999). The variants of Po mpe disease have been divided into 3 conventional groups, but there also appears to be a continuum of the disease. The 3 traditional groups are divided by the age of onset which typically correlates with the

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5 percent of GAA activity, such that the higher the GAA activity, the late r the age of onset (Hirschhorn 1995). The most severe form of Pompe disease results in cardio-resp iratory failure by 2 years of age and is associat ed with no residual GAA enzyme activity (reviewed in: (Hirschhorn 1995; Reuser, Kroos et al. 1995; Raben, Plotz et al. 2002)). They present with clinical symptoms including skelet al muscle weakness and hypotonia, cardiac hypertrophy, respiratory insufficiency, hepato megaly and macroglossia by 7 months of age. The left ventricular wall becomes th ickened, which prevents ventricular outflow, thus, initiating a cyclical cascade to further incr ease the size of the left ventricle (Seifert, Snyder et al. 1992). The cardiac involvement can be observed in an electrocardiogram by a shortened PR interval and a large QRS complex (Ruttenberg, Steidl et al. 1964), indicative of myocardial ischemia. Re spiratory muscle weakness is compounded by mechanical stress on the bronchi and l ungs due to the cardiac enlargement. Juvenile onset patients have 3-10% of normal GAA activity, with a mild (if any) cardiac involvement (Hirschhorn 1995). They present during the first and second decade with progressive proximal muscular weakness, mild he patomegaly and respiratory deficits. Respiratory failure is ultimately the cause of deat h for these patients, which is similar to the adult-onset group. Adult-onset patients develop symptoms between the third and seventh decade with limb-girdl e muscular weakness (Hug, Schubert et al. 1973). They typically have GAA enzyme activities ~15-20% of normal (Hirschhorn 1995). 30% of patients also have respiratory insufficiency at the time of diagnosis. The onset of respiratory symptoms cannot be predicted by level of enzyme activity

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6 (Hagemans, Winkel et al. 2005). However, most juvenile and adult onset patients become ventilator-dependent and ultimately die of respiratory insufficiency. Even though there are three classical groups of Pompe disease, there are some patients that do not fall within these parame ters. The continuum of Pompe disease was developed after the examination that several patients presented with clinical symptoms similar to infantile-onset patients, but with less severe cardiac involvement (Slonim, Bulone et al. 2000). These patients have survived past infanc y with the support of mechanical ventilation. Within this subset of patients, age of onset correlates with cardiac severity. Target Organs Skeletal and Cardiac Muscle Despite the systemic accumulation of lyso somal and cytoplasmic glycogen, skeletal muscle is the most obvious tissue affected by Pompe disease pathology and therefore, most widely studied in this population. Four hypotheses have been generated to explain the muscle involvement of Pompe disease (Hirschhorn 1995): 1. lysosomes rupture causing mechan ical stress on the myofibrils 2. a toxic metabolite is formed when lyso somes do not degrade as they should 3. calcium regulation is altered when the sarc omlasic reticulum membrane is depleted because excess lysosomal membranes are recruited 4. the normal metabolic properties of muscle inherently have more glycogen within the cell; when glycogen degradation is impaired, it is more severe in muscle that already has elevated glycogen compared to other tissues Brain and Spinal Cord Although glycogen accumulation in cardiac and skeletal muscle has clear pathological consequences, the presence of glycogen accumulation in the central nervous

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7 system (Mancall, Aponte et al. 1965; Hug 1967; Hogan, Gutmann et al. 1969; Gambetti, DiMauro et al. 1971; Ma rtin, de Barsy et al. 1973; Martin i, Ciana et al. 2001) has not been fully evaluated at the functional level. Supported by the overall sparing of cortical neurons, cognitive deficits have not been reported in infantile or later-onset patients. It does appear that motor neurons may be mo re prone to develop glycogen accumulation (Mancall, Aponte et al. 1965; Hogan, Gutmann et al. 1969), which could exacerbate the motor deficits caused by muscle weakness. Two case reports have demonstrated elevated spontaneous activity measured by EMG (Hogan, Gutmann et al. 1969), which could indicate both a muscular and neural mechanis m being altered. These patients had normal conduction velocity which is expected becau se Pompe disease patients have normal glycogen content in axons (Hogan, Gutmann et al. 1969; Gambetti, DiMauro et al. 1971) with no measured de-myelination. Case repo rts of neuronal involvement vary, but the anterior horns of the spinal cord emer ge as the most susceptible to glycogen accumulation with distortion of cell bodies pr esent (Clement and Godman 1950; Mancall, Aponte et al. 1965; Hogan, Gutmann et al . 1969; Martin, de Barsy et al. 1973; Manktelow and Hartley 1975). Th e clinical relevan ce of this glycogen accumulation in the spinal cord was evaluated with this Thesis. In addition to the presence of glycogen accumulation in the spinal cord of Pompe disease, murine studies have found that GAA is highly elevated (vs. other tissues) in the nervous system throughout development (Ponce, Witte et al. 1999). It has also been shown that transcription factors known to regulate GAA are necessary for neuronal differentiation (Ohtsuka, Ishibashi et al. 1999) . Therefore, the accumulation of glycogen

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8 may not be the only detrimental effect for Po mpe disease patients in the nervous system, as GAA also regulates unknown mechanis tic functions during development. Animal Models Multiple natural and genetically engineered models of Pompe disease can be utilized to study the disease progression and possible treatments. Naturally occurring models include Brahman (O'Sullivan, Healy et al. 1981) and Short horn (Jolly, Van-deWater et al. 1977; Howell, Dorling et al. 1981 ) cattle, cat (Sandstrom, Westman et al. 1969), dog (Walvoort, Slee et al. 1982), sheep (Manktelow and Hartley 1975) and quail (Usuki, Ishiura et al. 1986). However, th e cat, dog and sheep models have not been maintained. The cattle strains recapitulate the juvenile-onset pr ogression of Pompe disease (Howell, Dorling et al. 1981), but the obvious la rge size of cattle prevents treatment evaluation from being cost-effective. In addition, the long gestation period and single baby births in cattle ar e other determinants. The sma ll sizes of quail make them a useful model for evaluating treatments. Th is model has residual levels of enzyme activity, which offers a good representation of adult-onset Pompe disease. However, quail have two genes that encode GAA and they do not have a high homology with the human counterpart (Usuki, Ishi ura et al. 1986). To better en hance the understanding of Pompe disease, three mouse models were de veloped with various disease progressions (Raben, Nagaraju et al. 1998; Bijvoet, Van Hirtum et al. 1999; Raben, Nagaraju et al. 2000). The two most widely studied mouse models are the exon 13 knockout (Bijvoet, Van Hirtum et al. 1999) and the exon 6 knoc kout (Raben, Nagaraju et al. 1998). The exon 13 knockout was developed by targeted disr uption with the neomycin gene cassette in exon 13. This knockout ha s no detectable protein and GAA enzyme activity levels

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9 similar to infantile-onset patients, with glycogen accumulation visible by periodic acid Schiff staining. However, the exon 13 knoc kout does not show any objective clinical signs of muscle weakness until 9 months of age when posture and gait problems arise (Bijvoet, Van Hirtum et al. 1999). Alt hough the exon 6 knockout has similar glycogen accumulation and GAA activity compared to th e exon 13 knockout, it has a more severe clinical phenotype. This knockout was developed by insertion of the neo cassette into exon 6. At 3.5 weeks of age the exon 6 knockout has significantly le ss locomotor activity in the open field test when compared to their wild-type littermates (Raben, Nagaraju et al. 1998). In addition, our group has demonstrated contractile dysfunction in these mice by 3 months of age (Fraites, Schleiss ing et al. 2002). One hypothesis for the difference in thes e two knockout strain s is the genetic background. The exon 13 knockout is maintained on the 129SvJ x C57BL/6 or 129SvJ x FVB backgrounds (Bijvoet, Van Hirtum et al. 1999), while the exon 6 knockout is maintained on the 129SvJ x C57BL/6 backgroun d (Raben, Nagaraju et al. 1998). Each background strain could have various compensato ry mechanisms that alter the severity of the disease (Erickson 1996; Wilson 1996). Although the heterozygosity of the mixed genetic background can cause difficulties with characterizing the disease phenotype, it is representative of the heterozygosity of th e human population. This will allow us and other investigators to anticipa te the broad range of reactio ns in response to different treatment regimens. We have chosen to study the exon 6 knockout in our studies, because it has a critical phenotype that is ev ident by 1 month of age (Raben, Nagaraju et al. 1998). It is more practical to study this mouse because it develops symptoms earlier and therefore, we can uncover if gene therapy treatments are effectiv e at an earlier age.

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10 In addition to gene therapy, we have characte rized the ventilation of these mice starting at 6 months of age. Althou gh the previous functional m easures used to determine differences in GAA knockout mice and their wild-type controls did increase our knowledge of the GAA knockout phenotype, some measures were subjective (rotorod and open field test (Raben, Na garaju et al. 1998)), while ot hers required sacrificing the animal to obtain the data (Fra ites, Schleissing et al. 2002). By quantifying ventilation in the GAA knockout mouse with barometric plethysmography, we were able to obtain whole animal conscious physiologic measurements that can be utilized over time in the same animal. These experiments have in creased our understanding of the respiratory deficits in the Gaa-/mouse model, as well as the e ffectiveness of gene therapy on ventilation. Since respiratory insufficiency is the main cause of death in juvenile and adult-onset patients, the data generated from these experiments will be clinically relevant for treatment of Pompe disease. Treatments for Pompe Disease Enzyme Replacement Therapy Since the initial disc overy by Hers that Pompe disease was a result of a deficiency in a lysosomal enzyme (Hers 1963), investig ators have been working to exogenously supply GAA. Initially, GAA purified from Aspe rgillus niger was injected via both i.m. (Baudhuin, Hers et al. 1964) and i.v. (Hug a nd Schubert 1967) routes with no success. GAA isolated from human placenta showed nega tive results as well (R euser, Kroos et al. 1984). In 1984, a landmark study illustrate d that phosphorylated GAA isolated from Bovine testes could be taken up by cells in culture by the mannos e-6-phosphate receptor on the cell membrane (Reuser, Kroos et al. 198 4). It was later determined that this GAA was also incorporated into the lysosome and processed to the mature form, resulting in

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11 clearance of glycogen from Pompe disease cell lines (van der Ploeg, Kroos et al. 1987). In addition, Bovine testes GAA was also taken up in a perfused rat heart through the mannose-6-phosphate receptor, while human pl acental GAA was again ineffective, as it is not phosphorylated (van der Ploeg, van de r Kraaij et al. 1990). Bovine testes GAA was next administered in vivo to mice, wher e it was shown to increase GAA activity in every tissue but brain (Van der Ploeg, Kroos et al. 1991). Despite the success of using this purified form of GAA, interspecies antigen responses prevent Bovine GAA from use in human patients. For this reason, studies were pursued that isolated GAA from human urine to treat glycogen accumulation in cells (van der Ploeg, Bolhui s et al. 1988). These investigations proved successf ul (van der Ploeg, Bolhuis et al. 1988; Van der Ploeg, Loonen et al. 1988), but only a small amount of GAA can be purified from human urine. Again, a new approach was needed. Clonal ce ll lines were then used to produce larger amounts of purified GAA (Fuller, Van der Ploe g et al. 1995; Van Hove, Yang et al. 1996) which were used in a phas e I/II clinical tr ial (Amalfitano, Bengur et al. 2001). Recombinant human GAA produced in the milk of transgenic animals was also proven to be effective (Van den Hout, Reuser et al . 2000; Winkel, Kamphoven et al. 2003; Van den Hout, Kamphoven et al. 2004). Currently, enzyme replacement therapy (ERT) is available to some Pompe dis ease infantile and juvenile on set patients that qualify as clinical trial participants. The results from th e ERT clinical trials may allow for this to be an optional treatment for all Pompe disease patients, although it will clearly be quite costly. Even though the results are promising fo r ERT, it is important to note that GAA cannot pass through the blood brai n barrier. Therefore, if a central nervous system

