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Xanthine Oxidase Contributes to Mechanical Ventilation-Induced Diaphragmatic Oxidative Stress and Contractile Dysfunction

Permanent Link: http://ufdc.ufl.edu/UFE0022049/00001

Material Information

Title: Xanthine Oxidase Contributes to Mechanical Ventilation-Induced Diaphragmatic Oxidative Stress and Contractile Dysfunction
Physical Description: 1 online resource (106 p.)
Language: english
Publisher: University of Florida
Place of Publication: Gainesville, Fla.
Publication Date: 2008

Subjects

Subjects / Keywords: Applied Physiology and Kinesiology -- Dissertations, Academic -- UF
Genre: Health and Human Performance thesis, Ph.D.
bibliography   ( marcgt )
theses   ( marcgt )
government publication (state, provincial, terriorial, dependent)   ( marcgt )
born-digital   ( sobekcm )
Electronic Thesis or Dissertation

Notes

Abstract: Mechanical ventilation (MV) is used to mechanically assist or replace spontaneous breathing in patients who cannot sustain sufficient alveolar ventilation. The withdrawal of MV from patients is referred to as ?weaning? and problems in weaning patients from MV are common. Unfortunately, MV results in the development of diaphragmatic atrophy and contractile dysfunction that likely contribute to weaning difficulties. Importantly, oxidative stress has been critically linked to the signaling events responsible for the progression of MV-induced diaphragmatic atrophy. However, the sources of oxidants in the diaphragm during MV have not been fully elucidated. An intracellular enzyme, xanthine oxidase (XO), is capable of producing reactive oxygen species (ROS) in skeletal muscle and we hypothesized that XO plays a key role in MV-induced oxidative stress in the diaphragm. To test this postulate, we mechanically ventilated rats for 12 or 18 hours (MV) with a subset of animals that combined MV along with a XO inhibitor, oxypurinol (MVO). Indices of XO activity and protein, oxidative stress, contractile function, and atrophy were measured in the diaphragm following the experimental protocol. Our study reveals that XO activity is elevated in the diaphragm during MV and oxypurinol provides protection against oxidative injury and contractile dysfunction. Specifically, oxypurinol administration attenuated protein oxidation and lipid peroxidation in the diaphragm during MV. Further, XO inhibition attenuated MV-induced contractile dysfunction of the diaphragm at stimulation frequencies above 60 hertz at both 12 and 18 hours of MV. Together, these results reveal that XO-mediated production of oxidants is involved in MV-induced diaphragmatic oxidative stress and contractile dysfunction.
General Note: In the series University of Florida Digital Collections.
General Note: Includes vita.
Bibliography: Includes bibliographical references.
Source of Description: Description based on online resource; title from PDF title page.
Source of Description: This bibliographic record is available under the Creative Commons CC0 public domain dedication. The University of Florida Libraries, as creator of this bibliographic record, has waived all rights to it worldwide under copyright law, including all related and neighboring rights, to the extent allowed by law.
Thesis: Thesis (Ph.D.)--University of Florida, 2008.
Local: Adviser: Powers, Scott K.
Electronic Access: RESTRICTED TO UF STUDENTS, STAFF, FACULTY, AND ON-CAMPUS USE UNTIL 2010-05-31

Record Information

Source Institution: UFRGP
Rights Management: Applicable rights reserved.
Classification: lcc - LD1780 2008
System ID: UFE0022049:00001

Permanent Link: http://ufdc.ufl.edu/UFE0022049/00001

Material Information

Title: Xanthine Oxidase Contributes to Mechanical Ventilation-Induced Diaphragmatic Oxidative Stress and Contractile Dysfunction
Physical Description: 1 online resource (106 p.)
Language: english
Publisher: University of Florida
Place of Publication: Gainesville, Fla.
Publication Date: 2008

Subjects

Subjects / Keywords: Applied Physiology and Kinesiology -- Dissertations, Academic -- UF
Genre: Health and Human Performance thesis, Ph.D.
bibliography   ( marcgt )
theses   ( marcgt )
government publication (state, provincial, terriorial, dependent)   ( marcgt )
born-digital   ( sobekcm )
Electronic Thesis or Dissertation

Notes

Abstract: Mechanical ventilation (MV) is used to mechanically assist or replace spontaneous breathing in patients who cannot sustain sufficient alveolar ventilation. The withdrawal of MV from patients is referred to as ?weaning? and problems in weaning patients from MV are common. Unfortunately, MV results in the development of diaphragmatic atrophy and contractile dysfunction that likely contribute to weaning difficulties. Importantly, oxidative stress has been critically linked to the signaling events responsible for the progression of MV-induced diaphragmatic atrophy. However, the sources of oxidants in the diaphragm during MV have not been fully elucidated. An intracellular enzyme, xanthine oxidase (XO), is capable of producing reactive oxygen species (ROS) in skeletal muscle and we hypothesized that XO plays a key role in MV-induced oxidative stress in the diaphragm. To test this postulate, we mechanically ventilated rats for 12 or 18 hours (MV) with a subset of animals that combined MV along with a XO inhibitor, oxypurinol (MVO). Indices of XO activity and protein, oxidative stress, contractile function, and atrophy were measured in the diaphragm following the experimental protocol. Our study reveals that XO activity is elevated in the diaphragm during MV and oxypurinol provides protection against oxidative injury and contractile dysfunction. Specifically, oxypurinol administration attenuated protein oxidation and lipid peroxidation in the diaphragm during MV. Further, XO inhibition attenuated MV-induced contractile dysfunction of the diaphragm at stimulation frequencies above 60 hertz at both 12 and 18 hours of MV. Together, these results reveal that XO-mediated production of oxidants is involved in MV-induced diaphragmatic oxidative stress and contractile dysfunction.
General Note: In the series University of Florida Digital Collections.
General Note: Includes vita.
Bibliography: Includes bibliographical references.
Source of Description: Description based on online resource; title from PDF title page.
Source of Description: This bibliographic record is available under the Creative Commons CC0 public domain dedication. The University of Florida Libraries, as creator of this bibliographic record, has waived all rights to it worldwide under copyright law, including all related and neighboring rights, to the extent allowed by law.
Thesis: Thesis (Ph.D.)--University of Florida, 2008.
Local: Adviser: Powers, Scott K.
Electronic Access: RESTRICTED TO UF STUDENTS, STAFF, FACULTY, AND ON-CAMPUS USE UNTIL 2010-05-31

Record Information

Source Institution: UFRGP
Rights Management: Applicable rights reserved.
Classification: lcc - LD1780 2008
System ID: UFE0022049:00001


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XANTHINE OXIDASE CONTRIBUTES TO MECHANICAL VENTILATION-INDUCED DIAPHRAGMATIC OXIDATIVE STRE SS AND CONTRACTILE DYSFUNCTION By MELISSA ANN WHIDDEN 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 2008 1

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2008 Melissa Ann Whidden 2

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To my husband, family, and friends for their consta nt support and to the teachers and professors who have played a role in my education 3

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ACKNOWLEDGMENTS First, I thank my mentor Dr. Scott Powers for his guidance and continuous support during my graduate education. He is a great example as to the researcher I aspire to be. Also, I praise my supervisory committee members; Dr. Ste phen Dodd, Dr. David Cr iswell and Dr. Glenn Walter for their direction and support throughout my graduate studies I also thank Dr. Krista Vandenborne for presenting me the opportunity to be involved with an NIH T32 award. As a T32 trainee, I had the opportunity to expand my scientific career with meetings and workshops. Of course I wish to thank all laboratory members who have played a role in my achievements. Finally and most importantly, I am thankful for the experiences shared with my husband and family throughout my life and career. 4

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TABLE OF CONTENTS page ACKNOWLEDGMENTS...............................................................................................................4 LIST OF TABLES................................................................................................................. ..........8 LIST OF FIGURES.........................................................................................................................9 ABSTRACT...................................................................................................................................11 CHAPTER 1 INTRODUCTION................................................................................................................. .13 2 LITERATURE REVIEW.......................................................................................................17 Overview of Mechanical Ventilatio n-Induced Diaphragmatic Injury ...................................17 Introduction................................................................................................................... ..17 Diaphragm Response to Mechanical Ventilation............................................................18 Mechanisms of Diaphragmatic Dysfunction...................................................................19 Contractile dysfunction............................................................................................19 Atrophy.....................................................................................................................19 Protein synthesis and degradation............................................................................20 Oxidative stress........................................................................................................21 Summary.................................................................................................................................26 3 MATERIALS AND METHODS...........................................................................................31 Experiment 1: Animals.......................................................................................................... .31 Animal Model Justification.............................................................................................31 Animal Housing and Diet................................................................................................31 Experimental Design.......................................................................................................31 Animal Protocol...............................................................................................................32 Statistical Analysis..........................................................................................................3 3 Experiment 2: Animals.......................................................................................................... .33 Animal Model Justification.............................................................................................33 Animal Housing and Diet................................................................................................33 Experimental Design.......................................................................................................34 Animal Protocol...............................................................................................................34 Statistical Analysis..........................................................................................................3 5 Experiment 3: Cell Culture..................................................................................................... 35 Experimental Design.......................................................................................................35 Cell Protocol....................................................................................................................35 Statistical Analysis..........................................................................................................3 7 General Methods.....................................................................................................................37 Animal Measurements.....................................................................................................37 5

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Histological..............................................................................................................37 Functional.................................................................................................................38 Biochemical..............................................................................................................38 Cell Culture Measurements.............................................................................................40 4 RESULTS...................................................................................................................... .........45 Experiment 1................................................................................................................... ........45 Systemic Response to MV......................................................................................................45 XOR Characteristics............................................................................................................ ...45 XOR Localization............................................................................................................45 XDH and XO Activity Levels.........................................................................................46 XDH and XO Protein Levels...........................................................................................47 Enzyme Substrates and End Product......................................................................................48 Hypoxanthine..................................................................................................................48 Xanthine..........................................................................................................................49 Uric Acid...................................................................................................................... ...49 Redox Balance........................................................................................................................50 Protein Carbonyls............................................................................................................50 4-HNE..............................................................................................................................50 Total Glutathione.............................................................................................................51 Contractile Function........................................................................................................... ....51 Maximal Isometric Twitch Force....................................................................................51 Force-Frequency Response.............................................................................................52 Diaphragmatic Atrophy.......................................................................................................... 52 Cross-sectional Area........................................................................................................52 Experiment 2................................................................................................................... ........53 Systemic Response to MV......................................................................................................53 Redox Balance........................................................................................................................53 Protein Carbonyls............................................................................................................53 4-HNE..............................................................................................................................54 Contractile Function........................................................................................................... ....54 Maximal Isometric Twitch Force....................................................................................54 Force-Frequency Response.............................................................................................55 XOR Characteristics............................................................................................................ ...55 XDH and XO Activity Levels.........................................................................................55 XDH and XO Protein Levels...........................................................................................56 Enzyme Substrates and End Product......................................................................................56 Experiment 3................................................................................................................... ........57 Myogenic Cells................................................................................................................. ......57 Cell Viability...................................................................................................................57 XDH and XO Activity Levels.........................................................................................57 XDH and XO Protein Levels...........................................................................................58 Calpain Protein and Activity...........................................................................................59 6

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5 DISCUSSION................................................................................................................... ......85 Overview of Principal Findings..............................................................................................85 Mechanical Ventilation-induced Induction of XO Activity............................................85 XO Inhibition during MV Attenuates Di aphragmatic Oxidative Stress and Contractile Dysfunction...............................................................................................87 Antioxidant Administrati on during MV Fails to Attenuate XO Activation....................89 Hydrogen Peroxide Acti vates XO in Myotubes..............................................................91 Conclusions and Future Directions.........................................................................................93 LIST OF REFERENCES...............................................................................................................95 BIOGRAPHICAL SKETCH.......................................................................................................106 7

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LIST OF TABLES Table page 2-1 Summary of MV-induced changes in th e diaphragm observed during 3 hours of MV.....................................................................................................................................274-1 Total xanthine oxidoreductase (XOR), xanthine dehydrogenase (XDH), and xanthine oxidase (XO) activities in diaphragm from control, mechanically ventilated, and mechanically ventilated animals with oxypurinol.............................................................614-2 Maximal isometric twitch force produc tion in diaphragm strips obtained from mechanically ventilated and nonmechanically ventilated (control) animals with and without oxypurinol.............................................................................................................624-3 Maximal isometric twitch force produc tion in diaphragm strips obtained from control, mechanically ventilated, and mech anically ventilated animals with Trolox........704-4 Hypoxanthine, xanthine, and uric acid levels in control, mechanically ventilated, and mechanically ventilated animals with Trolox....................................................................71 8

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LIST OF FIGURES Figure page 2-1 Effects of prolonged MV on the diap hragmatic force-frequency response ( in vitro ) in rats......................................................................................................................................28 2-2 Potential sources of ROS production in the diaphragm during MV..................................29 2-3 Two proposed mechanisms of XO ac tivation in the diaphragm during MV.....................30 3-1 Experimental animal design used to determine if pharmacological inhibition of XO activity reduces MV-induced diaphragma tic oxidative stress and contractile dysfunction.........................................................................................................................42 3-2 Experimental animal design used to examine whether the administration of an exogenous antioxidant during MV preven ts XO activation in the diaphragm..................43 3-3 Experimental cell culture design used to determine if a ROS challenge activates XO in skeletal muscle myotubes..............................................................................................44 4-1 Xanthine oxidoreductase (XOR) localization in a contro l diaphragm muscle sample......63 4-2 Ratio of xanthine oxidas e (XO) activity to xanthine dehydrogenase (XDH) activity in diaphragm samples from experiment 1..............................................................................63 4-3 Fold changes (versus control) of XOR protein (150 kDa band) content in diaphragm samples from experiment 1................................................................................................64 4-4 Fold changes (versus control) of XO R protein (130 kDa) content in diaphragm samples from experiment 1................................................................................................64 4-5 Hypoxanthine levels in diaphr agm samples from experiment 1........................................65 4-6 Xanthine levels in diaphragm samples from experiment 1................................................65 4-7 Uric acid levels in diaphr agm samples from experiment 1................................................66 4-8 Protein carbonyl levels in dia phragm samples from experiment 1....................................66 4-9 Fold changes (versus control) of 4-hydroxynonenal (4-HNE) accumulation in diaphragm samples from experiment 1..............................................................................67 4-10 Total glutathione concentrations in diaphragm samples from experiment 1.....................67 4-11 Diaphragmatic force-frequency response ( in vitro ) of diaphragm samples from experiment 1................................................................................................................... ....68 9

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4-12 Type I fiber cross-sect ional area in diaphragm samples from experiment 1.....................68 4-13 Type IIa fiber cross-sectional area in diaphragm samples from experiment 1..................69 4-14 Type IIb/x fiber cross-se ctional area in diaphragm samples from experiment 1...............69 4-15 Protein carbonyl levels in dia phragm samples from experiment 2....................................72 4-16 Fold changes (versus control) of 4hydroxynonenal (4-HNE) accumulation in diaphragm samples from experiment 2..............................................................................72 4-17 Diaphragmatic force-frequency response ( in vitro ) of diaphragm samples from experiment 2................................................................................................................... ....73 4-18 Xanthine dehydrogenase (XDH) activity in diaphragm samples from experiment 2........73 4-19 Xanthine oxidase (XO) activity in diaphragm samples from experiment 2......................74 4-20 Fold changes (versus control) of XOR protein (150 kDa band) content in diaphragm samples from experiment 2................................................................................................74 4-21 Fold changes (versus control) of XOR protein (130 kDa band) content in diaphragm samples from experiment 2................................................................................................75 4-22 Xanthine dehydrogenase (XDH) activity in C2C12 myotubes from experiment 3.............76 4-23 Xanthine oxidase (XO) activity in C2C12 myotubes from experiment 3...........................77 4-24 Fold change (versus control) of XOR protein (150 kDa band) in C2C12 myotubes from experiment 3.............................................................................................................. 78 4-25 Fold change (versus control) of XOR protein (130 kDa band) in C2C12 myotubes from experiment 3.............................................................................................................. 79 4-26 Fold change (versus control) of total calpain-1 protein in C2C12 myotubes from experiment 3................................................................................................................... ....80 4-27 Fold change (versus control) of cleaved calpain-1 protein in C2C12 myotubes from experiment 3................................................................................................................... ....81 4-28 Fold change (versus control) of total calpain-2 protein in C2C12 myotubes from experiment 3................................................................................................................... ....82 4-29 Fold change (versu s control) of total II-spectrin protein (250 kDa) in C2C12 myotubes from experiment 3.............................................................................................83 4-30 Fold change (versu s control) of cleaved II-spectrin protein (145 kDa) in C2C12 myotubes from experiment 3.............................................................................................84 10

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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 XANTHINE OXIDASE CONTRIBUTES TO MECHANICAL VENTILATION-INDUCED DIAPHRAGMATIC OXIDATIVE STRE SS AND CONTRACTILE DYSFUNCTION By Melissa Ann Whidden May 2008 Chair: Scott K. Powers Major: Health and Human Performance Mechanical ventilation (MV) is used to m echanically assist or replace spontaneous breathing in patients who cannot su stain sufficient alveolar ventil ation. The withdrawal of MV from patients is referred to as weaning and problems in weaning patients from MV are common. Unfortunately, MV results in the development of diaphragmatic atrophy and contractile dysfunction that likely contribute to weaning difficulties. Importantly, oxidative stress has been critically linke d to the signaling events respons ible for the progression of MVinduced diaphragmatic atrophy. However, the s ources of oxidants in the diaphragm during MV have not been fully elucidated. An intracellula r enzyme, xanthine oxidas e (XO), is capable of producing reactive oxygen species (ROS) in skelet al muscle and we hypot hesized that XO plays a key role in MV-induced oxidative stress in th e diaphragm. To test this postulate, we mechanically ventilated rats fo r 12 or 18 hours (MV) with a subset of animals that combined MV along with a XO inhibitor, oxypurinol (MVO). Indices of XO activity and protein, oxidative stress, contractile function, and atrophy were measured in the diaphragm following the experimental protocol. Our study reveals that XO activity is elevated in the diaphragm during MV and oxypurinol provides protection against ox idative injury and c ontractile dysfunction. Specifically, oxypurinol administrati on attenuated protein oxidation and lipid peroxidation in the 11

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diaphragm during MV. Further, XO inhibition attenuated MV-i nduced contractile dysfunction of the diaphragm at stimulation frequencies above 60 hertz at both 12 and 18 hours of MV. Together, these results reveal that XO-media ted production of oxidant s is involved in MVinduced diaphragmatic oxidative stress and contractile dysfunction. 12

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CHAPTER 1 INTRODUCTION Mechanical ventilation (MV) is used clini cally to defend pulmonary gas exchange in patients who are unable to sustain sufficient alveolar ventilation. Respiratory failure, spinal cord injury, drug overdose, and surgery are among the various conditions that render the pulmonary system unable to maintain blood gas homeostasis (30, 44). While MV supports ventilation in periods of respiratory distress, removal from the ventilator, termed weaning, is often difficult. In fact, problems in weaning patients from MV are common, whereby ~25% of the MV population has trouble weaning from the ventilator (70). Regrettably, weaning difficulties account for almost half of the total time spent on the ventila tor (29), so understanding the mechanism(s) that contribute to weaning failure is imperative. There is accumulating evidence that weaning problems are linked to inspiratory muscle dysfunction which results in the inability of the respiratory muscles to maintain adequate ventilation. Specifically, our laboratory has shown that respir atory muscle weakness produced by prolonged MV is due to diaphragmatic atr ophy and contractile dys function (79, 80, 98, 108, 110). Moreover, we have demonstrated that MV-induced diaphragmatic contractile dysfunction is directly linked to oxidative stress (12, 31, 81, 110, 122, 132). The generation of reactive oxygen species (ROS) in the diaphragm during MV can damage proteins, lipids and DNA and interrupt normal cell signaling. In fact, oxidation of both actin and myosin occurs in the diaphragm within the first 6 hours of MV (132). This is significan t because oxidation of contractile proteins contributes to abhorrent excitation-cont raction coupling and decreased muscle force production (132). Therefore, it is necessary to identify the cellular pathways responsible for ROS production in the diaphragm during MV. 13

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Numerous ROS producing pathways exist in skeletal muscle and they include; nicotinamide adenine dinucleotide phosphate (NAD( P)H) oxidase, nitric oxi de synthase (NOS), heme-oxygenase-1 (HO-1), mitochondria, and xanthi ne oxidase (XO). With regards to MV, one or more of these pathways may contribute to the oxidative stress that is observed in the diaphragm. Our laboratory has already examin ed both the NOS and NAD(P)H oxidase pathways in the diaphragm and their contribution to ROS production during MV. Specifically, we revealed that MV-induced oxidative stress in the di aphragm is not due to in creases in nitric oxide (122). Likewise, we have show n that NAD(P)H oxidase is not a substantial source of oxidant production in the diaphragm during prolonged m echanical ventilati on (McClung et al., unpublished). Therefore, we have focused on xanthine oxida se (XO) as a potential pathway involved in the formation of ROS in the diaphragm during MV. Xanthine oxidoreductase (XOR) is an intracellular enzyme that has been localized in the cytosol of skeletal muscle fibers (40, 49). XOR is involved in purine catabolism where the enzyme catalyzes the reduction of hypoxanthine and xanthine to uric acid (40, 42). Importan tly, XOR exists in two interconvertible forms, xanthine dehydrogenase (XDH) and xanthine oxidase (XO) (39, 42, 130). In the dehydrogenase form, the enzyme utilizes nicotinamide adenine dinucleotide (NAD+) as it s electron acceptor in purine catabolism. However, in the oxidase fo rm, because molecular oxygen is used as the electron acceptor instead of NA D+, hypoxanthine and xanthine are reduced to uric acid and superoxide. Therefore, only the XO form is capable of producing ROS. In skeletal muscle, the level of superoxide production th at occurs due to this pathway is dependent upon the levels of XO present and the ratio of XO to XDH in the fiber. Since XO produces superoxide and is found in skeletal muscle of both rats and human s (40, 42) it may contri bute to ROS production in 14

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the diaphragm during MV. Therefore, these expe riments investigated the contribution of XO to ROS production in the diaphragm during MV and examined potential mechanism(s) by which XO may be activated in skeletal muscle. Ou r experiments were designed to achieve the following specific aims. Specific Aim 1: To determine if pharmacological inhibition of XO activity in the diaphragm reduces MV-induced oxidative stress and contractile dysfunction. Rationale: Our work indicates that MV results in diaphragmatic oxidative injury as a result of increased ROS production (12, 31, 80, 110, 122, 132). Despite the fact that there is significant evidence that XO plays an important role in oxidant production in diseased cardiac muscle (25, 27, 50, 68, 82, 103), it is unclear whether XO contribute s to oxidant production in the diaphragm during MV. Hypothesis: The administration of a XO inhibito r will attenuate diaphragmatic MVinduced oxidative stress a nd contractile dysfunction. Specific Aim 2: To ascertain whether the administ ration of an exogenous antioxidant during MV maintains redox balance in th e diaphragm and prevents XO activation. Rationale: The oxidation of cysteine residues on the dehydrogenase form of XOR results in the reversible conversion of XDH to XO and s ubsequent XO activation (85, 101). Therefore, it is feasible that during MV, XO may be activat ed via oxidants produced from pathways other than XO. Previous work in our laboratory has shown that the administration of Trolox, a watersoluble Vitamin E analog, reduces both oxidativ e stress and contractile dysfunction in the diaphragm during MV (12, 80, 81). However, it is unknown whether Trolox administration reduces XO activation in th e diaphragm during MV. 15