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12 component emerges in Pompe disease, ERT will not be able to decrease the excess glycogen content in the spinal cord or brain that may be causing a functional pathophysiology. A main goal of this thesis was to determine if a central component contributed to the respiratory in sufficiencies in Pompe disease. Gene Therapy A therapeutic approach that may be able to access the CNS through retrograde transport is gene therapy (M artinov, Sefland et al. 2002; Ka spar, Llado et al. 2003). The interest in gene therapy first arose as an alte rnative to ERT. ERT is limited by the cost of producing enzyme on a large scale and the need for infusions throughout the life of the patient and the possible immunogenic problems (Raben, Danon et al. 2003). The concept of gene therapy is based on a one time inj ection that would deliver the missing DNA to deficient cells. GAA would then be transcribed and could be used by the host cell and secreted to neighboring cells. Adenovirus was the first vector used to pr ovide the proof of concept studies that GAA could be delivered to defi cient cells both in vitro (Nicolino, Puech et al. 1998) and in vivo (Tsujino, Kinoshita et al. 1998) that re sulted in decreased glycogen content. Later studies demonstrated that GAA was being targeted to the lysosome, where it was processed to the mature 76 and 70 kDa forms (P auly, Johns et al. 1998). The first in vivo experiments were performed in quail wh ich contains two genes for GAA (Tsujino, Kinoshita et al. 1998). To identify if Ade novirus could be effective in the mammalian system, Adenovirus carrying the GAA gene was in jected into neonatal rats that resulted in increased activity of GAA that was present in lysosomes (Pauly, Johns et al. 1998). The next important study came when a mous e model of Pompe disease was developed (previously described, exon 6 knockout (Rabe n, Nagaraju et al. 1998)). Although the

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13 previous studies with adenovirus proved to transduce cells, the delivery of GAA was somewhat limited to surrounding tissues. Th erefore, for adenovirus to be effective, multiple injection sites would be needed to correct patients with Pompe disease. For this reason, the liver was targeted as a depot for GAA. Overexpression in the liver was hypothesized to saturate the liver GAA, whic h would then secrete the enzyme into the circulation where it could corr ect other tissues. This method was successful by reducing glycogen content of the hear t and diaphragm, as well as other tissues (Amalfitano, McVie-Wylie et al. 1999). Adenoviral vectors did provide the proof of concept that GAA could be delivered to deficient cells. However, multiple aut hors of these previous papers recommend against using adenovirus in hum ans with Pompe diseaseGSD II (Pauly, Johns et al. 1998; Amalfitano, McVie-Wylie et al. 1999). Ade novirus is immunogenic (Schnell, Zhang et al. 2001), and has resulted in one fatality in an adenoviral clinical tr ial (Raper, Chirmule et al. 2003). Its expression is known to be transient (Xu, Mizuguchi et al. 2005), which is an obvious downfall for Pompe disease patien ts, who need GAA activity throughout life. Adenovirus has safety issues, which must be overcome before they can be used for Pompe disease. The viral vector of choice for our studie s is adeno-associated virus (AAV). We have chosen this vector based on its safety and long-term expression of the packaged transgene (Fisher, Jooss et al. 1997). AAV has never been associated with any human pathogen or disease (review of AAV in: (Kotin 1994; Flotte and Carter 1995; Hildinger and Auricchio 2004)). Without a he lper virus, it develops as a latent infection in its host. Even though it is latent, DNA packaged into AAV can be expressed once AAV is in the

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14 body; this is without replication of the virus itself. Another safety feature of AAV is that almost the entire viral genome can be gutte d and replaced with a chosen promoter and transgene. With the viral genome replace d, immune responses are greatly reduced compared to other viral vectors, including aden ovirus. In addition to the safety of AAV, it is also resistant to chemi cal and physical treatment (Wright , Qu et al. 2003). This is quite beneficial for the production and purification of AAV stocks. AAV serotype 2 (AAV2) was the first AAV genome to be sequenced and subsequently used for gene therapy experi ments. The initial work on AAV as a viral vector for gene therapy was all perf ormed using AAV2. With AAV2, sustained expression (>12 months) of various transgenes was found in multiple different disease models (Herzog, Hagstrom et al. 1997; Song, Mo rgan et al. 1998; Chao, Mao et al. 2000; Chao, Monahan et al. 2001; Wang, Calcedo et al. 2005). As th e progress of AAV mediated gene therapy grew with the reality to have long term ge ne expression following AAV administration, multiple other serotype s of AAV were discovered which allowed for tissue specificity of the viral capsid (re viewed in: (Muzyczka and Warrington 2005)). Because the safety of the AAV2 genome had b een previously elucid ated, a process called pseudotyping was employed for using the ne w capsids of AAV. With psuedotyping, the AAV2 genome is inserted into the capsid of serotypes AAV1-9. This allows for different tissue tropism via the capsi d, with the known safety of the viral genome of AAV2. Our group and others have shown that AAV can be used to treat mouse models of Pompe disease (GAA-/-; exon 6) (Fraites, Schlei ssing et al. 2002; Ru cker, Fraites et al. 2004; Sun, Zhang et al. 2005). Fr aites et al was the first to demonstrate that both AAV2 and AAV1 could increase GAA activity in skeletal muscle of Gaa-/mice (Fraites,

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15 Schleissing et al. 2002). In terestingly, AAV1 increased GAA activity to a much higher degree compared to AAV2, although both were effective at recovering muscle, tested either by contractile function (AAV2) or glycogen conten t (AAV1). These responses were measured for up to 24 weeks post-inje ction. Our group has also demonstrated correction of Gaa-/embryonic diaphragm with delivery of AAV2 carrying the GAA gene to mice in utero (Rucker, Fr aites et al. 2004). GAA activity persisted in these mice, as well as clearance of glycogen visible by histology. The em bryonic data are impressive; however, injections into pre-term or neona tal mice prevent any i mmune response to the transgene, and do not give a clear picture for the effectiveness of GAA delivery in immune competent individuals. Importantl y, these experiments la id the groundwork for the possibility of in utero gene therapy. Nevertheless, the immune response to GAA will be important for clinical trials in patient s with no residual GAA. Patients with some GAA activity will be tolerized to GAA, and will not need to circumvent the immune system. Reports using the Gaa-/mouse have shown signifi cant antibody elevation to GAA delivered both by enzyme (Raben, Danon et al. 2003) and gene therapy (Cresawn, Fraites et al. 2005) in non-tole rized mice. However, the hear t and diaphragm of tolerized mice responded quite well to a portal vein injection of AAV5 and AAV8-GAA (Cresawn, Fraites et al. 2005). Due to the size of the AAV icosohedral ca psid (20 nm; (Xie 1999)), it cannot be transported across the blood brain barrier. To circumvent this obstacle, molecules that extravasate the blood brain barrie r and allow it to be less sele ctive have been utilized to access the central nervous system with AAV (Bat es, Hillman et al. 2002). Extravasation has it problems, as decreased blood pressure and fluid accumulation result. It is not

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16 known if these detriments would prevent th is therapy from being used in already unhealthy patients. However, a recent repo rt has demonstrated that following an intramuscular injection of AAV2, the cap sid can transfect the CNS via retrograde transport from the muscle (Kaspar, Llado et al. 2003). Adenovirus has also been shown to be transported in a retrograde fashion from both slow and fast twitch muscle fibers (Martinov, Sefland et al. 2002). Retrograde tr ansport is a new phenomenon in the gene therapy field, and has only recently been reported by one group using AAV2 who were able to correct a mouse model of ALS (Kaspa r, Llado et al. 2003). At the time of their publication, AAV2 was the predominant serotype used in gene therapy experiments. Since this short time, however, new sero types have been developed that have various tropisms for motor neurons, and glia, in addition to serotypes with high muscle tropism. The most exhaustive study to date compared serotypes 1,2 and 5 after direct injection into various regions of the brain and spinal co rd (Burger, Gorbatyuk et al. 2004). AAV1 and AAV5 were found to tran sduce many more neurons compared to AAV2, with AAV1 and AAV5 both observed to have anterograd e and retrograde transport. In contrast to the regions of neuronal transduction in the previous publication, AAV5 has been shown to transduce both glia and neurons in striatum and cortex (Davidson, Stein et al. 2000). These studies cannot be directly compared for two reasons: 1.) different regions of the CN S were injected with the vari ous serotypes of AAV and 2.) different promoters were used. However, it can be stated that both AAV1 and AAV5 are useful for targeting the CNS. The use of AAV3 and AAV4 has been ruled out as a possible use in the CNS, as both serotypes have little to no transduc tion of neural tissue (Davidson, Stein et al. 2000). Serotype 6 has shown similar tropism for the CNS as

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17 serotype 1, which is not surprising consid ering that have very similar homology (Huszthy, Svendsen et al. 2005). Serotypes 7 and 8 were isolated in 2002, with both having similar tropism (Gao, Al vira et al. 2002). As a re sult, studies have focused on AAV8. Although AAV8 transduces vastly more cel ls compared to all other serotypes, it also infects the gonads after an intravenous injection (Wang, Zhu et al. 2005), a detriment for the use of this serotype in clinical tr ials. AAV7 and AAV8 are not neutralized by human antibodies (Gao, Alvira et al. 2002), which is hypothesized to be a main reason for its strong ability to infect and transduce multiple tissues. In addition to the tropisms of the vari ous serotypes for the CNS, tropism for skeletal muscle has been ex tensively studied. Most re cently, serotypes 1,2,5,6,7 and 8 were been compared after an intravenous injection (Wang, Calcedo et al. 2005). Serotype 8 was up to 100 times more efficien t at transducing muscle compared to the others, but as mentioned previously, also infects the gonads. From this study, AAV2 and AAV5 had very low transduction efficiency comp ared to the others. However, De et al have demonstrated robust transduction of th e diaphragm after an intrapleural injection with AAV5 (De, Heguy et al. 2004). Fo llowing an intramuscular injection, AAV1 transduced the most muscle fibers when compared to AAV2 and AAV8 (Wang, Calcedo et al. 2005). In yet another comparison, sero types 1-5 were evaluated 8 weeks after an intramuscular injection. 1,3,4 and 5 all performed better than AAV2 (Chao, Liu et al. 2000). However, in a disease model, AAV1 and AAV5 were most effective at recovering the clinical phenotype (Chao, Liu et al. 2000). Clearly, the route of administration is very important for tissue tr opism. Nonetheless, serotypes 1,5 and 8 are capable of transducing muscle with relatively high affinity.

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18 It is a goal of this thesis to use a sero type that has high affinity for both motor neurons and muscle, without germline tran smission. AAV1 and AAV5 emerge as the best options for our studies. Summary Pompe disease is a lysosomal storage di sorder caused by deficiency of acid glucosidase. It has predomin antly been studied as a diseas e of the musculature, with respiratory insufficiency resulting as the majo r cause of death in later-onset patients. Preliminary data for this thes is led us to hypothesize that a neural component may also contribute to the muscular weakness. To th is end, we characte rized the pattern of breathing in the mouse model of Pompe diseas e, and quantified motor nerve activity to the diaphragm. In addition, the capacity of adeno-associated vi rus serotype 1 to transduce the diaphragm and ga in access to the central nervous system through retrograde transport was investigated. These experiment s identified a neural component contributes to diaphragmatic muscular weakness in Pomp e disease and proof of concept studies to utilize AAV1 as a possible treatment for the neural deficits of Pompe disease were initiated.

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19 CHAPTER 2 RESPIRATORY AND NEURAL CHARACTE RIZATION OF A MOUSE MODEL OF POMPE DISEASE Introduction The storage of glucose in the form of gl ycogen is important for all cells, including neurons, to have energy available in times of increased demand (Chen 1995; Ransom and Fern 1997). However, the physiological cons equences of excess glycogen accumulation within the central nervous system (CNS ) are currently unknown. This issue is particularly relevant with re spect to Glycogen Storage Diseas es, such as Pompe disease. Pompe disease is both a lysosomal and gl ycogen storage disorder resulting from deficiency of the enzyme acid -glucosidase (GAA) (H irschhorn 1995). GAA is normally active in the lysosome where it degrades excess glycogen by cleaving the -1,4 (Lejeune, Thines-Sempoux et al. 1963) and -1,6 (Jeffrey, Brown et al. 1970) glycosidic bonds. Without adequate GAA activity, massive amounts of glycogen accumulate in all cells. Despite systemic accumulation of lyso somal glycogen in Pompe disease, skeletal and cardiac muscle dysfunction have been cla ssically viewed as the principle basis for muscle weakness in this disorder (reviewe d in:(Hirschhorn 1995; Re user, Kroos et al. 1995; Raben, Plotz et al. 2002)). Respiratory dysfunction in Pompe disease is a significant clinical feature that contributes substantially to morbidity and mort ality. The most severe form of the disease is associated with complete absence of GAA activity, early onset of symptoms and cardio-respiratory failure by 8-10 months. Typi cally, a later onset of clinical symptoms

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20 is associated with partial GAA activity. While cardiac involveme nt is mild in these later onset patients, respiratory insufficiency leads to ventilator dependency and ultimately respiratory failure in juvenile and adult Pompe disease individuals (Hirschhorn 1995). Many animal models are available to study Pompe disease (O'Sullivan, Healy et al. 1981; Usuki, Ishiura et al. 1986; Raben, Nagaraju et al. 1998). Of these, the Gaa knockout mouse ( Gaa-/-) closely mimics human disease pathology, including the chronic aspects of disease progression (Rab en, Nagaraju et al. 1998). The Gaa-/mouse, developed by Raben in 1998, shows dia phragm glycogen accumulation as well as in vitro contractile weakness (Fraites , Schleissing et al. 2002; Cres awn, Fraites et al. 2005). Although it is generally accep ted that diaphragmatic muscle dysfunction is the primary reason for ventilation deficits in Pomp e disease, there has been no formal effort to discount a neural contribu tion to the pathophysiology of this disease. However, clinical case reports have demonstrated si gnificant glycogen accumulation in the central nervous system (CNS) (Mancall, Aponte et al. 1965; Gambetti, DiMauro et al. 1971; Martin, de Barsy et al. 1973), and absent or diminished deep tendon reflexes in Pompe disease patients (Hogan, Gutmann et al. 1969). In particular, spinal motoneurons seem to be susceptible to excessive glycogen accumu lation (Mancall, Aponte et al. 1965; Hogan, Gutmann et al. 1969). The physiological consequenes of glycogen accumulation in motoneurons are unknown, but the large, swol len appearance of th ese cells (Mancall, Aponte et al. 1965; Gambetti, Di Mauro et al. 1971) suggests that they may not respond to “normal” depolarizing synaptic input. A ccordingly, we postulated that GAA deficiency in the nervous system may contribute to respiratory insufficiency and tested the

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21 hypothesis that Gaa-/mice would exhibit reduced ventil ation that was associated with attenuated efferent phrenic motor discharge. Our results demonstrate that neural drive to the dia phragm is compromised in Gaa/vs. control mice. Accordingly, these findings raise important therapeutic considerations for Pompe disease since the only currently available strategy, enzyme replacement, cannot effectively target GAA deficiency and glycogen accumulation of the CNS (Kikuchi, Yang et al. 1998; Raben, Lu et al. 2001). Methods Animals The Gaa-/mouse was generated by targeted di sruption of exon 6 and is maintained on the C57BL/6 X 129X1/SvJ background (described previously: (Raben, Nagaraju et al. 1998). Contemporaneous gender matched C 57Bl/6 X 129X1/SvJ mice were used as controls for all experiments. Muscle-sp ecific hGAA mice (referred to as MTP mice: 18) were generated by developing a clone with th e tetracycline-responsiv e element linked to hGAA. This fragment was then microinject ed into FVB mouse embryos. The FVB mice were mated to Mck-t/-/mice (purchased from Jackson Laboratories, C57BL6/129 SVj background strain); their o ffspring were mated to Gaa-/mice to result in mice that express hGAA only in muscle (described prev iously in: (Raben, Lu et al. 2001). Mice were housed at the University of Florida sp ecific pathogen-free animal facility. The University of FloridaÂ’s Institutional An imal Care and Use Committee approved all animal procedures. Barometric Plethysmography A flow-through (0.5 L/min) barometric pl ethysmograph (Buxco Inc., Wilmington, NC) was used to measure the pa ttern of breathing in control, Gaa-/and MTP mice. A

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22 clear Plexiglas chamber (diameter 3.5” a nd height 5.75”) was calibrated with known volume signals prior to data collection. Signals were analyzed online using the Drorbaugh and Fenn equation (Drorbaugh and Fenn 1955) to provide a breath-by-breath display of ventilation. The following variab les were measured: inspiratory frequency, tidal volume, peak inspiratory flow, peak e xpiratory flow, inspiratory time, expiratory time, and expired carbon dioxide. Ventilati on data was collected in 10-s bins and metabolic data in 1-min bins. Ventilation was characterized in both genders to provide descriptive data for future studies using this relatively noninvasive technique. Howeve r, there is no evidence to date that gender differences exist in Po mpe disease patients. Nonetheless, it was important to verify this in the mouse model. Genders have only been separated when differences were detected between male and female mice. Body temperature was measured immedi ately prior to placing mice in the barometric plethysmograph. To account for any circadian differences in the pattern of breathing, mice were tested at the same time of the light cycle. Thirty minutes were allowed for acclimation to the chamber and the following 60 minutes were used as baseline. During both acclimation and base line mice were breathi ng normoxic air (21% O2, 79% N2). Following baseline, mice were expo sed to 10 minutes of hypercapnia (7% CO2, balance O2). Body temperature was measured again immediately following the exposure to hypercapnia. If post body temper ature differed by >0.1 degree Celsius, a temperature correction was applied to th e hypercapnic data using the Drorbaugh and Fenn equation.

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23 Hemoglobin, Hematocrit, Glucose and Sodium Blood Levels Venous tail blood was collected from anes thetized mice (2% isofluorane, balance O2) directly into a commercially available bl ood gas analysis cartri dge (I-stat, Heska Corporation; Ft. Collins, CO). The cartridge was used per the manufacturer’s instruction to measure hemoglobin, hematocrit, glucos e and sodium for each sample (~100 µL). Conscious Arterial Blood Sampling Anesthesia was induced with 3% isoflourane in a closed chamber. Mice were then placed in the supine position and fitted with a nosecone breathing 1-2% isofluorane in 50% O2 (balance N2). Neck fur was removed and the skin scrubbed with povidoneiodine followed by isopropyl alcohol. An incision (~1.5 cm) was made between the jaw and sternum. The left carotid artery was expos ed and tied at the most rostral point. A distal suture was used to ligate the artery a nd a catheter (mouse carotid catheter, Braintree Scientific) filled with heparinized saline wa s inserted into the artery with a needle beveled at 90 . The catheter was advanced, tied off and run under the skin to be exposed between the ears. Two sutures were used at th e ears to secure the catheter. The ventral skin incision was closed with 6.0 vicryl sutu re. Each catheter was attached to a tether ~20 inches long (0.033 outer diameter; renapulse tubing, Braintree Scie ntific; filled with heparinized saline) prior to removing mi ce from anesthesia. Once removed from anesthesia, mice were allowed to recover 310 hours in their home cage prior to blood sampling. Once mouse body temperatures returned to at least 36.5 C, mice were prepared for conscious blood samples. The experimenter would stand over the home cage for 10 minutes prior to sampling holdi ng the tethered catheter in place for the blood sample.

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24 During this time, mouse breat hing would return to a lower frequency. The tethered catheter was cut approximately 10 inches from the animal. Four blood drops (after the saline) were blotted onto a gau ze pad and the blood was then allowed to flow into the IStat cartridge for measuremen t of partial pressure of PO2 (I-stat, Heska Corporation; Ft. Collins, CO). The cartridge was used per the manufacturer’s instruction to measure partial pressure of PO2 (~100 µL). Different mice were used for hemoglobin/hematocrit measures and conscious blood sampling because a saline flush is required to keep the chronic catheter patent which coul d dilute hemoglobin/hematocrit. Glycogen Quantification Fed control and Gaa-/mice were sacrificed during the 3rd to 6th hour of the light cycle with an overdose of sodium pentabar bitol (150/mg/kg). Ce rvical spinal cord segments C3-C5 were harvested and immediately frozen in liquid nitrogen; then stored at -80 C until ready for biochemical analysis. Harvesting of cervical segments C3-C5 was performed by identifying the corr esponding cervical dorsal roots. Glycogen content was measured using a modification to the acid-hydrolysis method (Lo, Russell et al. 1970). This met hod was preferred because it was the most reproducible for glycogen standa rds, muscle and spinal tissu e; and could be performed with 15-20 mg of tissue. Tissue sample s were weighed and placed into a 1.5 mL microcentrifuge tube. A standard curve was generated by using 5 serial dilutions starting with 800 µg glycogen (Roche Diagnostics Corp oration; Indianapolis , IN). Three hundred µL of 30% potassium hydroxide (w/v) saturate d with sodium sulfate was added to each tube, and then heated at 90 C for 3 hours. Samples were cooled on ice for 1 min, and 900 µL of 95% ethanol added. Samples we re kept on ice for 30 minutes, followed by

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25 centrifugation for 30 min (14000 rpm) at 4 C. The supernatant was discarded and 120 µL of distilled water was used to re-suspend the pellet. Twenty µL of each sample was added to 40 µL of 5% phenol, followed by 500 µL of 98 % sulfuric acid. Each sample was run in triplicate and incubated for 24 hour s at room temperature. 200 µL of each sample solution (sample, phenol, sulfuric acid) was added to a 96-well microplate and read at 490 nm absorbance (µQuant microplate reader; Bio-Tek, Inc., Winooski, VT). Values were calculated according to the sta ndard curve and are reported relative to mg wet weight (µg glycogen/mg wet weight tissu e). Importantly, normalizing to wet weight was appropriate because the wet:dry weight ratio was similar for control (3.59) and Gaa-/(3.52) spinal cord. Retrograde Labeling of Phrenic Motoneurons Mice were anesthetized with 2% isoflurane and restrained in the supine position on a warmed operating surface. A laporatomy was performed for access to the peritoneal surface of the diaphragm. The neuronal re trograde tracer fluorogold (4% diluted in sterile phosphate buffered salin e, Fluorochrome, LLC, Denver, CO) was then applied to the diaphragm (~75 µL) using a small artist’s br ush. Care was taken to apply the tracer sparingly only to the diaphragm in order to minimize leakage to liver and surrounding tissues. Complete coverage of the diaphrag m was determined when all regions were yellow in color from the fluorogold. Abdomina l muscle and skin were closed and mice removed from isoflurane and allowed to recover in their home cages. Forty-eight hours after fluorogold applicati on, mice were anesthetized with sodium pentobarbital (60 mg/kg, i.p. ), and then systemically perfused via the heart with 4% paraformaldehyde in phosphate buffered saline. The cervical spinal cord (C3-C5) was

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26 removed, paraffin-embedded and sectione d in the transverse plane at 10 µ m. Embedding for spinal cords was performed by a modified method first described by Guth and Watson (Guth and Watson 1968). Tissue was placed in to a solution of 45 mL 100% ethanol, 2.5 mL of glacial acetic acid and 2.5 mL of 40% formaldehyde for 4 hours. Next, the spinal cords were placed into 95% ethanol for 30 minutes followed by 4 washes (30 minutes each) of 100% ethanol. Samples were then cleared with cedarwood oil for 2 hours and placed directly into paraffin for final embeddi ng procedures. Care was taken to remove the residue left by cedarwood oil every 30 mi nutes throughout the clearing protocol. Flurogold-labeled phrenic motoneurons were identified by fluorescence microscopy. Histological Glycogen Detection in Motoneurons Spinal tissues were stained with standard Periodic Acid Schiff (PAS) methods for histological detection of glyc ogen according to the protocol originally published by Guth & Watson (Guth and Watson 1968). Briefly, spin al cord sections were deparrafinized, incubated in 1% periodic acid for 5 minutes, ri nsed in distilled water, placed into Schiff’s reagent for ~8 minutes, rinsed in sulphurous acid for 6 minutes, rinsed in tap water, dehydrated, cleared and coverslipped. In Vitro Diaphragmatic Contractility Mice were anesthetized with sodium pe ntobarbital (1.5 g/kg body weight). The diaphragm was excised and placed into a dissecting chamber with Krebs-Henseleit solution equilibrated with a 95% O2 / 5% CO2 gas mixture. The diaphragm strip was suspended vertically between two lightwei ght plexiglass clamps connected to force transducers (Model FT03, Grass Instruments, West Warwick, RI) in a water jacketed tissue bath containing Krebs-Hensele it solution equilibrated with a 95% O2 / 5% CO2 gas (bath 37 C, pH 7.4, osmality 290mOsm). Tran sducer output was amplified and