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Hypothesis: Trolox administration during MV w ill maintain redox balance in the diaphragm and attenuate MV-induced XO activation. Specific Aim 3: To determine if a ROS challenge (hydrogen peroxide) activates XO in skeletal muscle myotubes. Rationale: In non-muscle cell lines, hydrogen peroxide (H2O2) has been shown to modulate the reversible conversion of XDH to XO (85), s uggesting a possible role for free radical production via other pathways in the induction of XO activity. However, it is unknown whether hydrogen peroxide activates XO in skelet al muscle myotubes. By utilizing murine myotubes in vitro, we will examine the mechanism(s) by which XO is activated in skeletal muscle. Hypothesis: Treatment of skeletal muscle myotubes with hydrogen peroxide will increase XO activity via the reversible conversion of XDH to XO. 16

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CHAPTER 2 LITERATURE REVIEW Mechanical ventilation (MV) is used to ach ieve satisfactory pulmonary gas exchange in patients incapable of maintaini ng adequate alveolar ventilation. The withdrawal of MV from patients is referred to as weaning and problem s in weaning from MV are common. Numerous studies indicate that MV-induced diaphragmatic weakness, due to both atrophy and contractile dysfunction, is an important contributor to weaning difficulties. Although the specific mechanisms responsible for MV-induced diaphragmatic weakness remain unknown, it is now clear that oxidative st ress in the diaphragm plays a major role in regulating the signaling pr ocesses leading to MV-induced diaphragmatic dysfunction. It follows that understanding the sources of oxidant production in the dia phragm during prolonged MV is important. Hence, this forms the rationale for th e experiments contained within this dissertation. Specifically, our experiments were designed to investigate the role that a specific oxidant production pathway (i.e. xanthine oxidase) plays in MV-induced oxidative injury in the diaphragm. This chapter will discuss the importance of our experimental work and will develop the ideas behind our hypotheses based up on our prior research and the wo rk of others. Specifically, this review will be divided into two concis e segments: 1) an overview of MV-induced diaphragmatic injury; and 2) a detailed discussi on of oxidant producing pathways in skeletal muscle. Overview of Mechanical Ventilation -Induced Diaphragmatic Injury Introduction Mechanical ventilation (MV) is used clini cally for patients with respiratory distress. Specifically, when an individual is unable to su stain adequate alveolar ventilation on their own, 17

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MV is required to maintain adequate gas exchange. Unfortunately, problems in weaning patients from MV are common (70). Failure to wean pati ents from MV is a problem because increased time on the ventilator increases complications and the incidence of morbidity and mortality outcomes (17, 70, 121, 125, 126). In addition, longer stays in the intensiv e care unit for extra weaning time result in excess health care co sts (70). Thus, the identification of the mechanism(s) responsible for MV-induced diaphragmatic weakness is imperative. Diaphragm Response to Mechanical Ventilation Our laboratory has extensively examined MV in a rat model. We have documented the changes that occur in the diaphragm muscle that likely contribute to re spiratory muscle weakness and weaning difficulties. Our findings are presen ted in Table 2-1 in a time course dependent manner (12, 23, 31, 79-81, 98, 108-110, 122, 123, 132). While our laboratory uses a rat model of MV, there is abundant evidence from other an imal models (rabbits, pigs, and baboons) along with recent data from human research that s upport the concept that pr olonged MV results in diaphragmatic dysfunction (4, 10, 32-34, 51, 69, 71, 93, 99, 100, 104-106, 124, 129, 131, 133). Collectively, these studies clearly document the damaging effects of MV on the diaphragm. While all animal models of MV report a wide array of detrimental effects on the diaphragm, limited human MV studies exist. B ecause of the invasive nature of obtaining a biopsy from a human diaphragm, human MV studies are difficult to conduct. However, a recent study demonstrates that prolonge d MV results in diaphragmatic atrophy in humans (71). Specifically, Levine et al. observed an approximate 40% decrease in cross-sectional area across both type I and type IIa diaphragm fibers in pa tients ventilated between 18-72 hours (71). In another human study, investigators found that twitch transdiaphragma tic pressure in individuals with disease was 50% lower than healthy patient s following MV (129). Fi nally, a retrospective 18

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analysis of postmortem data obtained from neona tes who received ventilatory assistance for 12 days or more reported diffuse diaphragmatic fiber atrophy, which was not observed in locomotor muscles (58). Mechanisms of Diaphragmatic Dysfunction Contractile dysfunction Problematic weaning is often associated with respiratory muscle weakness. In baboons, Anzueto and colleagues demonstrated a decr ease in both maximal diaphragmatic force production and endurance after 11 days of MV (4). In a rat model of MV, Le Bourdelles et al. documented a significant 60% reduction in maximal diaphragmatic specific force following 48 hours of MV (69). Our laboratory has shown th at the MV-induced decline in diaphragmatic submaximal and maximal specific force increases as a function of time on the ventilator (Figure 2-1) (98). Specifically, maximal diaphragmatic specific force is approximately 18% and 46% lower in animals ventilated for 12 and 24 hours re spectively, when compared to control animals (98). Atrophy MV-induced diaphragmatic atrophy has been observed in both animal and human experiments (4, 10, 33, 58, 69, 71, 79, 80, 110, 131). Wh ile atrophy occurs during many types of skeletal muscle disuse, the rate of atrophy during MV is extraordinarily fast when compared with hindlimb muscle unloading (69, 79, 131). For example, only 12 hours of MV results in a significant reduction in diaphragma tic myofiber size. Work by Sh anley et al. revealed a 15% reduction in myofiber cross-sec tional area across all four fibe r types in the rat diaphragm following 18 hours of MV (110). To achieve this level of atrophy in hindlimb skeletal muscle, muscles would need to be unloaded for more th an 96 hours (120). As mentioned above, Levine 19

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and colleagues found a similar reduction in myofib er size in the human diaphragm following 18 72 hours of MV (71). Protein synthesis and degradation Diaphragmatic atrophy can result from a dec line in protein synthesis (66) and/or an increase in protein degradation ( 13). The decrease in protein s ynthesis often observed in models of skeletal muscle disuse is characterized by alterations in tr anslational initiation and elongation and/or a decrease in cellular RNA (48, 66, 89, 90). Our laboratory has observed a decrease in protein synthesis in as few as 6 hours following MV (109). With respect to the protein synthetic signaling pathway, McClung and colleagues found that MV increases the activation of the translational repressor protein 4E-BP1 and decreas es the activity of the translational initiation factor p70s6kinase (81). Both the increase in 4E-BP1 and the decrease in p70s6kinase signify early events in skeletal muscle atrophy (81). When protein sy nthesis is reduced and myofiber myosin heavy chain content is minimized during MV, diaphragmatic atrophy ensues. Nevertheless, although MV results in decreased protein synthesis in the diaphragm our laboratory has demonstrated that the development of atrophy during MV is primarily due to an increase in protein degradation (23, 80, 81, 110). There are several proteolytic systems that contribute to skeletal muscle degradation during MV, including the lysosomal, calpa in, caspase-3, and proteasome pathways (8, 97, 127). While currently it is not known whether the lysosomal pathway plays a major role in MV-induced atrophy (reviewed in 97), both calpain and caspase pr oteases are believed to cleave proteins that anchor the contractile elements to the myofilame nt (19). Thus, during MV, calpain and caspase3 activation may act as the initial step in muscle protein lo ss during diaphragmatic atrophy. Following the release of myofibrillar proteins, the ubiquitin proteasome pathway appears to be 20

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the major proteolytic system responsible for sk eletal muscle protein degradation. Protein degradation by this pathway invol ves substrate recognition via the coordinated action of a threeenzyme system and protein degradation by the 26S proteasome (23). Ou r laboratory has shown that MV increases diaphragmatic levels of tw o ubiquitin ligases and increases 20S proteasome activities including chymotrypsin and trypsin-like activities (23). Oxidative stress ROS are produced in inactive skeletal muscles and oxidative stress-induced cellular injury can contribute to muscle atrophy (23, 61-65). Si nce the diaphragm muscle is completely inactive during controlled MV (98, 106) the potential for ROS production exists. In skeletal muscle, when ROS production exceeds the antioxidant capac ity, oxidative stress occurs (96). We have shown that within 6 hours after the initiation of MV, oxidative in jury in the diaphragm occurs (132). Oxidative stress can alter the structure, function and/or re gulation of lipids, proteins, and DNA in the cell. Therefore, it is important to identify the sources of ROS that contribute to redox imbalances in the muscle cell. Numer ous ROS producing pathways exist in the cell (Figure 2-2) and they include; 1) nicotinam ide adenine dinucleotide phosphate (NAD(P)H) oxidase, 2) nitric oxide syntha se (NOS), 3) heme oxygenase-1 (HO-1), 4) mitochondria, and 5) xanthine oxidase (XO). A brief overvie w of each of these pathways follows. NAD(P)H Oxidase NAD(P)H oxidase is a membrane-associated enzyme that catalyzes the one-electron reduction of molecular oxygen into superoxide usi ng either NADH or NADPH as the electron donor (52). It has been shown that the calcium-sensitive PKC-ERK1/2 pathway can activate NAD(P)H oxidase activity in cells (41). While our laboratory has found that NAD(P)H oxidase inhibition via apocynin reduces MV-induced oxidative stress a nd contractile dysfunction, diaphragmatic NAD(P)H oxidase activ ity is only increased 4% during MV when compared with controls (McClung et al., unpublished). Although the contribution of NAD(P)H 21

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oxidase to MV-induced oxidative st ress appears to be small, it may act as an up-stream regulator of other pathways that generate ROS. Thus, NAD(P)H oxidase inhibition could reduce both MV-induced oxidative stress and contractile dysfunction by down-regulating other ROS producing pathways. Nitric Oxide Synthase Three isoforms of NOS exist (54, 112), however, only neuronal NOS (nNOS) and endothelial NOS (eNOS) are expressed in diaphragm muscle (54, 112, 122). Endogenous nitric oxide production via NOS can re sult in the formati on of reactive nitrogen species, including peroxynitrite (ONOO-). Reactive nitrogen species are associated with cellular injury including mitochondrial dysfunction, lipid pe roxidation, and nitrosylat ion of proteins (7, 112, 118). In reference to MV-induced oxidative stress in the diaphragm muscle, Van Gammeren et al. demonstrated that nitric oxide is not involved in MV-induced oxidative injury in the diaphragm (122). Heme Oxygenase-1 Heme oxygenase-1 (HO-1) is an intracellular enzy me localized in the microsomal fraction of the cell (55). HO-1 cat alyzes the rate-limiting step in the degradation of heme resulting in the generation of car bon monoxide, biliverdin, and free iron (119). Biliverdin can then be further reduced to bilirub in. Since bilirubin posses the ability to quench singlet oxygen, scavenge hypochlorou s acid, and inhibit lipid peroxida tion (18), the possibility of radical formation exists due to the release of free iron. Transiti onal metal ions such as iron are capable of converti ng superoxide and hydrogen peroxide to the highly reactiv e hydroxyl radical (67). However, our laboratory has recently s hown that HO-1 acts as an antioxidant in the diaphragm by demonstrating that hemin-induced overexpression of HO-1 is associated with reduced oxidative stress during MV (Falk et al., unpublished). More importantly, HO-1 22