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27 differentiated by operational amplifiers and underwent A/D conversion for analysis using a computer-based data acquisition syst em (Polyview, Grass Instruments). In vitro contractile measurements began with determ ination of the muscleÂ’s optimal length for isometric tetanic tension development. Th e muscle was field-stimulated (Model S48, Grass Instruments) along its entire length with platinum electrodes. Muscle length was progressively increased until maximal isom etric twitch tension was obtained. Once the highest twitch force was achieved, maximum is ometric tetanic tension was measured at optimal length and stimulated at 10, 20, 40 80, 100, 150 and 200 Hertz, while measuring the force produced (N/cm2) at each frequency. At the end of the study, the muscle strip length and weight were determined in order to calculate the normalized force generated. Efferent Phrenic Nerve Recordings Anesthesia was induced with 3% isoflouran e in a closed chamber. Mice were then placed in the supine position and fitted with a nosecone breathing 2% isofluorane in 50% O2 (balance N2). Neck fur was removed and the skin scrubbed with povidone-iodine followed by isopropyl alcohol. An incision wa s made from the jaw to the sternum and the trachea was cannulated with a 20-gauge angi ocath (cut to 1 inch) and connected to the ventilator (Model SAR-830/AP, CWE, Incor porated). Ventilator settings were manipulated to produce partial pressures of arterial CO2 between 45-55 mmHg. A jugular catheter (0.033 outer diameter; renapulse tubing, Braintree Scientific) was implanted and used to transition the mice fr om isofluorane to urethane (1.0 1.6g/kg) anesthesia. Anesthesia was supplemente d (0.2 g/kg urethane) as necessary. The adequacy of anesthesia was monitored init ially by observing move ments during toe pinch and the palpebral reflex. After mice were paralyzed (see below), anesthesia was monitored by observing phrenic nerve responses to toe pinch.

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28 A carotid arterial catheter (mouse carotid catheter, Braintree Sc ientific) was then inserted to enable withdrawal of 0.15ml samples for measuring arterial PO2 and PCO2 (IStat portable blood gas analyzer). Mice we re vagotomized bilaterally and paralyzed (pancuronium bromide; 2.5 mg/kg, i.v. ). Rectal temperature was maintained (37-38°C) with a rectal thermistor and servo-controll ed heating pad. The ri ght phrenic nerve was isolated using a ventral approach and was placed on a bipolar tungs ten wire electrode (0.005 inch diameter). Nerve electrical activ ities were amplified (2000x) and filtered (100-10,000 Hz; Model BMA 400, CWE, Incorpor ated). When monitoring spontaneous inspiratory activity in the phrenic neurogram, the amplified signal was full-wave rectified and smoothed with a time constant of 100 ms, digitized and recorded on a computer using Spike2 software (Cambridge Electronic De sign; Cambridge, UK). The amplifier gain settings and signal processing methods were id entical in all experime ntal animals. The 30-s prior to each blood draw were analyzed for the mean phrenic inspiratory burst amplitude, fictive breathing frequency and th e inspiratory time from these digitized records. At the conclusion of the experiment, mice were eu thanized with a bolus i.v. injection of urethane. Comparisons of efferent phrenic burst ampl itude between experimental groups (i.e. control vs. Gaa -/-) are a fundamental aspect of this study that merits a brief discussion for the validity of the approach. In these experi ments, the distal end of the phrenic nerve was cut to measure efferent extr acellular activity of compound action potentials in the phrenic nerve. However, factors that are essentially beyond the control of the experimenter ( e.g. subtle differences in nerve-electrode contact, nerve diameter, etc. ) have the potential to influence the amplitude of the efferent phr enic burst. Accordingly, many investigators

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29 have used normalization procedures to expr ess phrenic burst amp litude relative to a “maximum” phrenic activity output (e .g. during severe hypercapnia or hypoxia resulting in gasping). However, that approach begi ns with the assumption that maximum is the same or similar between groups. If the ability to recruit phrenic mo toneurons is impaired, then normalizing phrenic activity to a maxi mum will have the effect of eliminating physiologically meaningful differences between experimental groups. It was hypothesized that efferent phren ic discharge is reduced in Gaa-/mice, and therefore, we felt it was not appropriate to normalize the burst amplitudes. Rather, the data are presented as a “raw” voltage. Statistics Statistical significance for this project was determined a priori at p<0.01. The stringent p-value was chosen because the findings of this study may have profound clinical relevance and accord ingly we wanted to minimize the possibility of a Type I error. Ventilation data were analyzed using a three-way anal ysis of covariance (ANCOVA). Ratios of volume:bodyweight we re not used, as body mass ratios can introduce bias and this method does not have the intended eff ect of removing the influence of body mass on the data (Packard and Boardman 1999). By using the ANCOVA method, bodyweight is analyzed as a co-variate for all respiratory volume data, which more accurately removes the influence of bodyweight on our data. For baseline measures, gender, strain and age were used as factors while the hypercapnic data was analyzed using gender, strain and time (mi nutes 1-10 of hypercapnia) as factors. To compare ventilation data of B6/129, Gaa-/and MTP mice, student’s t-test with Bonferroni correction were applied. A 2-way ANOVA was used for diaphragm contractile data of B6/129, Gaa-/and MTP mice. Hemoglobin, hematocrit, glucose,

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30 sodium and conscious arterial PO2 levels were analyzed us ing the student’s t-test. Glycogen quantification was analyzed us ing a two-way ANOVA and t-test with Bonferroni correction for post-hoc measurements. Phrenic inspiratory burst amplitude, breathing frequency and the rate of rise of the phrenic burst were extracted from the phrenic neurogram. These variable s and anesthetized arterial PaCO2 were analyzed with the one-way ANOVA and Fische r’s LSD test for post-hoc analysis. All data are presented as MEAN±SEM. For all respiratory volume da ta described, the analysis of co-variance was applied with bodyweight as the co-variate to remove th e influence of this variable on ventilation. However, in some cases, all assumptions were not met for the ANCOVA. In these instances, volume data were analyzed with ANOVA. We did not f eel it was appropriate to normalize respiratory volumes to bodyweight in these cases fo r three reasons: 1) Packard and Boardman have eloquently i llustrated that norm alizing physiological variables to bodyweight does not always have the intended effect of removing the influence of bodyweight (Packard and Bo ardman 1999), 2.) when the ANCOVA was applied (baseline tidal volume for all groups and hypercapnic responses at 6 months of age), respiratory volumes of Gaa-/were clearly lower (p<0.01) than controls and 2) in many disease states associated with ventil atory dysfunction (anorex ia, cystic fibrosis, COPD), bodyweight decreases with disease progression, but respiratory volumes are not typically normalized to bodyweight in these pa thological states (Nishimura, Tsutsumi et al. 1995; Ionescu, Chatham et al. 1998; Gonzalez-Moro, De Miguel-Diez et al. 2003). Consequently, we feel that ventilation deficits in Po mpe disease could also be

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31 confounded by bodyweight normalizations, especi ally in older populations associated with augmented weight loss. Results Table 2-1. Body weight values for control a nd Gaa-/mice at 6, 12 and >21 months of age. *= Gaa-/different from control values for all age groups. =male different from female for all age groups. No age effect detected. 6 MONTHS 12 MONTHS >21 MONTHS CONTROL: male (g) 39.0 ± 2.3 38.8 ± 2.7 36.2 ± 2.1 Female (g) 30.0 ± 0.9 38.8 ± 2.1 37.9 ± 3.8 GAA-/-:* male (g) 31.5 ± 2.2 32.2 ± 1.7 25.2 ± 1.2 Female (g) 25.2 ± 1.2 25.9 ± 1.9 22.0 ± 0.7 General Features of Gaa-/Mice Body weight was recorded as an overall measure of the health status of the Gaa-/mice (Table 2-1). At all ages examined, the Gaa-/mice weighed significantly less than their wild-type controls. No age-related gender differences were observed, although males weighed significantly more than fema les at the three age intervals studied. Glycogen Quantification and PAS Stai ning of the Cervical Spinal Cord Glycogen was elevated at all ages in the cervical spinal cords (C3-C5) of Gaa-/mice, and differences were even more pronounced at >21 months compared to 6 months in Gaa-/mice (Figure 2-1). This was also obs erved in an independent series of experiments, in which the spinal segments were not restricted to the cervical level (Figure 2-2). Consistent with these re sults, correlative histochemi stry using the PAS method demonstrated significant reaction product in neuronal cell bodies th roughout the cervical spinal cord gray matter which was most obvi ous in motoneurons (Figure 2-3A). To relate this observation to phrenic motoneurons, we used an indirect approach in which we

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32 applied the retrograde tracer, fluorogold, to the peritoneal surface of the diaphragm. This approach enabled the identification of a sm all cluster of neurons (Figure 2-3B) which were localized in a region of the ventral gray matter similar to where phrenic motoneurons have been described in the sp inal cord (Goshgarian and Rafols; Prakash, Mantilla et al. 2000). Visualization of cells in the same area in adjacent PAS-stained sections demonstrated large neurons with prominent PAS-positive droplets throughout the cell body cytoplasm (Figure 2-3C). Comp arable neurons from PAS-stained sections of control specimens showed neurons with virtually no PAS-positive inclusions (Figure 2-3D). Figure 2-1. Glycogen quantifica tion for unspecified spinal segments. *=Gaa-/different from control, #=24 month different fr om 6 month, glycogen measured by µg glycogen/mg wet weight. n=4/group.

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33 Figure 2-2. Glycogen quantification for cer vical spinal segments C3-C5. *=p<0.01: different from control. N=6/group. Figure 3A ug glycogen/mgww 0 10 20 30 40 control Gaa -/* * *6 12 >21 months

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34 Figure 2-3. Histological st aining for glycogen in phr enic motoneurons. 2-3A 2-3B 2-3C 2-3D

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35 Figure 2-4. Minute Ventilation : Expi red CO2 ratio for control and Gaa-/mice at 6 (A), 12 (B) and >21 (C) months of age. *= Gaa-/different from control. Ve/VCO2 5 10 15 20 control male control female GAA-/male GAA-/female * 1C. 1B. Ve/VCO2 5 10 15 20 * 1A. Ve/VCO2 5 10 15 20 * 2-4A 2-4B 2-4C

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36 Ventilation Respiratory measurements were made in awake, unrestrained mice using the technique of barometric plethysmography (F uller, Golder et al. 2006). This method provides a reliable and accurate measure of re lative differences in ventilatory function across time and between experiment al groups (DeLorme and Moss 2002). Gaa-/mice appeared to be hypoventilating based on the minute ventilation/expired CO2 ratio, which normalizes minute ventilation to metabolic CO2 production. This meas ure was attenuated at baseline in Gaa-/mice vs. wild-type controls (Figure 2-4). Baseline minute ventilation, breathing frequency, tidal volume, peak inspiratory flow, peak expiratory flow and tidal volume/inspiratory ti me ratio were also decreased in Gaa-/mice compared to controls at all ages studied (Table 2-2, Fi gure 2-4). The only age differences detected were lower frequency at >21 months (vs. 6 months) and elevated tidal volume at >21 months (vs. 6 months). No strains by age interactions were detected in our analyses. Hypercapnic challenge was used as a respir atory stimulus to test the capacity to increase ventilation in Gaa-/mice. The 10 minute response to hypercapnia was lower for Gaa-/mice vs. controls at each age for minut e ventilation (Figure 2-4), as well as frequency, tidal volume, peak inspiratory fl ow, peak expiratory flow and the tidal volume/inspiratory time ratio. Interestingly, gender differences were detected only in the 6 month age group, whereby females had a di fferent response to hypercapnia for all respiratory variables tested (Table 2-3).