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inhibition in the diaphragm during MV does not alter the norm al progression of MV-induced oxidative stress (Fal k et al., unpublished). Mitochondrial oxidants While the primary function of the mitochondria is to produce ATP, electron leak from the electron transport chain may occur at complexes I and III resulting in the formation of superoxide and subsequent ly hydrogen peroxide (3, 16, 96). However, Fredriksson and colleagues obser ved negligible mitochondrial damage in the diaphragm following 5 days of MV (32). Nonetheless, due to the limited evidence, the role of mitochondrial generated oxidants in the diaphrag m during MV should be further investigated. Xanthine Oxidase Xanthine oxidoreductase (XOR) is an intracellular enzyme localized primarily in the cytosol and it is involved in purine catabo lism (40). XOR occurs as a homodimer and each subunit contains four redo x centers; a molybdenum cofactor, a flavin adenine dinucleotide (FAD), and two iron sulf ur cluster sites (15, 40, 46). Mammalian XOR catalyzes the hydroxylation of hypoxant hine and xanthine to uric acid at the molybdenum center of the enzyme. Reducing equivale nts introduced into the enzyme ar e transferred via the two iron sulfur cluster sites to the FAD cofactor where the reduction of nicotinamide adenine dinucleotide (NAD+ ) or molecular oxygen occurs (60). Impor tantly, XOR exists in two interconvertible forms, xanthine dehydrogenase (XDH) and xant hine oxidase (XO) (39, 42, 130). XDH utilizes NAD+ as its electron acceptor in purine catabolis m. However, XO uses molecular oxygen as its electron acceptor instead of NA D+, so hypoxanthine and xanthine are reduced to uric acid and superoxide (O2 .) (shown below) (130). Therefore, only the XO form is capable of producing ROS. XO XO Hypoxanthine + O2 Xanthine + O2 .Xanthine + O2 Uric acid + O2 .23

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The two forms of XOR (XDH and XO) can be interconverted reversibly by sulfhydryl oxidation or irreversibly by proteolysis of XDH to XO (1, 9, 11, 22, 25, 27, 40, 50, 84, 85, 92, 101, 102, 113, 114, 130). XOR is initially synthesized as a 150 kDa protein from which both XDH and XO are derived (85, 130). XDH only ex ists as the 150 kDa band of XOR while XO can be reflected in the 150 kDa band as well as an active 130 kD a band of XOR. The mechanisms responsible for the reversible or irreversible conversion of XDH to XO are not completely understood. Under baseline conditions, the majority of the 150 kDa protein contains XDH activity. Reversible formation of XDH to XO can be achieved by the oxidation of cysteine residues on XOR, a process that changes enzymatic function but not molecular weight (Figure 23) (85, 101). It is feasible that during MV, diaphragmatic XO activity co uld be up-regulated via the oxidation of XDH to XO. As mentioned abov e, pathways other than XO may generate ROS during prolonged MV and production of oxidants from one or more of those pathways may result in the oxidation of XDH. The subsequent XO activation and further ROS production through purine catabolism would contribute to oxidative damage in the diaphragm during MV. Further, it is also possible that XO activati on during MV may occur th rough the proteolytic conversion of XDH to XO (22, 27, 85, 101, 102, 113, 114, 130). The irreversible proteolysis of XDH occurs at a specific site leading to the fo rmation of an active 130 kD a form of XO (Figure 2-3) (130). While the protease responsibl e for this conversion has been examined by investigators since the late 1960s, debate continue s as to the specific identity of this protease (22, 102, 113, 114, 130). While some investigations suggest that a calpain-independent calcium (Ca2+) activated protease functions in the cleavag e of XDH to XO (1, 92, 102, 113), it has also been theorized that calpain is responsible for the cleavage in various tissues (39, 42). Since 24

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calpain activity is increas ed in the diaphragm during MV (110), it is a potential candidate for the protease responsible for the cleavage. Attention was first directed towards XO when Granger and colleagues examined the role of XO-derived ROS production in ischemia-reperf usion injury (37, 38, 82). These researchers found that during the course of ischemia, transm embrane ion gradients were dissipated allowing elevated cytosolic concentrations of calcium into the cell. The increase in intracellular calcium activated a protease that irreversibly converted XDH into XO. At the same time, cellular ATP was catabolized to hypoxanthine which accumulated over the duration of ischemia. Upon reperfusion and the restorati on of oxygen, hypoxanthine was reduced to uric acid, superoxide, and hydrogen peroxide (37, 38, 82). Since then several studies ha ve examined the role of XOmediated ROS production in other tissues unde r varying physiological states (1, 11, 25, 28, 36, 42, 43, 45, 50, 53, 56, 68, 78, 84, 103, 117, 128). In doing so, these studies have used allopurinol or oxypurinol to e ffectively inhibit XOR activity. Allopurinol (1,5-dihydro-4Hpyrazolo (3,4-d) pyrimidine-4-one) is a struct ural analogue of hypoxanthi ne and inhibitor of xanthine oxidoreductase (88). It has a half-life of approxi mately 2 hours and is quickly metabolized to oxypurinol which has a half-life of ~18-30 hours (88). In cardiac tissue XO has been found to contribut e to abnormal excitation-contraction (EC) coupling and cardiac remodeling in heart failure (103). For inst ance, Saliaris and colleagues found that XO inhibition preserved the positive in otropic effects of dobutamine that were lost during heart failure (103). Likewise, Isabelle et al. found th at XO inhibition prevents the myocardial production of superoxi de anions in cocaine-induced le ft ventricular dysfunction in male rats (50). In this model, XO inhibiti on prevented cocaine-induced cardiac alterations by restoring cardiac output, str oke volume, and fractional shor tening (50). Finally, in an 25

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26 atherosclerosis study, McNally et al. reported that XO is responsible for increased ROS production in response to oscillator y shear stress that occurs at sites of circulation that are vulnerable to disease (84). Recent evidence indicates that XO is found in skel etal muscle fibers of humans (42). In a study by Vina and colleagues, they found that XO is responsible for the free radical production and tissue damage that occurs during exhaustive exercise in humans (128). In COPD patients, XO inhibition reduced both glutathione oxidation and lipid peroxidation during strenuous exercise (45). Finally, it has been shown th at XO contributes to oxi dant production in the diaphragm during contraction (115) However, the XO pathway has not been studied with regard to oxidant production in the dia phragm during MV. This important void forms the basis for the current experiments. Summary Our laboratory has developed a rat animal m odel for the study of MV-induced atrophy and contractile dysfunction of th e diaphragm muscle (12, 20, 79, 80, 98, 108, 110). By utilizing aseptic techniques and maintaining blood gas homeostasis, we have repeatedly demonstrated that this is an excellent animal model for examining the mechanisms that contribute to diaphragmatic injury during MV. Our laboratory has investigat ed the effects of oxidativ e stress, proteolysis, and atrophy on diaphragm myofiber function during various durations of MV. However, the oxidant pathways that contribute to MV-induced oxidative injury in the diaphragm are currently unknown. Since XO can generate superoxide in the presence of oxygen and the purine substrates, determining if XO contributes to MV-induced oxidative stress in the diaphragm is important and may lead to a therapeutic strate gy to alleviate MV-induced diaphragmatic injury.

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Table 2-1. Summary of MV-induc ed changes in the diaphragm observed during 3 hours of MV. Measurement 3 hrs 6 hrs 12 hrs 18 hrs 24 hrs Oxidative stress ND Caspase-3 activity ND ND ND Protein synthesis ND ND Protein degradation ND ND Contractile dysfunction ND ND Atrophy ND ND Myonuclear apoptosis ND ND ND ND; no data available, ; not significantly different from control value, ; increased vs. control value, ; further increased, ; decreased vs. control value, and ; further decreased. Data are from re ferences (12, 23, 31, 79-81, 98, 108-110, 122, 123, 132).27

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Stimulation frequency (Hz) 020406080100120140160180Specific force (N/cm2) 0 5 10 15 20 25 Control 12 hrs MV 18 hrs MV 24 hrs MV * Figure 2-1. Effects of prolonged MV on the diaphragmatic force-frequency response ( in vitro ) in rats. Values are means SE. Compared with control, MV (all durations) resulted in a significant (*P < 0.05) reduction in diaphragmatic specific force production at all stimulation frequencies (Redrawn from 98). 28

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Figure 2-2. Potential sources of ROS productio n in the diaphragm during MV. Prolonged MV may cause an increase in one or more of the following sources of oxidants in the diaphragm: 1) NAD(P)H oxidase 2) nitric oxide synthase 3) heme oxygenase-1, 4) mitochondria, and 5) xanthine oxidase. Through these mechanisms, MV may cause an increase in the level of oxidative stress and subsequent injury to the diaphragm. 29

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Figure 2-3. Two proposed mechanisms of XO ac tivation in the diaphragm during MV. 1) MV can activate ROS-producing pathways that result in the oxidat ion and reversible conversion of XDH to XO (dashed arrows). 2) MV can activate a cellular protease that can irreversibly cleave XDH to XO (dotted arrows). Following either activation mechanism, XO catalyzes the reduction of hypoxanthine to xanthine and superoxide (O2 .-) and subsequently reduces xanthine to uric acid and additional superoxide (O2 .-). 30

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CHAPTER 3 MATERIALS AND METHODS This chapter will be divided into two secti ons. Section one includes the experimental designs used in each of our experiments that are intended to determine if XO contributes to oxidative stress and contractile dysfunction during MV and establ ish the mechanism(s) by which XO is activated in the diaphragm. In the subs equent section, we will provide the methodological details associated with each experiment al protocol and measurement technique. Experiment 1: Animals Animal Model Justification To address our first specific aim and determ ine if the pharmacological inhibition of XO activity in the diaphragm reduces MV-induced ox idative stress and cont ractile dysfunction, we used adult (4-6 month old) female Sprague-Dawley (SD) rats in experiment 1. The animals were 4-6 months of age (young adult) at the time of s acrifice. The SD rat was chosen due to the similarities between the rat and human dia phragm in both anatomical and physiological parameters (5, 6, 86, 87, 94, 95). Animal Housing and Diet All animals were housed at the University of Florida Animal Care Services Center according to guidelines set forth by the Institutional Animal Care and Use Committee. The Animal Care and Use Committee of the University of Florida has approved these experiments. Animals were maintained on a 12:12 hour reve rse light-dark cycle and provided food (AIN93 diet) and water ad libitum throughout the experimental period. Experimental Design In experiment 1, adult rats were randomly assigned to one of the following groups; 1) acutely anesthetized control (C ON) (n = 8), 2) acutely anesthetized control with oxypurinol 31

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(CONO) (n = 8), 3) 12 hours of MV (12MV) (n = 8), 3) 12 hours of MV with oxypurinol (12MVO) (n = 8), 4) 18 hours of MV (18MV) (n = 8), and 5) 18 hours of MV with oxypurinol (18MVO) (n = 8) (Figure 3-1). Animal Protocol Animals in the control groups were acutely an esthetized with an intraperitoneal (i.p.) injection of sodium pentobarbital (60 mg/kg). After reaching a su rgical plane of anesthesia, the control animals were sacrificed immediately and a section of the costal diaphragm was used for contractile measurements while the rest was stored at C for subsequent analyses. Animals in the MV groups were acutely anesth etized with sodium pentobarbital (60 mg/kg IP). After reaching a surgical plane of anesth esia, the animals were tracheostomized utilizing aseptic techniques and mechanically ventilated with a controlled pressure-driven ventilator (Seimens) for 12 or 18 hours with the following settings: upper airway pressure limit: 20 cmH2O, PEEP: 1 cmH2O, pressure control level above PEEP: 4-6 cmH2O, and respiratory rate: 80 bpm (12, 123). We choose 12 and 18 hours of mechanical ventilation in order to establish a time course for these measurements because these periods of MV are associated with diaphragmatic oxidative stress, contractile dys function, and myofiber atrophy. Animals in the oxypurinol groups (CONO, 12MVO, and 18MVO) received two intraperitoneal injecti ons of oxypurinol (25 mg/kg body weight ) 24 hours prior to treatment. Further, an additional 25 mg/kg body weight was administered via an i.p. injection 12 hours prior to sacrifice in the CONO animals and via infusion over a 10 min peri od before the initiation of MV in the 12MVO and 18MVO groups. This protoc ol has been reported to effectively inhibit XO activity in rat skeletal muscle (57). Animal s in the MV group received an i.p. injection of saline and surgical preparation, pr ocedures, and animal monitoring were performed as previously 32

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described (31). Following the completion of eac h experimental protocol, the animals were immediately sacrificed and a section of the costal diaphr agm was used for contractile measurements while the rest was stored at C for subsequent analyses. Statistical Analysis Group sample size was determined using a power analysis of preliminary data from our laboratory. Comparisons between groups were made by a one-way ANOVA and, when appropriate, a Tukey HSD test was performed. Significance was established at P < 0.05. Experiment 2: Animals Animal Model Justification To address our second specific aim and to as certain whether the administration of an exogenous antioxidant during MV prevents diaphr agmatic XO activation, adult (4-6 month old) female Sprague-Dawley (SD) rats were used for experiment 2. The animals were 4 months of age (young adult) at the time of sacr ifice. The rationale for select ing the rat as an experimental model was discussed previously. Animal Housing and Diet All animals were housed at the University of Florida Animal Care Services Center according to guidelines set forth by the Institutional Animal Care and Use Committee. The Animal Care and Use Committee of the University of Florida has approved these experiments. Animals were maintained on a 12:12 hour reve rse light-dark cycle and provided food (AIN93 diet) and water ad libitum throughout the experimental period. 33

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Experimental Design In experiment 2, adult rats were randomly assigned to one of the following groups; 1) acutely anesthetized control (CON) (n = 8), 2) 12 hours of MV (12MV) (n = 8), and 3) 12 hours of MV with Trolox (12MVT) (n = 8) (Figure 3-2). Animal Protocol Animals in the control group were acutely anesth etized with an intrap eritoneal in jection of sodium pentobarbital (60 mg/kg). After reaching a surgical pl ane of anesthesia, the control animals were sacrificed immediately and a sec tion of the costal diaphragm was used for contractile measurements while the rest was stored at C for subsequent analyses. Animals in the MV groups were acutely anesth etized with sodium pentobarbital (60 mg/kg IP). After reaching a surgical plane of anesth esia, the animals were tracheostomized utilizing aseptic techniques and mechanically ventilated with a controlled pressure-driven ventilator (Seimens) for 12 hours with the following se ttings: upper airway pressure limit: 20 cmH2O, PEEP: 1 cmH2O, pressure control level above PEEP: 4-6 cmH2O, and respiratory rate: 80 bpm (12, 123). Animals in the Trolox group (12MVT) received an infusion of Trolox (20 mg/kg) over a 5minute period, 20 minutes prior to the start of MV. Additionally, Trolox was infused continuously at a rate of 4 mg/kg per hour for the en tirety of the ventilatio n treatment. We have previously used this protocol to effectively maintain redox balance in the diaphragm during MV (12, 80, 81) Animals in the MV group received an i.p. injection of saline and surgical preparation, procedures, and animal monitoring we re performed as previously described (31). Following the termination of each experimental group, the animals were immediately sacrificed 34

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and a section of the costal diaphragm was used for contractile measurements while the rest was stored at C for subsequent analyses. Statistical Analysis Group sample size was determined using a power analysis of preliminary data from our laboratory. Comparisons between groups were made by a one-way ANOVA and, when appropriate, a Tukey HSD test was performed. Significance was established at P < 0.05. Experiment 3: Cell Culture To answer our third specific aim and determine if a ROS challenge activates XO in skeletal muscle myotubes, we used the C2C12 myogenic cell line for experiment 3. The C2C12 myogenic cell line was chosen because of the bi ochemical and functional similarities between these myotubes and rodent skeletal muscle fibers (26, 35, 72, 74-76). Cells were maintained in a temperature and gas controlled incu bator throughout the experiments. Experimental Design In experiment 3, an independent observati on was made by placing cells in one of the following groups; 1) no treatment (CON) (n = 6), 2) hydrogen peroxide (H2O2) (n = 6), 3) leupeptin (LEU) (n = 6), 4) hydr ogen peroxide and leupeptin (H2O2 + LEU) (n = 6), 5) siRNA for calpain-1 (Cal-1 siRNA) (n = 6), and 6) hydrogen peroxide and si RNA for calpain-1 (H2O2 + Cal-1 siRNA) (n = 6) (Figure 3-3). Cell Protocol Myogenic cells were cultured according to Li (73). Briefly, myot ubes were cultured in DMEM supplemented with 10% fetal bovine serum and gentamicin at 37C in the presence of 5% CO2. Myoblast differentiation was initiated by replacing the growth medium with differentiation medium supplemented with 2% horse serum. Differentiation was allowed to 35

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continue for 96 hours before experimentation, ch anging to fresh media every 24 hours. At the time of harvest, myotubes were washed 3x in PBS buffer (pH 7.4) before the addition of cell lysis buffer. After the addition of lysis buffer, myotubes were collected and centrifuged at 1100 g for 10 min at 4 C. The supernatant was collected and stored at C for subsequent analyses. We chose 100 M hydrogen peroxide (H2O2) for the oxidative challenge in these experiments. This concentration of H2O2 has been used in the lite rature in order to induce a stress response in C2C12 myotubes (74, 116). In addition, this concentration may have relevance to in vivo cellular levels. Therefore, we chose this concentration to induce oxidative stress in the C2C12 cells. H2O2 treatment was administered 4 hours prio r to harvest in the hydrogen peroxide, hydrogen peroxide plus leupeptin, and the hydrog en peroxide plus siRNA for calpain-1 cell groups. We choose 4 hours for our incubation pe riod because preliminary data showed that while XO activation was induced in our cell line following H2O2 treatment for 1 hour, XO activity was significantly in creased following the 4 hour incubation period. Myotubes in the leupeptin (LEU) groups were cultured in horse serum combined with 120 M leupeptin and were incubated for 4 hours alone or in combination with H2O2 prior to harvest. Leupeptin was selected as a general calpain inhibitor to examine the irreversible proteolytic conversion of XDH to XO. We c hoose 120 M leupeptin for these experiments because this concentration has been shown to effectively inhibit calpain activity in C2C12 cells (21). Lastly, myotubes in the short interference RNA (siRNA) groups were grown in horse serum and were transfected with siRNA for calpain -1 three days prior to harvest or the addition of H2O2. We included siRNA for calpain-1 as anot her means to effectively inhibit calpain activity however siRNA was specific for calpain-1 activity. We have previously shown that 36

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siRNA for calpain-1 reduces calpain-1 mRNA by 65% and active calpain-1 protein levels by 53% in the C2C12 cell line (unpublished observation). Statistical Analysis Group sample size was determined using a power analysis of preliminary data from our laboratory. Comparisons between groups were made by a one-way ANOVA and, when appropriate, a Tukey HSD test was performed. Significance was established at P < 0.05. General Methods Animal Measurements Histological Immunohistochemistry. Diaphragm samples were remove d and fixed in OCT and stored at C. On the day of analysis, sections from frozen diaphragm samples were cut at 10 microns with a cryotome (Shandon Inc., Pittsburgh, PA) and allowed to air dry at 25 C for 30 minutes. Slides were fixed in acetone (4 C) for 5 minutes and allowed to air dry for an additional 30 min at 25C. Slides were wash ed 2 times for 2 minutes in phosphate buffered saline (PBS) and then blocked with normal goat serum (5%) blocking solution for 30 minutes. Sections were incubated in primary xanthine oxidase antibody diluted at 1:50 in TBS for 60 minutes at 25C. This was followed by two washes in PBS buffer for 2 minutes at 25 C. Secondary antibody incubation was done at a con centration of 1:40 (Rhodamine Red goat antirabbit IgG) for 60 min at 25 C in the dark. Final rinses in PB S-T (3x) at 2 minute intervals were performed on all sections. Slides were then mounted in fluorescent mounting medium with Dapi (Vector Laboratories) and images were acquire d via a monochrome camera (Qimaging Retiga) attached to an inverted fluorescent microscope (Axiovert 200, Xeiss). 37

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Myofiber Cross-Sectional Area. Sections from frozen diaphr agm samples were cut at 10 microns using a cryotome (Shandon Inc., Pittsburgh, PA) and stained for dystrophin, myosin heavy chain (MHC) I and MHC type IIa proteins for fiber cross-sec tional area analysis (CSA) as described previously (79). CSA was dete rmined using Scion software (NIH). Functional Contractile Properties. Twitch characteristics and the fo rce-frequency response of a strip of costal diaphragm were performed in vitro and normalized to cross-sectional area (CSA) as previously described (24, 98, 107). Biochemical Xanthine Dehydrogenase and Xa nthine Oxidase Activities. The activities of XDH and XO were assayed using a fluorometric assay according to Beckman et al. (9). The principle of the assay involves the conversion of pterin into the fluorescent product isoxanthopterin. The rate of product formation with oxygen as the electron acceptor represents the activity of XO, whereas the combined activities of XO and XDH are measured with methylene blue as the electron acceptor. In brief, a section of the costal di aphragm was homogenized in a buffer containing 50 mM potassium phosphate, 0.1 mM EDTA, 0.25 M sucrose, and 0.2 mM PMSF (pH 7.4). The homogenates were centrifuged at 40,000 g for 30 min at 4C. The supernatant was collected and assayed immediately. Briefly, XO activity was dete rmined by the addition of 10 M of pterin to 1 mg protein lysate while 10 M of methylene blue was a dded to determine total XO + XDH activity. The conversion of pt erin to the fluorescent product isoxanthopterin was monitored fluorometrically at an excitation of 345 nm and an emission of 390 nm at 37C. Hypoxanthine and Xanthine Levels. Hypoxanthine and xanthine were measured using the Amplex Red xanthine/xanthine oxidase assa y kit from Invitrogen (Eugene, Oregon) as 38