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37 Figure 2-5. Frequency (5A, 5D, 5G), tidal volume (5B, 5E, 5H) and minute ventilation (5C, 5F, 5I) in 6 (left panel), 12 (middl e panel) and >21 (right panel) month control and Gaa-/mice at baseline and 10 minutes of hypercapnia. *=Gaa-/different from control. †=male diffe rent from female. Control 6 months: n=10/group. Gaa-/6 months: n=8 males, n=10 females. 12 months: n=8/group for all groups. Control >21 months: n=5 males, n=6 females. Gaa/>21 months: n=7 males, n=8 females. 2B. 0.0 0.1 0.2 0.3 0.4 0.5 Tidal Volume (mL)}* 2E. 0.0 0.1 0.2 0.3 0.4 0.5 }* 2C. Minutes Minute Ventilation (mL/min) 0 50 100 150 200 }* 2F. Minutes 0 50 100 150 200 B6/129 male B6/129 female GAA male GAA female }* 2H. 0.0 0.1 0.2 0.3 0.4 0.5 }* 2I. Minutes 0 50 100 150 200 }* 2A. Frequency (breaths/min) 75 125 175 225 275 325 375 }* 6 MONTH 2D. 75 125 175 225 275 325 375 }* *12 MONTH 2G. 75 125 175 225 275 325 375 }*>21 MONTH 5A. 5B. 5C. 5D. 5E. 5F. 5G. 5H. 5I.

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38 Table 2-2. Baseline ventila tion characteristics. *=p<0.01: different from control. €=p<0.01: >21 months diffe rent from 6 months. FREQUENCY (BREATHS/ MIN) TV (ML/ BREATH) MV (ML/ MIN) PIF (ML/ SEC) PEF (ML/ SEC) TV/TI (ML/ SEC) 6 MO. Control 239 ± 7 0.27 ± 0.00 64.8 ± 3.7 5.9 ± 0.2 3.4 ± 0.2 3.4 ± 0.2 Gaa-/197 ± 6* 0.21 ± 0.00* 41.6 ± 2.3* 3.3 ± 0.2* 2.2 ± 0.1* 1.7 ± 0.1* 12 MO. Control 252 ± 7 0.31 ± 0.00 77.3 ± 3.4 6.7 ± 0.2 4.4 ± 0.2 3.9 ± 0.2 Gaa-/186 ± 7* 0.23 ± 0.00* 43.2 ± 2.3* 3.6 ± 0.1* 2.3 ± 0.1* 2.1 ± 0.1* >21 MO. Control 225 ± 7€ 0.33 ± 0.00€ 73.4 ± 3.2 6.3 ± 0.3 4.6 ± 0.2 3.7 ± 0.2 Gaa-/168 ± 7*€ 0.25 ± 0.01*€ 41.8 ± 3.8* 3.5 ± 0.3* 2.3 ± 0.2* 2.1 ± 0.2* Table 2-3. Six month mean response to hype rcapnia. Ten minute mean response to hypercapnia (7% CO2, balanced O2). * = Gaa-/different from control. = male different from female for each variable presented. FREQUENC Y (BREATHS/ MIN) TV (ML/ BREATH ) MV (ML/MIN ) PIF (ML/SEC ) PEF (ML/SEC ) TV/TI (ML/SEC ) Contro l Male (n=10) 324 ± 5 0.40 ± 0.01 134.9 ± 4.2 8.3 ± 0.2 7.2 ± 0.3 5.2 ± 0.1 Female (n=10) 321 ± 3 0.40 ± 0.01 130.8 ± 3.1 7.8 ± 0.3 6.8 ± 0.2 5.8 ± 0.1 Gaa-/Male (n=8) 249 ± 4* 0.33 ± 0.01* 83.3 ± 2.5* 4.8 ± 0.1* 4.8 ± 0.2* 3.2 ± 0.1* Female (n=10) 291 ± 5* 0.33 ± 0.01* 98.2 ± 2.9* 5.6 ± 0.1* 5.3 ± 0.2* 3.7 ± 0.1*

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39 Table 2-4. Body weight as a possible covariate to respiratory volumes. TIDAL VOLUME MINUTE VENTILATION TIDAL VOLUME/ INSPIRATORY TIME MINUTE VENTILATION/ VCO2 BASELINE CV NS NS NS HYPERCAPNIA 6 MONTH CV CV CV NA 12 MONTH NS NS NS NA >21 MONTH NS NS NS NA CV=bodyweight is a co variate (p<0.01) NS=bodyweight is not significant NA=not applicable for this variable Hemoglobin and Hematocrit Hypoxia results in elevated hemoglobin (Hb) and hematocrit (Hct) levels (Hartmann, Krafft et al. 2005). Because our data suggest signifi cant hypoventilation, Hb and Hct were measured in 12 month control and Gaa-/mice (Table 2-5). Both Hb and Hct were elevated in Gaa-/mice, most likely due to insuffi cient arterial partial pressure of O2. In addition, glucose and sodium leve ls did not vary between control and Gaa-/mice, suggesting that Hb and Hct differences did not reflect plasma volume differences. Table 2-5. Hemoglobin, hematocrit, sodium and glucose levels for control and Gaa-/mice at 12 months. *=p<0.01: different from control. n=9/GROUP HEMOGLOBIN (G/DL) HEMATOCRIT (%)SODIUM (MMOL/L) GLUCOSE (MG/DL) Control 13.5 ± 0.3 39.8 ± 0.9 144.5 ± 0.8 180.4 ± 16.3 Gaa-/15.3 ± 0.4* 45.0 ± 1.1* 143.4 ± 0.8 176.8 ± 11.4 Conscious Arterial Blood Sampling For verification of hypoxemia in Gaa-/mice, conscious arterial blood samples were collected in 12 month Gaa-/and control mice for measurement of partial pressures of O2.

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40 In agreement with the hypothesis that Gaa-/mice are hypoventilating, Gaa-/mice had lower PO2 vs. controls (Table 2-6). Table 2-6. Conscious arterial bl ood sampling to measure arterial partial pressure of O2 in control and Gaa-/mice at 12 months. *=p<0.01 : different from control. N=6/GROUP PARTIAL PRESSURE OF O2 (MMHG) Control 98.5 1.9 Gaa-/83.3 2.7* Table 2-7. Bodyweights of muscle specific Ga a mice used in ventilation experiments. BODYWEIGHT (G) 40.6 2.5 Muscle-Specific hGAA Mice We next wanted to quantify respiratory f unction in transgenic animals with musclespecific correction of GAA activity (MTP mice). To first obtain an index of diaphragm muscle function, we measured in vitro contractility of diaphr agms from control, Gaa-/and MTP mice. Control and MTP mice had similar forces produced by comparable stimulation frequencies, while the Gaa-/produced a significantly smaller force for each frequency (Figure 2-6-A). These data confir m that the normal glycogen levels in MTP diaphragm muscle (MTP vs. control; 1.7 ± 1.3 vs. 1.4 ± 0.2 µg/mgww) correspond to diaphragm muscle that is functi onally similar to controls. Despite apparently normal functional diaphr agm muscle (Figure 2-6-A), the pattern of breathing was altered in the MTP mice. Minute ventilation during baseline was similar in MTP and Gaa-/mice, and both were significantly reduced compared to controls (Figure 2-6-B). Furthermore, th e response to hypercapnia was attenuated in MTP mice (p<0.01), although these mice s howed a greater response than the Gaa-/mice

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41 2-6-A 2-6-B m i nu t e ven til a ti on ( m L/ m i n ) 0 40 80 120 160 control muscle specific Gaa Gaa -/baselinehypercapnia • • •* Figure 4BFigure 4A Frequency (Hz) Tw10204080100150200 Force (N/cm 2 ) 0 5 10 15 20 25 30 control Gaa -/muscle specific hGaa †

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42 Figure 2-6. Diaphragm contra ctile function (n=3/group) and minute ventilation for control (n=8), Gaa-/(n=8) and MTP (n=6) mice. Airflow tracings from representative control (Figure 2-6C:baseline, Figure 2-6-D:hypercapnia), Gaa-/ (Figure 2-6-E:baseline, Figure 26-F:hypercapnia) and MTP (Figure 26-G:baseline, Figure 26-H:hypercapnia) mice. ______ = 1 sec. *=p<0.01 different from control. •=p<0.01 all groups different from each other. Table 2-8. Phrenic neurophysiology characteristics for control, Gaa-/and MTP mice. *=p<0.01, different from control. RATE OF RISE (MV/S) FREQUENCY (BREATHS/S) AMPLITUDE (MV) Control (n=9) 346 ± 86 167 ± 14 52.8 ± 14.1 Gaa-/-(n=9) 44 ± 15* 107 ± 14* 6.6 ± 1.7* MTP (n=6) 101 ± 27* 124 ± 17* 11.8 ± 1.8* (p<0.01; Figure 2-6-B). Airf low tracings are depicted for baseline and hypercapnia in a representative contro l (Figure 2-6-C-D), Gaa-/(Figure 2-6-E-F) and MTP (Figure 2-6-GH) mouse demonstrating the robus t difference between mice.

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43 Efferent Phrenic Activity To determine whether the comp romised ventilation seen in Gaa-/and MTP was associated with reduced phrenic motor out put, we measured efferent phrenic nerve activity in Gaa-/-, MTP and control mice at 12 months of age. At similar arterial PCO2 levels (see legend, Figure 2-7-A), Gaa-/and MTP mice had signi ficantly lower phrenic inspiratory burst amplitudes (Figure 2-7-A). The neurogram recordings from Gaa-/and MTP mice also revealed less frequent bursts, and an attenuated slope of the integrated inspiratory burst (i.e. slower “rate of rise”, Table 2-8). Parenthetically, it should be noted that only the more robust of the Gaa-/mice studied were able to tolerate the neurogram recording protocol. For this reason, the true mean phrenic amplitude shown in Figure 27-A may actually be even lower across all Gaa-/mice. A representative neurogram for a control (Figure 2-7-B), Gaa-/(Figure 2-7-C) and a MTP (F igure 2-7-D) mouse are shown in Figure 2-7. Discussion This study of a murine model of Pomp e disease has reveal ed several novel observations pertaining to GAAdeficiency and concomitant respiratory involvement. First, ventilation is reduced in Gaa-/mice as revealed by qua ntitative barometric plethysmography. Second, cervical spinal cord glycogen is elevated in Gaa-/mice and PAS staining demonstrated prominent glyc ogen inclusions in cervical motoneurons, including phrenic motoneurons indirectly id entified by retrograde fluorogold tracing. Third, Gaa-/mice have greatly attenuated phrenic output relative to wild-type controls of

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44 Figure 2-7. Thirty second mean phrenic inspiratory burst amplitude for control, Gaa-/and muscle specific hGaa mice with similar arterial PaCO2 values (2-8-A). *=different from control. Raw phrenic amplitude (top trace) and rectified, integrated trace (bottom trac e) from a representative control (Figure 2-8-B), Gaa-/(Figure 2-8-C) and MTP (Figure 2-8D) mouse. _________=1 sec. comparable age. Lastly, MTP mice also exhibited breathing impairments and phrenic neurogram features simila r to those revealed in Gaa-/mice, despite apparently normal diaphragmatic contractile function. To our know ledge, these are the first formal lines of evidence suggesting that resp iratory weakening in the Gaa-/mouse, and by extrapolation in Pompe disease patients, may represent a combination of both neural and muscular deficits. The Gaa-/Mouse Model and Altered Ventilation Our first major finding demonstrates that Gaa-/mice have a ventilatory phenotype similar to juvenile and adult onset Pompe di sease patients. It is unclear, however, why ventilation for Gaa-/mice did not get progressively worse as the mice aged. 2-7-A burst amplitude (mV) 0 20 40 60 control PaCO2 = 53.7 ± 2.0 mmHg Gaa-/-PaCO2 = 53.7 ± 1.6 mmHg MTP PaCO2 = 51.8 ± 1.3 mmHg * * 2-7-B 2-7-C 2-7-D .0 .0 .0 .1 .1.1 .25 -.25 -.25 .25 -.25 .25 mV mV mV