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described by the manufacturer and normalized to protein concentration. Br iefly, a section of the costal diaphragm was homogeni zed 1:30 (wt/vol) in phosphate bu ffered saline (PBS). Samples were centrifuged at 3500 g for 30 min at 4C. After collect ion of the resulting supernatant, diaphragmatic protein content was assessed by th e method of Bradford (Sigma, St. Louis). Approximately fifty microliters of the supernat ant was reacted with the assay cocktail solution for 30 min at 37C in the dark. Absorbance wa s read at 560 nm. Hypoxanthine and xanthine standards were prepared and concentrations were calculated based on the standard curves. Uric Acid Levels. Uric acid levels in the diaphragm were assessed using the uric acid assay kit from BioAssay Systems (Hayward, CA) as described by the manufacturer. Briefly, a section of the costal diaphragm was homogeni zed 1:30 (wt/vol) in phos phate buffered saline (PBS). Samples were centrifuged at 3500 g for 30 min at 4C. After collection of the resulting supernatant, diaphragmatic protein content was assessed by the method of Bradford (Sigma, St. Louis). Approximately twenty microliters of th e supernatant was reacted with the assay cocktail solution for 30 min at room temperature. Absorb ance was read at 590 nm. A uric acid standard was prepared and concentrations were calculated based on this standard. Western Blot Analysis. Protein content was determined in all samples via Western Blot analysis. Diaphragm samples we re homogenized in a buffer cont aining 5 mM Tris (pH 7.5) and 5 mM EDTA (pH 8.0) with a protease inhibitor cocktail (Sigma, St. Louis). Homogenates were centrifuged at 1500 g for 10 min at 4C. After centrifugati on, the supernatant was collected and a protein assay (Bradford) was performed. Protei ns from the supernatant fraction were separated via polyacrylamide gel electrophores is, transferred to a nitrocellu lose membrane, and incubated with primary antibodies directed against the protein of interest. 4-hydroxynonenal (4-HNE) was probed as a measurement indicative of oxidative stress while XOR was performed to visualize 39

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both XDH and XO induction. Membranes were pr obed with Alexa Fluor 680 IgG secondary, scanned, and analyzed using the Odyssey system (Li-Cor Biosciences) using direct infrared fluorescent detection with a wide linear dynamic range. Protein Carbonyls. Protein carbonyls were analyzed as an indicator of protein oxidation. Diaphragmatic protein carbonyls were measured in 40-50 mg total costal diaphragm muscle using a commercially available kit (Zenith T echnology Corporation, Dunedin, NZ) according to the manufacturers instructions. Total Glutathione. Total glutathione content was measured as an indicator of nonenzymatic antioxidant status using a co mmercially available kit according to the manufacturers instructions (Cayman Chemical, Ann Arbor, MI). Briefly, a section of the costal diaphragm was homogenized in 100 mM phospha te buffer containing 1 mM EDTA and 0.05% bovine serum albumin. Sa mples were centrifuged 10,000 g for 15 min at 4C. The supernatant was deproteinated with an equal volume of metaphosphoric acid, centrifuged, and combined with 50 l triethanolamine per milliliter of supernatant. Fifty microliters of deproteinated sample were reacted with the provided assay cocktail in the dark for 25 min. Absorbance was read at 405 nm. Standards were prepared using oxidized glutathione (GSSG), which is reduced to 2 mol equivalents of glutathione (GSH ) under the assay conditions, and c oncentrations were calculated based on this standard curve. Cell Culture Measurements Trypan Blue Exclusion Assay. The trypan blue assay was us ed to assess cellular viability upon exposure to hydrogen peroxide (H2O2). Briefly, an aliquot of cell suspension was diluted 1:1 with 0.4% trypan blue and cells were counted with a hemocytometer. Results are 40

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expressed as the percentage ratio of viable (unstained) cells in hydr ogen peroxide treated samples vs. untreated samples. Xanthine Dehydrogenase and Xa nthine Oxidase Activities. The activities of XDH and XO were assayed using a fluorometric assay according to Beckman et al. (9). Western Blot Analysis. Protein content was determined in all samples via Western Blot analysis. Myotubes were lysed in a buffe r containing 30 mM Tris-HCL (pH 7.5), 250 mM Sucrose, 150 mM NaCl, 1 mM DTT, and protease inhibitor cocktail (Sigma, St. Louis). Lysates were centrifuged at 4 C for 10 minutes at 16000 g. After centrifugation, the supernatant was collected and a protein assay (Bradford) was perf ormed. Proteins from the supernatant fraction were separated via polyacrylamide gel electrophores is, transferred to a nitrocellulose membrane, and incubated with primary antibodies directed against the protein of interest. XOR was probed to visualize both XDH and XO induc tion during the experiments. Calpain activity was assessed by analyzing both total (non-active) and cleaved (active) calpain-1 and total (non-active) calpain2. In addition, -II spectrin was measured to obtain a mo re defined picture of calpain regulation with hydrogen peroxide treatment. Blots were imaged using an Odyssey system (Li-Cor Biosciences), using direct infrared fluorescen t detection with a wi de linear dynamic range. 41

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42 Figure 3-1. Experimental animal design used to determine if pharmacological inhibition of XO activity reduces MV-induced diaphragma tic oxidative stress and contractile dysfunction. Measurements from MV and MVOxypurinol were made vs. CON. We addressed Aim 1 by having two levels of MV at two different durations.

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Figure 3-2. Experimental animal design used to examine whether the administration of an exogenous antioxidant during MV prevents XO activation in the diaphragm. Measurements from MV and MVTrolox were made vs. CON. We addressed Aim 2 by having two levels of MV. 43

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Figure 3-3. Experimental cell cu lture design used to determine if a ROS challenge activates XO in skeletal muscle myotubes. We addre ssed Aim 3 by using hydrogen peroxide to stimulate a ROS challenge to examine th e reversible conversion of XDH to XO in skeletal muscle cells. In addition, leupetin and siRNA for calpain-1 were used in conjunction with hydrogen per oxide to ascertain whethe r calpain is the protease responsible for the irreversib le conversion of XDH to XO in skeletal muscle cells. 44

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CHAPTER 4 RESULTS Experiment 1 Systemic Response to MV To determine if the pharmacological inhibiti on of XO activity in the diaphragm reduces MV-induced oxidative stress and contractile dysf unction, we measured XOR protein and activity levels, key markers of oxidativ e stress, and contractile charac teristics of our experimental animals. Animals used in experiment 1 were 4 months of age a nd weighed between 280-320 g before the experimental procedures. Neither the 12-hr or 18-hr MV protocols altered body weight (P<0.05). Heart rate (HR) systolic blood pressure (SBP), and body temperature (T) were maintained relatively constant during the MV protocols (HR range= 300 beats/min; SBP= 70 mmHg; T= 36-37C). The arte rial partial pressures of O2 (PaO2) and CO2 (PaCO2) were also maintained relatively consta nt during MV. Specifically, PaO2 ranged from 65-100 mmHg whereas PaCO2 ranged from 32-42 mmHg. In addition, at the completion of the MV protocol, there were no visual abnormalities of the lungs or peritoneal cavity, no evidence of lung infarction, and no evidence of infection, indicati ng that our aseptic su rgical technique was successful. XOR Characteristics XOR Localization Xanthine oxidoreductase (XOR) was been characterized in skeletal muscle of both humans and rats (40, 42). Figure 4-1 illustrates that XOR is primar ily localized in the cytosol of rat diaphragm muscle. XOR, stained red, appe ars diffuse throughout the diaphragm muscle fibers as the nuclei, which are stained blue, are localized al ong the sarcolemma. 45

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XDH and XO Activity Levels Under normal conditions, mammalian XOR is synthesized as xanthine dehydrogenase (XDH). XDH uses NAD+ as its electron accepto r in the hydroxylation of hypoxanthine and xanthine to uric aci d. To determine diaphragmatic levels of XDH activity, total XOR activity and XO activity were independently measured in our experimental groups. XDH activity was calculated by subtracting the level of XO activity from total XOR activity. Compared to control, 12 hours of MV resulted in a significant decrease in diaphr agmatic XDH activity (P<0.05) (Table 4-1) and oxypurinol admi nistration restored XDH activity to control values. Prolonged (18 hrs) MV tended to increas e diaphragmatic XDH activity above control (Table 4-1), although this increase did not reach stat istical significance. Nonetheless, 18 hours of MV resulted in a significant increase in total XOR activity (Table 4-1). Given th at XDH activity is determined by the subtraction of XO activity from total XOR activity, as total activity increased in the diaphragm, so did XDH. Oxypurinol administration in the 18-hr MV animals resulted in a significant increase in both XDH and total XOR activ ities above control (Table 4-1). However, when XDH activity is expressed as a percent of total XOR activit y, both MV durations resulted in a decrease in diaphragmatic XDH activity below control and XO i nhibition restored XDH activity to control values. In certain physiological conditions, XDH can be converted to XO. It is in this XO form that during the reduction of the pur ine substrates to uric acid, supe roxide is produced. Compared to control, diaphragmatic XO activity significantl y increased 25% and 43% in the MV animals at 12 and 18 hours, respectively (P<0.05) (Table 4-1). Oxypurinol administration significantly attenuated the increase in diaphragmatic XO activity back to control valu es. When XO activity is expressed as a percent of total XOR activity, MV resulted in a significant increase in 46

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diaphragmatic XO activity. Furthermore, XO inhibition via oxypuri nol administration significantly attenuated the MV-induced increase in diaphragmatic XO activity. Since the level of superoxide production that occurs in skel etal muscle by the XO pathway is dependent upon the levels of XO to XDH in the muscle, we calculated th e ratio of XO to XDH in our experimental groups. Compared to c ontrol, both 12 and 18 hours of MV resulted in a significant increase in XO/XDH activity in the diaphragm (P<0.05) (Figure 4-2). While oxypurinol administration in a group of control animals did not affect the ratio, it significantly attenuated the increase in XO to XDH activity in the diaphragms of our MV animals. Collectively, these results show that MV resu lts in a significant increase in diaphragmatic XO activity and oxypurinol is an e ffective pharmacological agent to inhibit XO activity in the diaphragm during MV. XDH and XO Protein Levels To further understand the ro le of the XO pathway in oxida nt production in the diaphragm during MV, we measured the protei n level of XOR. XOR exists in two alternative forms that are derived from the same gene product (47). Ini tially, XOR is synthesized as a 150 kDa protein from which both XDH and XO are derived (130). Under normal conditions, XOR exists as XDH at a molecular weight of 150 kDa. However, XDH can be reversibly converted to XO via sulfhydryl oxidation which changes the enzyma tic activity of XDH to XO but does not change its weight (101). Compared to control, only prolonged (18 hrs) MV resulted in a significant increase in the 150 kDa band of XOR in the diaphragm (P<0.05) (Figure 4-3). Moreover, compared to control, oxypuri nol administration in both the 12 and 18 hr MV experimental groups resulted in a significant increase in the 150 kDa band of XOR in the diaphragm (P<0.05) 47

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which was not observed in a group of oxypurinol-tre ated control animals. Thus, prolonged MV increases XOR protein (150 kDa) expression in the diaphragm. Whereby both XDH and XO may exist in the diaphragm muscle as a 150 kDa band of XOR, only XO exists as a 130 kDa band of XOR. During activating conditions, XDH can be irreversibly converted to XO via proteolytic m echanisms. The proteolytic processing of XDH results in the cleavage of an approximately 15-20 kDa fragment, resulting in the formation of an active 130 kDa form of XO (22, 130). Compared to control, 12 and 18 hrs of MV resulted in a significant increase in the 130 kDa band of XOR in the diaphragm (P<0.05) (Figure 4-4). In addition, oxypurinol administ ration did not statistica lly alter XO protein levels in the diaphragms of either control or mechanically ventilated anim als. Therefore, both MV durations resulted in a significant increase in the protein levels of XO (130 kDa) in the diaphragm and thus the irreversible proteolytic convers ion of XDH to XO is significan tly increased in the diaphragm during MV. Enzyme Substrates and End Product Hypoxanthine Both XDH and XO catalyze the last two step s of purine catabolism resulting in the formation of uric acid from hypoxanthine and xant hine. Thus an increase in diaphragmatic XO activity would result in the depletion of purine substrates. Compared to control, MV resulted in a significant decrease in hypoxant hine levels in the diaphragm (P<0.05) (Figure 4-5). Twelve hours of MV resulted in an 11% decrease in hypoxanthine and 18 hours resulted in a 38% decrease. Oxypurinol administ ration failed to decrease hypoxan thine levels during prolonged (18 hrs) MV and there was only a trend (P=0.09) for oxypurinol to attenuate the decrease in hypoxanthine levels at 12 hours of MV. The depletion of hypoxanthine during MV coincides 48

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with the MV-induced increase in diaphragmatic XO activity. Finally, XO inhibition did not significantly protect hypoxant hine levels during MV. Xanthine Similar to hypoxanthine, both MV durations re sulted in a significant decrease in xanthine levels in the diaphragm (P<0.01) (Figure 4-6). Twelve hours of MV resulted in a 31% decrease in xanthine, whereas 18 hours resulted in a 41% decrease. XO i nhibition via oxypurinol administration significantly attenuated the decrease in xanthine at both 12 and 18 hours of MV. Thus both MV durations resulted in a significant decrease in the le vel of purine substrates in the diaphragm. However, unlike hypoxanthine, oxy purinol administration significantly attenuated the MV-induced decrease in xant hine levels in the diaphragm. Uric Acid In the presence of the purine substrat es and molecular oxygen, XO catalyzes the production of uric acid and superoxide. Accordi ngly, the level of uric acid formation in the diaphragm would increase in concomitant with an increase in diaphragmatic XO activity. Compared to control, 12 and 18 hrs of MV resulted in a significant 15% and 42% increase in uric acid formation in the diaphragm, respectiv ely (P<0.05) (Figure 4-7). Although oxypurinol administration tended to attenuate the increase in uric acid at the 12 hour time point, it did not reach significance (P=0.09). In contrast, oxypurinol administration significantly attenuated the formation of uric acid in the diaphragms of our 18-hr MV animals. Thus MV results in a significant increase in uric ac id, an end product of both XDH a nd XO enzymatic activity, in the diaphragm. This coincides with the MV-indu ced increase in diaphragmatic XO activity and depletion of the purine subs trates in the diaphragm. 49

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Redox Balance Protein Carbonyls Since MV results in an increase in dia phragmatic XO activity, the potential for ROS production by the XO pathway exists in the diaphr agm. Protein carbonyl formation is a general indicator of protein oxidation a nd commonly used to measure oxida tive injury. Twelve hours of MV resulted in a sign ificant increase in prot ein carbonyl formation in the diaphragm when compared with control (P<0.05) (Figure 4-8) and there was a tre nd (P=0.07) for oxypurinol administration to attenuate this in crease. Prolonged (18 hrs) MV further elev ated the levels of protein carbonyl formation in the diaphragm wh en compared with 12 hrs of MV (P<0.05). While oxypurinol administration si gnificantly attenuated the increas e in protein carbonyls at the 18-hr time point, protein carbonyl formation in the diaphragm remained significantly elevated when compared with control. Thus, XO inhib ition attenuates some of the MV-induced protein oxidation in the diaphragm 4-HNE Lipid peroxidation occurs as a response to oxidative stress, and among the end-products it produces several biologically active aldehydes. 4-hydroxynoneal (4-HNE) is an unsaturated hydroxyalkenal that is generated during the lipid peroxidation cascade. Furthermore, 4-HNE is the primary adduct formed during th is process and is commonly us ed to assess protein oxidative damage. Compared to control, both MV dur ations resulted in a significant increase in diaphragmatic 4-HNE (P<0.05) (Figure 4-9). XO inhibition significa ntly attenuated the accumulation of 4-HNE as oxypurinol treated animal s had basal levels of diaphragmatic 4-HNE. Therefore, diaphragmatic XO activity contributes to MV-induced oxidative stress by the fact that 50

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XO inhibition reduces protein oxidation and signi ficantly attenuates lipid peroxidation in the diaphragm. Total Glutathione Glutathione (GSH) is the major nonezymatic antioxidant component of the cell and depletion of cellular stores is generally considered indicative of oxidative stress. Following 12 and 18 hours of MV, there was a significant depl etion of total GSH in the diaphragm (P<0.05) (Figure 4-10) and oxypurinol administration failed to attenuate this decrease. Consequently, MV results in the depletion of the antioxidant defense of the diaphr agm and XO inhibition does not rescue this depletion. Contractile Function Maximal Isometric Twitch Force To determine if XO-mediated ROS produc tion in the diaphragm contributes to diaphragmatic contractile dysfunction, we m easured maximal isometric twitch force and analyzed the force-frequency response of dia phragm strips obtained from our experimental animals. Maximal isometric twitch forces of in vitro costal diaphragm strips from mechanically ventilated and nonmechanically ventilated (control) animals with and without oxypurinol are presented in Table 4-2. Maxima l isometric twitch force was si gnificantly reduced following 18 hrs of MV when compared with control and cont rol with oxypurinol. During prolonged (18 hrs) MV, there was a trend (P=0.07) for XO inhibition via oxypurinol administration to attenuate the drop in maximal isometric twitch force. Accord ingly, prolonged MV results in a significant decrease in maximal isometric twitch force of the diaphragm and oxypurinol administration during MV appears to restor e some of this decrease. 51

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Force-Frequency Response Mean maximal force-frequency responses of in vitro costal diaphragm strips from mechanically ventilated and nonmechanically ve ntilated (control) anim als with and without oxypurinol are presented in Fi gure 4-11. Both 12 and 18 hours of MV shifted the forcefrequency response down when compared with c ontrol (CON) and control animals treated with oxypurinol (CONO). Twelve and ei ghteen hours of MV resulted in a significant reduction in the specific force of the diaphragm compared to the control groups at all stim ulation frequencies. Specifically, both MV durations resulted in a 23 % decrease in maximal di aphragmatic specific force. Treatment with oxypurinol prior to MV (12MVO and 18MVO) attenuated the MVinduced diaphragmatic dysfunction at stimulation frequencies above 60 hertz. So XO inhibition attenuated some of the MV-induced contra ctile dysfunction that is normally observed. Diaphragmatic Atrophy Cross-sectional Area To evaluate the impact of XO inhibiti on on MV-induced diaphragmatic atrophy, we compared the cross-sectional areas (CSA) of di aphragmatic myofibers across our experimental groups. Compared to control, we observed a si gnificant decrease (P<0.05) in diaphragmatic type I fiber CSA following 12 and 18 hours of MV (Figur e 4-12). While oxypuri nol administration at both MV time points appears to attenuate the decrease in type I my ofiber cross-sectional area, it did not reach statistical signifi cance (P=0.1). Diaphragmatic t ype IIa fiber CSA followed the same pattern as the type I fibers, whereby bot h 12 and 18 hours of MV re sulted in a significant reduction in type IIa fiber CSA in the diaphr agm (P<0.05) (Figure 4-13). There was a trend (P=0.07) for oxypurinol administra tion to attenuate the decrease in type IIa CSA following 12 hours of MV. Finally, significant decreases in CSA for type IIb/x fibers were observed in 52

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diaphragms from both MV groups (Figure 4-14) although no significant di fferences existed in fiber CSA between control and oxypurinol tr eated MV animals. Therefore, while XO contributes to both MV-induced oxidative stress and contractile dysfunction, XO inhibition does not appear to attenuate diaphragmatic atrophy during MV. Experiment 2 Systemic Response to MV To ascertain whether the administration of an exogenous antioxidant during MV prevents diaphragmatic XO activation, we administered Trol ox to a subset of MV animals and measured oxidative stress, contractile func tion, and XOR protein and activity levels in the diaphragms of our experimental animals in expe riment 2. All of the animals were 4 months of age. No significant differences existed in body we ight between the groups (CON = 0.305 g 0.005, 12MV = 0.294 g 0.003, and 12MVT = 0.298 g 0.004) before the experiment, and the 12-hr experimental protocol did not alter body wei ght (P<0.05). Heart rate (HR), systolic blood pressure (SBP), and body temperature (T) were ma intained within a normal physiological range during the MV protocol (HR range= 300 b eats/min; SBP= 70 mmHg; T= 36-37C). The arterial partial pressures of O2 (PaO2) and CO2 (PaCO2) were also maintained relatively constant during MV. Specifically, PaO2 ranged from 60-89 mmHg whereas PaCO2 ranged from 34-46 mmHg. In addition, at the completion of the MV protocol, there were no visual abnormalities of the lungs or peritoneal cavity, no evidence of lung infarction, and no evidence of infection, indicating that our aseptic surgical technique was successful. Redox Balance Protein Carbonyls In experiment 1, we demonstrated that the irreversible proteolytic conversion of XDH to XO is increased in the diaphragm during MV to induce XO activity. This proteolytic conversion 53

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is thought to occur subsequent to the reversible conversi on of XDH to XO via sulfhydryl oxidation of cysteine residues (14, 83). To as certain whether the reversible conversion via oxidation is a required step by which diaphrag matic XO activity is increased during MV, we used Trolox to attenuate the oxida tive stress that occurs during ventilation. By attenuating the MV-induced diaphragmatic oxidative stress, we theorized that any sulfhydrl oxidation of cysteine residues on XDH would be eliminated. To demonstrate the effectiveness of Trolox as an appropriate pharmacological agent to restor e diaphragmatic redox balance, protein carbonyl formation and 4-HNE levels were measured. Twelve hours of MV resulted in a significant increase in protein carbonyl formation in the di aphragm (P<0.05) when compared with control (Figure 4-15). More importantly, Trolox administ ration significantly attenuated the increase in protein carbonyls in the diaphragm. 4-HNE Compared to control, 12 hours of MV resulted in a significant increas e in diaphragmatic 4HNE (P<0.05) (Figure 4-16). Tr olox administration significantly attenuated the accumulation of 4-HNE as Trolox treated animals had basal levels of diaphragma tic 4-HNE. Thus we have shown that Trolox administration during MV effectively inhibits diaphragmatic oxidative stress by the complete attenuation of protein oxidation and lipid peroxidation in the diaphragm of Trolox-treated MV animals. Contractile Function Maximal Isometric Twitch Force Maximal isometric twitch force and force-fr equency responses were measured in our experimental groups to further demonstrate th e effectiveness of Trolox in maintaining redox balance. Maximal isometric twitch forces of in vitro costal diaphragm strips from control, 54

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mechanically ventilated, and mechanically ventilat ed animals with Trolox are presented in Table 4-3. Maximal isometric twitch force was signi ficantly reduced 30% following 12 hrs of MV when compared with controls (P<0.05) and Trol ox administration significantly attenuated the drop in diaphragmatic maximal isometric twitch force. Force-Frequency Response The mean maximal force-frequency responses of in vitro costal diaphragm strips from control, mechanically ventilate d, and mechanically ventilated an imals with Trolox are presented in Figure 4-17. Twelve hours of MV shifted the force-frequency response down when compared with control. In fact, 12 hours of MV resulted in a significant re duction (P<0.05) in the specific force of the diaphragm compared to the control group at all stimulation frequencies. Treatment with Trolox during MV significan tly attenuated the MV-induce d diaphragmatic contractile dysfunction at all stimulation frequencies. Th erefore, Trolox administration is a potentially useful therapeutic approach to attenuate MV-induced diaphr agmatic contractile dysfunction. XOR Characteristics XDH and XO Activity Levels To examine the reversible conversion of XDH to XO in the diaphragm during MV, we measured total XOR, XDH, and XO activities in the diaphragms of our control, mechanically ventilated, and mechanically ventilated animals with Trolox. If the re versible conversion of XDH to XO is present in the diaphragm during MV, then we would expect to find an increase in diaphragmatic XDH activity along with a concomita nt decrease in XO activity in our Troloxtreated MV animals when compared with our regular MV animals. Compared to control, MV resulted in a significant decr ease in diaphragmatic XDH activ ity (P<0.05) (Figure 4-18) along with an increase in XO activity (P<0.05) (Figure 4-19). Trolox ad ministration failed to attenuate the changes in both XDH and XO activity in the diaphragm. Because Trolox administration 55