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45 Nevertheless, the Gaa-/mouse model used in this inve stigation is presently the most compelling available model of this disorder . Previous studies have shown glycogen accumulation in all tissues of the Gaa-/mouse examined thus far (Raben, Nagaraju et al. 1998), and other physiological measures appear st rikingly similar to what is seen in the patient population (Hirschhorn 1995). Cardiac pathology in these mice is not as severe as in infants with Pompe disease (Seifert, S nyder et al. 1992), but instead more closely corresponds to juvenile and adult patients (Hirschhorn 1995). It should be recognized that the Gaa-/mouse line in this study is maintained on a genetically diverse background, as it was de veloped by crossing the B6 and 129 mouse strains (Raben, Nagaraju et al. 1998). While it is possibl e that this diverse genetic background could result in modifier genes w ith different compensatory mechanisms depending on the genetic proportion of each strain resulting in the Gaa-/mice, importantly, B6 and 129 inbred mice have si milar minute ventilation values at baseline and in response to hypercapnia (Tankersley, F itzgerald et al. 1994). In addition, we have maintained this line for over five years with brother/sister mating to assume as homogeneous a background as possible. To circumvent any issues involving this background strain, we tested large numbers of mice. In addition, we followed the guidelines set forth by the Banbury Confer ence for studying genetically altered mice (Silva 1997). To obtain a comprehensive and complementar y analysis of resp iration in control and Gaa-/mice, we combined barometric plethysmography with phrenic neurophysiology breathing performance. Baro metric plethysmography is a useful tool for studying the ventilatory behavior in conscious mice, although some technical

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46 differences of opinion have been presented in the literature (Enhorni ng, van Schaik et al. 1998; Mortola and Frappell 1998). The ventilat ory responses in mice, however, are quite reproducible (DeLorme and Moss 2002). Although body mass differed between control and Gaa-/mice, we did not normalize minute ventilation to body mass as no differences in lung volume were anticipated. However, lung volumes were not determined, so some interpretative caution is thus warranted. To more accurately rem ove the influence of body mass on ventilation, the analysis of covariance was used to analyze the data, and minute ventilation was normalized to expired CO2 for a more appropriate normali zation of ventilation between groups. Characterization of the ventilation of Gaa-/mice will be useful for future studies that measure the efficacy of various therapies. An important point for future studies is that male and female Gaa-/mice differ in their response to hypercapnia at 6 months of age. Therefore, these data should be cons idered when performing experiments utilizing potential treatments for Gaa-/mice (in this age group). Although no gender differences have been described in Pompe disease, ther e may be simply too few patients who have been thoroughly evaluated. Evidence for a Neural Contribution to Respiratory Deficits in Gaa-/Mice Since multiple tissues are affected by GAA deficiency (10), it was predicted that glycogen should also be accumulated in CNS, and relevant to our respiratory data, especially in the region of the spinal co rd where the phrenic cell bodies reside (C3-C5). Excess glycogen within the sp inal cord (including phren ic motoneurons) led us to quantify inspiratory phrenic burst amplitude between control and Gaa-/mice. Wellestablished electrophysiologica l techniques were utilized which require a reduced

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47 preparation in which phrenic motoneuron output can be directly measured (Baker and Mitchell 2000; Janssen and Fregosi 2000). Although normalizations to maximum (during gasping) and/or minimum (the phrenic bur sting that emerges following apnea) are commonly performed in various neurophysiologi cal manipulations in rats (Baker and Mitchell 2000; Janssen and Fregosi 2000), it was not suitable to normalize across the groups of healthy and genetically altered mice. Normalizations of this sort are intended for studying healthy, normal animals. It shoul d be noted that the variability was higher for control animals vs. Gaa-/(Figure 5), suggesting that healthy animals do have more divergence in their phrenic burst amplitude (and normalizations in these mice may have reduced the variability). N onetheless, both the phrenic amplitude associated with the onset of bursting following apnea (B6/129: 31.9 0.3 mV, Gaa-/-: 3.7 0.9 mV) and hypercapnic responses (B6/129: 68.7 20.0 mV, Gaa-/-: 14.0 4.8 mV) were measured between the two groups demonstrating that the range of phrenic burst amplitude was attenuated in the Gaa-/mice. Overall, the pairing of conscious barometric plethysmography measurements with the redu ced neurophysiological pr eparation offers a well-rounded assessment for the control of ventilation. The Gaa-/mice had severely blunted phrenic burst amplitude compared to controls during standard conditions. These data provided direct eviden ce of a neural deficit in Gaa-/mice. Importantly, we only compared the inspiratory phrenic burs t amplitude of mice with similar PaCO2 levels. In addition to excess glycogen within the spinal cord, the neural pathol ogies could also be the result of lack of GAA ac tivity throughout development. The role of GAA activity in the developing nervous system is currently unknown, although GAA expression is highest in neural tissue compared to all other ti ssues during development (Ponce, Witte et al.

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48 1999), and regulators of GAA are known to be necessary for neural development (Ohtsuka, Ishibashi et al. 1999). In conclusion, we measured the phrenic nerve activity which is the final motor output of the respiratory system. Despite mark ed differences in the phrenic nerve activity between control and Gaa-/mice, the differences could be resulting from higher (neural) respiratory inputs and/or impairment of ch emosensory afferents due to the hypothesized chronically elevated PaCO2 and attenuated PaO2 levels. It should be noted that during conditions of higher respiratory drive (B6/129 PaCO2: 87.2 5.3 mmHg, Gaa-/PaCO2: 92.3 7.4 mmHg) both groups were able to incr ease phrenic inspiratory burst amplitude, but the Gaa-/mice continued to have lo wer output (B6/129: 68.7 mv 20.0, Gaa-/-: 14.0 mv 4.8). Importantly, the final end product of the phrenic nerve activ ity was altered in Gaa-/mice, demonstrating a neural defi cit of respiratory control. Contribution of Diaphragm Musc le in Ventilation Deficits To determine that muscular dysfunction is not the only contributor to ventilation deficits due to GAA deficiency, we utilized a double transgenic mouse that expresses hGAA only in skeletal muscle (maintained on the Gaa-/background). Since these mice have normal muscle function (Fig 4A), we hypothesized that any differences in ventilation between MTP and control mice would reflect differences in the neural control of the respiratory muscles. Consistent w ith this suggestion, ventilation was similar between MTP and Gaa-/mice during quiet breathing (both le ss (p<0.01) than controls). When respiratory drive was elevated with hypercapnia, the ventilatory response of MTP mice was still less than the control strain, but elevated compared to Gaa-/mice (Figure 4B). Thus, both muscle and neural component s contribute to vent ilation deficits under

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49 conditions of elevated respir atory drive. The increased ventilation in response to hypercapnia would have induced use of the intercostal and abdominal muscles, which should be functionally normal in MTP mice (but not in Gaa-/mice). These functional intercostal and abdominal muscles may have be en a factor in the improved ventilation of MTP vs. Gaa-/mice during hypercapnia. In add ition, MTP mice would have been less likely to have skeletal deformities of the ri b cage which may have contributed to this ventilatory response. Since the MTP mice were derived by crossing Gaa-/mice with the FVB strain, it is appropriate to note the stra in differences between the Gaa-/and MTP mice. FVB mice are genetically distinct from the B6 and 129 strains that were used to develop the Gaa-/mouse. However, locomotor strength is similar for C57BL/6 and FVB mice (Hesselink, Gorselink et al. 2002). Nonetheless, ve ntilation data comparing MTP mice to Gaa-/and controls demonstrates that diaphragmatic muscle dysfunction alone could not account for breathing deficits resulti ng from GAA deficiency. Therapeutic Implications Further evaluation to identify th e morphology and cell number of the Gaa-/phrenic nucleus in mice and phrenic elect rophysiology in humans are impor tant next steps. It is hypothesized that glycogen accumulation in the motoneuron cell body may result in larger motoneurons. According to the He nneman size principle, larger motoneurons require greater excitatory synaptic input for depolarization to occur than smaller motoneurons (Henneman, Somjen et al. 1965). Thus, it is possible that the motoneurons themselves are functional, but the probability they will be depolarized is lower in Pompe disease individuals. Potentia l therapies which will target both skeletal muscle and motoneurons should be the ultimate goal for Pompe disease. Interventions for Pompe

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50 disease can be assessed by comparing treated Gaa-/mice to values reported in this study. Importantly, the current available therapy for Pompe disease, enzyme replacement, cannot be used to treat GAA deficiency of the central nervous system because enzymes do not cross the blood-brain barrier (Kikuc hi, Yang et al. 1998; Raben, Danon et al. 2003). The results of enzyme replacement ther apy have varied across the Pompe disease patient population (Van den Hout, Kamphoven et al. 2004), and these findings provide some explanations for current human studies. Data from these experiments have reinforced that the Gaa-/mouse is a useful model to study mechanisms of respiratory distress caused by GAA deficiency. Ventilation is lower for Gaa-/mice compared to controls , glycogen is elevated in Gaa-/spinal cord, and phrenic inspiratory burst amplitude is reduced in Gaa-/mice. Findings from this study supports the hypothesis that ne ural deficits contri bute to respiratory insufficiency in Pompe disease, and these conclusions may lead to better treatment possibilities, especially for j uvenile and adult-onset patients.

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51 CHAPTER 3 CHARACTERIZATION OF THE INTRAT HORACIC INJECTION USING ADENOASSOCIATED VIRUS SEROTYPE 1 Background Diaphragmatic muscle weakness is a primar y symptom in Pompe disease that leads to respiratory insufficiency in adult-on set patients. The muscle weakness in Gaa-/mice is due to both contractile dysfunction and diminished phrenic nerve activation of the muscle, suggesting that adultonset Pompe disease patients have a similar mechanism of weakness (Chapter 2). Currently there is no cure for Pompe disease, although both gene and enzyme replacement therapies have emerge d as possibilities for treatment of this devastating disease (van der Ploeg, Bolhui s et al. 1988; Cresawn, Fraites et al. 2005; Mah, Cresawn et al. 2005). Enzyme replacemen t therapy is not suitable to treat the central nervous system deficits in Pompe di sease, as it does not pass through the blood brain barrier (Raben, Danon et al. 2003). Fortunately, gene therapy has demonstrated promise as a therapy for neuromuscular diseases (Kaspar, Llado et al . 2003; Xu 2005). The concept of gene therapy is based on a one time injection to deliver DNA to cells resul ting in transcription of the delivered gene. Specifically, our group has chosen adeno-associated viral vect ors due to their safety and long-term expression of the packaged tran sgene (Fisher, Joos s et al. 1997). AAV carrying the acid-alpha glucosidase gene has reduced glycogen accumulation in mouse models of Pompe disease (GAA-/-; exon 6) (Fraites, Schleissing et al. 2002; Rucker, Fraites et al. 2004; S un, Zhang et al. 2005).

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52 The intrathoracic injection was developed as a relatively non-invasive technique to deliver AAV (with a transgene) to the diaphr agm; with the long-te rm goal of treating respiratory deficits in Gaa-/mice. Importantly, AAV2 is transported to motoneurons following intramuscular injection (Kaspar, Llado et al. 2003). For this reason, the intrathoracic injection was hypothesized to target both the diaphragm and phrenic motoneurons. Materials and Methods: Animals All mice used in these studies were B6/129 mi ce bred at the University of Florida. Animals were housed at the University of Fl orida specific pathogen-free animal facility and procedures were done in accordance with the Institutional Animal Care and Use Committee approved guidelines. Virus Preparation Recombinant AAV vectors was packaged by the University of Florida Powell Gene Therapy Vector Core as prev iously described (Zolotukhin, Po tter et al. 2002). Purified recombinant AAV stocks were characterized by SDS/polyacrylamide gel electrophoreses with silver-stain and particle count. Total particle titer was determined by the dot blot method as previously described (Z olotukhin, Byrne et al. 1999). Survival Surgery for Intrapleural Injection Animals were anesthetized with 2% isoflu rane and restrained in the supine position on a warmed operating surface. The surgical area which includes a 1 cm radius around the sternum was shaved and clipped; then scrubbed three times with Povidone-iodine scrub alternating with sterile saline. The xiphoid process was elevated with forceps. At such time, the ribs were palpated with a sterile cotton swab to detect the 4th and 5th ribs.

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53 An injection was made into the intrathoracic space between ribs 4 and 5, just where the ribs meet the sternum with a 29-gauge in sulin syringe. The needle had PE-10 tubing around the needle, allowing only 4 mm of the n eedle to penetrate into the body cavity, to prevent puncturing the heart or lungs. 100µ L of AAV-CMV-LacZ diluted in Lactated Ringer’s Saline (Baxter Healthcare Corporat ion, Deerfield, IL) was injected into the intrathoracic space. Mice were then held upright for 10 minutes to increase the probability of AAV infecting the diaphragm. They were then removed from isoflurane and allowed to recover in their home cages. Tissue Harvesting 4 weeks after the AAV-CMV-LacZ intraple ural injection, mice were euthanized with an overdose of pentibarbitol (150 mg/kg). The cervical roots C3-C5 were identified and the corresponding spinal cord was isolated and removed from the spinal column. A segment from the most rostral portion of C3-C5 will (~25 mg) was immediately frozen in liquid nitrogen and used for DNA isolation. The middle segment (~3 mm) was frozen in tissue freezing medium to be cut (20 µm s ections) and immunohistochemistry for LacZ performed. The most caudal segment was plac ed immediately in a FastPrep tube (MP Biomedicals; Irvine, CA) containing 100 µL of water and sterile homogenizing beads for the LacZ protein activity assa y. The diaphragm was also ha rvested and stored following the protocol listed above for the same assays. Immunostaining for LacZ Coronal sections of cervical spinal cord were cut at 20 µm each. The diaphragm and rib cage were harvested from 4% paraformaldehyde cardiac perfused mice. The tissue (cord sections or whole diaphragm) was then fixed for 5 minutes with 2% formaldehyde and 0.2% glutaralde hyde. Next, the tissue were washed 3 times with 1X

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54 phosphate buffered saline. Then the X-gal staining solution (MP Biomedicals; Irvine, CA) incubated on the tissue for 24 hours. DNA Isolation A section of the cervical spinal cord a nd diaphragm was used (~25 mg) to isolate DNA. The tissue was added to 500µL of D NAzol (Molecular Research Center, Inc; Cincinnati, OH) in a 1.5 mL microcentrif uge tube and homogenized by hand with a Teflon drummel. Twenty µL of proteinase K (Sigma-Aldrich; St. Louis, MO) was added to the solution containing DNAzol and ho mogenized sample, which was incubated overnight at room temperature. Next, 1.0 mL of 100% ethanol was added to each tube. The tubes were inverted 3 times and sat at room temperature for 5 minutes. The samples were then centrifuged for 5 minutes at 5000 x g to pellet the DNA. The pellet was washed 2 times with 75% ethanol; sedime nting the pellet between each wash by centrifugation at 1000 x g. Afte r the final centrifugation, the supernatant was removed and 50 µL of sterile water was added to solu blize the DNA. After the mixture had sat at room temperature for at least 30 minutes, the DNA was quantified by reading it at 260 nm. Samples were stored at -20°C. Polymerase Chain Reaction (PCR) One hundred nanograms of each DNA sample was used in the PCR reactions. PCR was carried out in 25µ L reactions with 100 pmol/microliter of the specific oligonucleotide primers and ten microliter s of 10X MasterMix (Eppendorf, Brinkmann, Westbury, NY). The primers were 5'-AGCTGGCGTAATAGC-3' and 5'TCGACGTTCAGACGTAGTCA-3' and were de signed to generate a 850 base-pair product. PCR temperature profiles were: a single cycle at 94°C for ten minutes; 30 cycles of 60 seconds 94°C (d enaturing), 60 seconds at 60° C (annealing), and 120 seconds

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55 at 72°C (extension); and a final cycle at 72°C for ten minutes. The PCR products were electrophoresed on a 1.5% agarose gel containi ng 0.4 micrograms/mL ethidium bromide. LacZ Protein Activity A section of cervical spinal cord (C3-C5; anterior to posterior section) weighing ~25 mg was harvested and immediately placed into a FastPrep (MP Biomedicals; Irvine, CA) tube filled with sterile beads and 100µ L of sterile water for homogenization. The tubes were put into the FastPrep (40 sec, level 4.0) and homogenized lysates were transferred to a 1.5 mL microcentrifuge tube and centrifuged at 20,000 x g for 5 minutes. The clarified lysate was transferred to a ne w 1.5 mL microcentrifuge tube and the pellet discarded. Clarified lysate (10 µL) was used in the Galacto-Star chemiluminescent reporter gene assay system for detection of -galactosidase activity (Tropix Inc., Bedford, MA.). Protein concentrations of lysates were determined using a standard Bradford assay based on the binding properties of detecti on of Coosmassie Brilliant Blue G-250 dye reagent (Bio-Rad, Hercules, CA). The Cooma ssie dye allows for an increasingly intense blue color to be detected at 620 nm waveleng th with increasing am ounts of total protein in the well. A standard curve was generate d using seven concentra tions of bovine serum albumin and corresponding A620 values. Va lues for the Galacto-Star assay were reported as relative light units/µg protein. Fluorescent Imaging of Mouse Diaphragm An intrathoracic injection w ith 100µL of IR Dye 800 (Li-Cor Biosciences; images green) was administered to an anesthe tized mouse (2% isofluorane, balance O2). Twenty minutes post-injection, the mouse was placed in the prone position on the Odyssey Infrared Imaging System (Li-Cor Bioscience s; Lincoln, NE) and imaged with a 680 nm

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56 and 780 nm laser. The 680 nm wavelength de tects the auto-fluores cence of the animal while the 780 nm wavelength dete cts the injected IR Dye 800. Fluorescent Labeling of Motoneurons Mice were anesthetized with 2% isoflurane and restrained in the supine position on a warmed operating surface. The surgical area was shaved and clipped; then scrubbed three times with Povido ne-iodine scrub alternating with sterile saline. A laporatomy was performed for access to the cauda l diaphragm. Four percen t fluoro-gold was painted on the diaphragm (~75 µL). Complete coverage of the diaphragm was determined when all regions of the diaphragm were yellow in colo r (from the fluoro-gold). Abdominal muscle and skin was closed with 6-0 suture. Mi ce were then removed from isoflurane and allowed to recover in their home cages. Fo rty-eight hours after fl uoro-gold application, mice were given a lethal dose of sodium pe ntabarbitol and cardiac perfused with 4% paraformaldehyde. Spinal cord was exci sed 24 hours later and placed into rising gradients of 10, 20 and 30% sucrose to cr yoprotect the tissue pr ior to sectioning. Co-labeling of AAV with Motoneurons Mice were administered the intrathoraci c injection with AAV1-CMV-GFP (1 x 1011 particles). Four weeks postinjection, fluoro-gold was pain ted onto the diaphragm of the same mice (method shown above) to identif y motoneurons. Forty-eight hours after fluoro-gold application, mice we re given a lethal dose of sodium pentabarbitol and cardiac perfused with 4% paraformaldehyde. Spinal cord was excised 24 hours later and placed into rising gradients of 10, 20 and 30% sucrose to cryoprotect the tissue prior to sectioning. The DAPI filter was used to detect fluor-gold labeling and the FITC filter to detect GFP positive cells in the 40µm sections.

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57 Statistics The one way ANOVA was used for all meas ures that were quantified. P<0.05 was considered significant. Results Intrathoracic Injection: Dilution Media Mice were injected with AAV1-CMV-LacZ (1 x 1011 particles) diluted in either lactated ringer’s saline, lactated ringer’s saline with 20% gel solution (this solution was the highest percentage of gel that could also be pulled into the insulin syringe), or lactated ringer’s solution with 10% iode xonol (contrasting agent that is denser than water). Diaphragm -Galactosidase activity was elevated for each treated group compared to controls and tended to be elevated in the ge l treated group (Figure 3-1). Spinal cord (C3C5) -Galactosidase activity was only elevated in the gel and lactated ringer’s groups vs. controls (Figure 3-2). Intrathoracic Injection: Targeting of Mouse Diaphragm To verify the distribution for the 100 µL intrathoracic injection, mice were injected with the IR Dye 800 which is detected at the 780 nm wavelength. The figure demonstrates the intrathoracic injection doe s effectively cover the diaphragm (Figure 33). In addition to fluorescent detection of the intrathoracic injection, mice were injected with AAV1-CMV-LacZ (1 x 1011 par ticles) diluted in 100 µL of Lactated ringer’s solution 4 weeks prior to euthanasia. Lactated ringer’s was the preferred dilution media because it is the safest and -galactosidase activity was detected in both diaphragm and spinal cord following the preliminary AAV1-CMV-LacZ injections.

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58 Figure 3-1. Media comparision for diaphragm -Galactosidase activity from mice injected with 1 x 1011 particles of AAV1-CMV-LacZ.

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59 Figure 3-2. Media comparision for spinal cord -Galactosidase activity from mice injected with 1 x 1011 particles of AAV1-CMV-LacZ.

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60 Figure 3-3. Intrathoracic injection with 100µL of IR Dye 800 (Li-Cor Biosciences; images green). Figure 3-4. X-Gal staining of mouse diaphr agm following intrathoracic injection with AAV1-CMV-LacZ (left panel) or phospha te buffered saline (right panel).

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61 Diaphragm was stained with the X-Gal solution to detect active LacZ in the muscle. The figure (Figure 3-4) demonstrates even distribution of positive (blue) X-Gal fibers. DNA Detection of Control Gene LacZ DNA was detected in harvested dia phragm from all mice tested that were injected with 1 x 1011 particles of AAV1-CMV-LacZ via intrathoracic inje ction (Figure 3-5). The same region of diaphragm was harvested in each mouse. Figure 3-5. PCR product run on a 1.5 % agar ose gel from diaphragm DNA amplified with primers specific for the LacZ gene. DNA was isolated from cervical spinal cord (C3-C5) of control mice and mice injected with AAV1-CMV-LacZ . Primers specific for LacZ and its promoter, CMV were used to amplify the specific DNA to identify if AAV1-CMV-LacZ was present in this region of the spinal cord which contains the phrenic nucleus. If retrograde transport of AAV1 were to occur (from the diaphrag m) with this injection, the DNA would be present in the phrenic motoneurons. Importa ntly, this technique is testing for positive DNA is all cell types within the spinal co rd and is not specif ic for motoneurons. ladder neg control DIAPHRAGM 10 pg lacZ 0.1 pg

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62 Figure 3-6. PCR product run on a 1.5 % agaros e gel from isolated spinal cord (C3-C5) DNA amplified with primers specific for LacZ. Protein Activity of Control Gene Protein activity for the LacZ gene ( -galactosidase) was qua ntified in diaphragm from mice injected with AAV1-CMV-LacZ (1 x 1011 particles) or phosphate buffered saline (controls). Protein ac tivity was elevated in mice injected with AAV1-CMV-LacZ vs. sham injected controls (Figure 3-7). Spinal cord (C3-C5) was also harvested for quantification of -galactosidase activity. Protein activity was higher in mi ce injected with AAV1-CMV-LacZ (1 x 1011 particles), but the amplitude of elevati on (as measured by the % of control) was attenuated compared to diaphragm (Figure 3-8). Labeling of Motoneurons Positive cells were detected containing fluoro-gold in the ventral spinal cord (C3C5). In addition to fluoro-gold, the cells we re morphologically identi fied as motoneurons (Figure 3-9). Green fluorescent protein positiv e cells were not detected in motoneurons (Figure 3-10). ladder neg control SPINAL CORD 10 pg lacZ 0.1 pg

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63 ng beta gal/10 ug protein 0 2 4 6 8 10 control (PBS injected) AAV1-CMV-LacZ injected * Figure 3-7. -galactosidase activity of diaphragm muscle from mice injected with AAV1-CMV-LacZ or phosphate buffered saline.

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64 ng beta gal/10 ug protein 0.0 0.2 0.4 0.6 0.8 1.0 control: 0.73 AAV1-CMV-LacZ: 0.95 * Figure 3-8. -galactosidase activity of spinal cord (C3-C5) from mice injected with AAV1-CMV-LacZ or phosphate buffered saline.

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65 Figure 3-9. Fluoro-gold id entified motoneurons.

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66 Figure 3-10. FITC filter used for identification of GFP positive cells. No cells detected. Discussion Two major findings emerge from these AAV1 experiments: 1) the intrathoracic injection (titer: 1 x 1011 particles) results in transfec tion of mouse diaphragm and as a result, control protein is translated in this tissue and 2) at this titer, the intrathoracic injection results in detectable transgene in th e cervical spinal cord, but is undetectable in any neural cells via histological methods. Th ese results provide a rationale for utilizing the intrathoracic injection for studies aime d at diaphragmic transfection of various transgenes including, but not limited to aci d-alpha glucosidase, dystrophin, and IGF. Although results from these studies have iden tified that an intrathoracic injection of AAV1-CMV-LacZ (titer; 1 x 1011 particles) does not result in high levels of transgene activity in the spinal cord, it is unknown if a higher titer, different purification process of

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67 AAV1 or larger volume of injection would ha ve augmented the ability of AAV1 to be transported to motoneurons. Multiple repor ts have revealed similar results to our findings with intramuscular in jection (Glatzel, Flechsig et al. 2000; Martinov, Sefland et al. 2002; Wang, Muramatsu et al. 2002), althoug h these studies were not investigating phrenic motoneurons. It may be necessary in the future to administer a direct spinal injection of AAV-CMVhGaa to target the phrenic motor nuc leus for recovery of neural deficits in Gaa-/mice. Although not as applicable to the clinic (vs. intrathoracic injection), these experiments would differen tiate neural vs. muscular recovery of ventilation. Recently, a new clone of the AAV2 capsid was developed, which resulted in high transgene expression in mot oneurons following a muscular injection (Xu 2005). The University of Florida has the capabilities to develop similar capsid mutations, which may someday be used in Gaa-/mouse studies with the goal of complete recovery of ventilation deficits (muscula r and neural) via intrathoracic injection. Although these new AAV capsids with high tropism for motoneur ons offer an exciting possibility for neuromuscular disease treatment, the safe ty of these capsids will need to be characterized.

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68 CHAPTER 4 METHOD OF BAROMETRIC PLETHYSMOGRAPHY What is the Principle of Barometric Plethysmography? The original published observations us ing barometric plethysmography were performed by Chapin in 1954 measuring ve ntilation in hamsters (Chapin 1954) and Drorbaugh and Fenn in 1955 measuring ventil ation in premature infants and cats (Drorbaugh and Fenn 1955). In its most basi c sense, barometric plethysmography is measuring an increase in pressure during in spiration that is proportional to changes in tidal volume. A pressure difference is detect ed because air is humidified and warmed to body temperature upon inspiration which creates an increase in the pressure of the entire chamber. The tidal volume measured by barometric plethysmography is relatively comparable to that measured with pneumotachography (Drorbaugh and Fenn 1955; Malan 1973; Wong and Alarie 1982). However, there are multiple sources of error that can alter the tidal volume calculation (Epste in and Epstein 1978; Chaui-Berlinck and Bicudo 1998; Mortola and Frappell 1998). In most instances th ese errors are similar for different animals placed into the chamber, such that differences in the pattern of breathing in two mice measured in the sa me chamber can be considered “real” differences. The frequency of breathing meas ured with barometric plethysmography is rarely influenced by sources of error, so can be compared with confidence between animals tested (Chaui-Berlinck and Bicudo 1998; Enhorning, van Schaik et al. 1998).

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69 Figure 4-1. Theoretical ba sis for plethysmography. The animal is at 100% relative humidity (c omplete saturation). If the chamber is also at 100% relative humidity, then volum e will not change during breathing and no pressure changes will be measured (left panel) . If relative humidity is different between the chamber and the animal, the air brought into the lungs will be warmed and saturated with water which will increase the volume in th e thoracic cavity resulting in a measurable change in pressure. BoyleÂ’s Law is used stating that for a closed container at constant temperature, pressure times volume is constant. Initial chamber pressure times initial chamber volume must equal the final chamber pressure times the final chamber volume. P1 x V1 = P2 x V2; to solve for V2, V2 =V1(P1/P2) Mortola and Frappell, 1998

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70 The differences in water vapor between the chamber and the lungs must be accounted for to measure tidal volume. Here airway volume is equivalent to V2 (from above): (1) airway volume = chamber volume [air way temperature (barometric pressure – partial pressure of water in the chamber)]/c hamber temperature (barometric pressure – partial pressure of water in the airways)] The pressure signal needs to be multiplied by a correction factor because for the same tidal volume, as the difference between the lung and chamber conditions converges, the lower the pressure signa l. To account for this: (2) correction factor = airway volum e/(airway volume – chamber volume) The pressure signal is also influenced by the size of the cham ber; the larger the chamber, the smaller the pressure signal. To account for this, a factor (known here as K) is determined by performing a calibration pr ior to measurements. A known volume is injected into the chamber and the change in pressure for the known change is volume is calculated. (3) K = known change in volume/change in pressure Tidal volume can then be calculated: (4) tidal volume = pressure signal x correction factor x K Body temperature values are substituted for airway values: (5) tidal volume = pressure signal x K x [body temperature(barometric pressure – partial pressure of water in the chamber)]/[[body temperat ure(barometric pressure – partial pressure of water in the chamber)] – [chamber temper ature(barometric pressure – partial pressure of water in the body]]

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71 Sources of error for barometr ic plethysmography include: Thermal and Pressure Drifts Thermal and pressure drifts can be si gnificant (Chaui-Berlinck and Bicudo 1998; Enhorning, van Schaik et al. 1998 ). However, in our experi ments we use a flow through system which reduces the amount of drift (Wong and Alarie 1982). In addition, most chambers eventually equilibrate, but the ti me preceding the equilibration can result in erroneous tidal volume calcula tions. A figure is included to demonstrate how the time for expiration (B) is erroneous ly shortened by a thermal drift compared to the time for inspiration (A). Figure 4-2. Thermal drift. Inaccurate Measurements of Body or Chamber Temperature Differences in temperature account for a la rge portion of the calculation used for tidal volume (Malan 1973; Enhorning, van Scha ik et al. 1998). Therefore, inaccurate temperatures will increase the variability of the tidal volume measurement. During Mortola and Frappell, 1998

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72 average conditions of 24°C (chamber) and 37° C (body), inaccurate temperature readings of 1°C would result in approximately a 7% over or under estimation of tidal volume. A larger difference in the chamber and body temp erature would result in a lower percent error of tidal volume. In our conditions the chamber is typically 22°C and the body temperature is 37°C, so our conditions would create a decreased pe rcent error of tidal volume if an inaccurate body temperature we re measured (Chaui-Berlinck and Bicudo 1998). The Assumption that Warmed, Satura ted Air from the Airways Reaches Equilibrium Immediately Once It Returns to the Chamber Air from the lungs does not immediately retu rn to equilibrium (Peslin, Duvivier et al. 1995), although the calculations are base d on an immediate return to chamber conditions. Sensors can be used to measure the temperature and vapor pressure at the nostrils during expiration, but th is can be very distracti ng to the animal. Without accounting for the difference for time to equilib rium, tidal volume can be underestimated. However, this underestimation would be similar for all animals placed into the chamber. A comparison of tidal volumes between animal s is still possible, although the absolute values may not always be comparable to other mice measured with a different chamber setup. The Assumption That When Flow Is Zero During the Breath, That This Is the End of Inspiration The end of inspiration is measured w ith barometric plethysmography when the airflow is zero (in between inspiration and expi ration). Although this is most always the case for the end of inspiration, it is important to be aware that under certain conditions (i.e. panting (Johanson and Pier ce 1971)) this is not the ca se. Under these conditions

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73 tidal volume would be overestimated. However, to induce panting in rodents anesthesia is typically required, so this type of e rror is not expected on our experiments. Alterations in Frequency and Airway Re sistance Can Alter the Tidal Volume Calculation There is not complete agreement on this source of error (Malan 1973; Enhorning, van Schaik et al. 1998; Mortola and Frappell 19 98). In a series of experiments frequency and airway resistance were al tered in a step-wise fashion by using a machine, and tidal volume was calculated. The frequencies and re sistances were not physiological for small animals (and it was not a living animal that warmed and saturated the air), so it is uncertain how much this source of error may contribute to a specific experiment. However, these authors found that by increa sing frequency or resi stance, that tidal volume was also measured to be elevated. Th is is important for our experiments because the GAA-/mice have a decreased frequency compared to controls, so it is possible that the decreased frequency could contribute to the decreased tidal volume that we measure.

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74 CHAPTER 5 CONCLUSIONS AND FUTURE DIRECTIONS The successful treatment for Pompe di sease will ultimately depend on a thorough understanding of the mechanisms resulting in muscle weakness (cardiovascular, respiratory and locomotor muscle). The discovery of a neural mechanism that contributes to respir atory deficits in Gaa-/mice is contrary to th e current hypothesis that muscle dysfunction alone results in respiratory failure in Pompe disease. For this reason, these findings may help to treat adult-onset patients more effectiv ely. In addition, new therapies can be developed to target neural de ficits that may be present in Pompe disease individuals. Ongoing efforts target the use of AAV as a vector to deliver the hGaa gene to cure Pompe disease. Work in our laboratory is ai med at identifying the safest, longest lasting and effective gene therapy treatments fo r genetic diseases by first elucidating mechanisms of success and failure in animal models. The accurate characterization of these animal models is of utmost importance for translation to clinical applications. Importantly, work described in this thesis in cludes whole animal, conscious measures of ventilation which will be valuab le for quantifying recovery in Gaa-/mice following various treatments as they are developed. The success of gene replacement ther apy for Pompe disease will depend on multiple factors including the ability to generate high-expressing, non-immunogenic vectors with the ability to transfect cardia c and skeletal muscle and motoneurons. The discovery that AAV2 could be transported to lumbar motoneurons following a muscle

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75 injection was a significant cont ribution to the field of gene therapy (Kaspar, Llado et al. 2003). However, other groups have been unabl e to repeat these experiments (including work described in this thes is) (Glatzel, Flechsig et al. 2000; Martinov, Sefland et al. 2002; Wang, Muramatsu et al. 2002). Thes e issues have led to the cloning and characterization of a lternative recombinant AAV capsids, which have resulted in higher transgene expression in motoneurons fo llowing a muscular injection (Xu 2005). Importantly, these new capsids offer the possi bility to treat not only Pompe disease, but many other neuromuscular disorders resulting in respiratory failure. In addition to rAAV vector development studies, the search con tinues for candidate promoters and other gene regulatory elements capable of yielding high le vels of transgene expression with varying tissue specificities. Future work will involve utilizing new rAAV vectors in our animal model of Pompe disease for the long-term goal of transitioning this therapy to the clinic. The finding that a neural deficit may contri bute to respiratory insufficiency in Gaa-/mice necessitates the evaluation of juvenile and adult-onset patients to identify if this pathophysiology is present in Pompe disease individuals. Additional experiments in the Gaa-/mouse will also be necessary to elucid ate the time course and reversibility of respiratory insufficiency following therapy re gimes. Although we have identified a new concept in the mechanisms of respiratory de ficits in Pompe disease, there are clearly many more avenues that need to be addresse d to further elucidate the disease progression.

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89 BIOGRAPHICAL SKETCH Lara DeRuisseau went to the University of Utah from 1997-1999 for her undergraduate education. She th en transferred to the Florid a State University where she completed a bachelorÂ’s degree in exercise science. Her masterÂ’s degree was completed under the direction of Dr. J. Michael Overton at the Florida State Un iversity Program in Neuroscience. She will complete her Ph.D . degree under the direction of Dr. Barry Byrne and Dr. David Fuller at the University of Florida.