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during MV did not attenuate dia phragmatic XO activity, it appears that the revers ible conversion of XDH to XO via oxidation does not occur in th e diaphragm during MV when oxidative stress in prevented with Trolox. XDH and XO Protein Levels Because the reversible conversion only ch anges enzymatic activity and not molecular weight, Trolox administration du ring MV should not alter the pr otein expression of the 150 kDa band of XOR or directly change the 130 kDa ba nd produced via the irreversible proteolytic process. No significant differe nces existed in the 150 kDa band of XOR (Figure 4-20) in the diaphragm between our experimental groups. In addition, Trolox administration did not attenuate the MV-induced induction of the 130 kDa band of XOR in the diaphragm (P<0.05) (Figure 4-21). Enzyme Substrates and End Product Given that Trolox administration during MV failed to alter diaphragmatic XO activity, purine substrate and uric acid leve ls would not be expected to ch ange in our MV animals treated with Trolox. The levels of hypoxanthine, xanthine, and uric aci d for control, mechanically ventilated, and mechanically ventilated anim als with Trolox are presented in Table 4-4. Compared to control, 12 hours of MV resulted in a significant decrea se in both hypoxanthine (~16%) and xanthine (~19%) in the diaphrag m (P<0.05) and Trolox administration did not rescue these levels. With regards to uric acid, co mpared to control, 12 hours of MV resulted in a significant 17% increase in uric acid formati on in the diaphragm (P<0.05). Again, Trolox administration failed to attenuate this increase in the diaphragm during MV. The fact that Trolox administration failed to alter the MV-induced changes in the levels of diaphragmatic hypoxanthine, xanthine, and uric acid, further demons trate that the revers ible conversion of XDH 56

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to XO via oxidation does not occur in the di aphragm during MV when redox balance is maintained with Trolox. Experiment 3 Myogenic Cells Cell Viability To determine if a ROS challenge activates XO in skeletal muscle myotubes, we used the C2C12 myogenic cell line in our third experiment. We performed cell viability measurements on C2C12 myotubes treated with and wi thout hydrogen peroxide (100 M) for 4 hours. These measurements were performed to determine if the hydrogen peroxide treatment was toxic to our cell line. Importantly, our results revealed that hydrogen peroxide treatment was not toxic to myocytes in our experimental model as indicated by the failure of myotubes to take up Trypan blue (97% viable). XDH and XO Activity Levels To investigate the mechanism(s) responsible for XO activation, we utilized a myogenic cell line to determine the effect of a ROS challenge (H2O2) on XO activation in skeletal muscle myotubes. Both XDH and XO activities were assesse d in our cultured myot ubes to determine if there was a similar increase in XO activ ity in cell culture compared to the in vivo animal experiments. In doing so, we examined the i rreversible conversion of XDH to XO in skeletal muscle cells. Hydrogen peroxide was used to mediate the oxidation of the cysteine residues on XDH and both leupeptin and short interference RNA (siRNA) for calpa in-1 were used to inhibit the activity of calpain which is up-regulated with hydrogen peroxi de treatment. Calpain was examined as the protease responsible for the irreversible proteolytic processing of XDH to XO. 57

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There were no significant di fferences in XDH activity in our experimental cell groups treated with or without hydrogen peroxide (Figure 4-22). However, there was a 21% increase in XO activity in cells treated with hydrogen peroxi de (P<0.05) (Figure 4-23). Moreover, siRNA for calpain-1 failed to attenuate the hydroge n peroxide-induced increase in XO activity in skeletal muscle myotubes. However, hydrogen peroxide-induced XO activation was significantly attenuated with leupeptin. So while it appear s that hydrogen per oxide induction of XO activity in C2C12s involves calpain activity or anothe r protease inhibited by leupeptin, it appears that calpain-1 is not i nvolved in the increase in XO activ ity in skeletal muscle myotubes with a hydrogen peroxide challenge. XDH and XO Protein Levels Measurements were also made to assess XOR protein content in our cultured myotubes to determine if there was a similar induction of XOR in cell culture compared to the in vivo animal experiments. There were no significant differences between our cell treatment groups in the protein expression of the 150 kDa band of XOR (F igure 4-24). However, when compared to control, hydrogen peroxide treated cells exhibited an approximate 1.4 fold increase in the 130 kDa band of XOR (Figure 4-25). The increase in the 130 kDa band of XOR was also observed in cells treated with both hydr ogen peroxide and siRNA for ca lpain-1, although not when cells were treated with hydrogen peroxide and leupeptin or when cells were treated with leupeptin or siRNA for calpain-1 alone. The observation th at compared to hydrogen peroxide treatment alone, cells treated with hydroge n peroxide and siRNA for calpain-1 exhibited the same significant increase in the 130 kD a band of XOR and cells treated with hydrogen peroxide and leupeptin prevented the induction of the 130 kD a band, suggests that a calpain protease is involved in XO activation in skel etal muscle myotubes but specifi cally eliminates calpain-1 as 58

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the protease responsible for the irreversible proteolytic conversion of XDH to XO in skeletal muscle cells. Calpain Protein and Activity To demonstrate the effectiveness of both leupe ptin and siRNA to i nhibit calpain activity, measurements for calpain protein abundance and activity were made in our experimental cell groups. Compared to control, leupeptin treated cells exhibited a signific ant reduction in total calpain-1 protein (Figure 4-26). There was a trend for both si RNA for calpain-1 (P=0.08) and hydrogen peroxide plus siRNA for calpain-1 (P =0.06) to reduce tota l calpain-1 protein expression in skeletal muscle myotubes. As exp ected, compared to control, cells treated with hydrogen peroxide resulted in a significant increase in the cleaved or active band of calpain-1 (Figure 4-27). However, cells tr eated with either leupeptin or siRNA for calpain-1 resulted in a significant reduction in cleaved cal pain-1. Furthermore, cleaved calpain-1 remained depressed below control values when cells where treated wi th both hydrogen peroxide and either inhibitor. Finally, there were no significant differences between our cell treatment groups in the protein expression of total calpain-2 (Figure 4-28). Lastly, to further demonstrate the effectiv eness of our calpain i nhibitors, the calpain specific degradation product of -II spectrin was measured. The intact form of -II spectrin exists as ~250 kDa protein and upon degradation yields a 145 kDa (calpain-specific) cleaved band. Thus, in vivo calpain activity can be determined by probing for the calpain-specific -II spectrin cleaved degradation produ ct. There were no significant differences between our cell treatment groups in the protein expression of total -II spectrin (Figure 4-29). Compared to control, cells treated with hydrogen peroxide resulted in a signi ficant 1.5 fold induction of the calpain-specific cleaved -II spectrin band (145 kDa) (Figure 4-30). On the other hand, there 59

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60 were no increases in the calpain-specific cleaved -II spectrin degradation product when cells were treated with either leupeptin, hydrogen peroxide and leup eptin, siRNA for calpain-1, or hydrogen peroxide and siRNA for calpain-1. Thus, leupeptin and siRNA for calpain-1 were completely effective in preventing the incr ease in calpain activity observed with hydrogen peroxide treatment.

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Table 4-1. Total xanthine oxidor eductase (XOR), xanthine dehydr ogenase (XDH), and xanthine oxida se (XO) activities in diaphrag m from control, mechanically ventilated, and m echanically ventilated animals with oxypurinol. Activity (mol/min/g) CON CONO 12MV 12MVO 18MV 18MVO Total XOR 0.131 0.004 0.148 0.004 0.126 0.004 0.126 0.004 0.165 0.007* 0.183 0.008*# XDH 0.075 0.006 0.088 0.010 0.056 0.003*# 0.078 0.002 0.085 0.008 0.126 0.013*# (57%) (59%) (44%) (62%) (52%) (69%) XO 0.056 0.003 0.060 0.005 0.070 0.003* 0.048 0.005 0.080 0.005*# 0.057 0.006 (43%) (41%) (56%) (38%) (48%) (31%) Values are expressed as mean SE. The values in parent heses represent the percent activity of total XOR activity. *Significantly different versus CON (P<0.05). #Significantly different from CONO (P<0.05). Significantly different from 12MV (P<0.05). Significantly different from 12MVO. Significantly different from 18MV (P<0.05) CON= Control; CONO= Control with oxypurinol; 12MV= 12 hrs mechanical ventil ation; 12MVO= 12MV with oxypurinol; 18MV= 18 hrs mechanical ventilation; 18MVO= 18MV with oxypurinol.61

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Table 4-2. Maximal isometric twitch for ce production in diaphragm strips obtained from mechanically ventilated and nonmechanically ventilated (control) animals with and without oxypurinol. Experimental Group Maximal Isometric Twitch Force, N/cm2 Control 10.1 0.3 Control with oxypurinol 10.1 0.1 18 h of MV 6.3 0.4*# 18 h of MV with oxypurinol 8.2 0.8 Values are expressed as mean SE. *Significantly decreased ve rsus Control (P<0.001). #Significantly decreased versus Control with oxypurinol (P<0.001).62

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CON(+,+) CON(+,-)Figure 4-1. Xanthine oxidoreductase (XOR) localization in a control diaphragm muscle sample. XOR appears to be localized in the cytosol. Red stain= XOR and blue stain= nuclei. (+,+); Both primary and secondary anti body application; (+,-); Primary antibody application alone. Figure 4-2. Ratio of xanthine oxidase (XO) activity to xanthi ne dehydrogenase (XDH) activity in diaphragm samples from experiment 1. Values are mean fold change SE. *Significantly increased versus CON (P<0.05). #Significantly increased versus CONO (P<0.05). Significantly decreased versus 12MV (P<0.001). Significantly decreased versus 18MV (P<0.01). CON= Control; CONO= Control with oxypurinol; 12MV= 12 hrs mechanical ventilation; 12MVO= 12MV with oxypurinol; 18MV= 18 hrs mechanical ventilation; 18MVO= 18MV with oxypurinol. 63

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Figure 4-3. Fold changes (versu s control) of XOR protein (150 kDa band) content in diaphragm samples from experiment 1. Values are mean fold change SE. *Significantly increased versus CON (P<0.01). #Significantly increased versus CONO (P<0.05). Significantly increased versus 12MV (P<0.05). CON= Control; CONO= Control with oxypurinol; 12MV= 12 hrs mechanical ventilation; 12MVO= 12MV with oxypurinol; 18MV= 18 hrs mechanical ven tilation; 18MVO= 18MV with oxypurinol. Figure 4-4. Fold changes (versus control) of XOR protein (130 kDa) content in diaphragm samples from experiment 1. Values are mean fold change SE. *Significantly increased versus CON (P<0.05). #Significantly increased versus CONO (P<0.05). Significantly increased versus 12MVO (P <0.05). CON= Control; CONO= Control with oxypurinol; 12MV= 12 hrs mechanical ventilation; 12MVO= 12MV with oxypurinol; 18MV= 18 hrs mechanical ven tilation; 18MVO= 18MV with oxypurinol. 64

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Figure 4-5. Hypoxanthine levels in diaphragm samples from experiment 1. Values are mean SE. *Significantly decreased versus CON (P<0.05). #Significantly different versus CONO (P<0.05). Significantly decreased versus 12MV (P<0.01). Significantly decreased versus 12MVO (P<0.001). CON= Control; CONO= Control with oxypurinol; 12MV= 12 hrs mechanical ven tilation; 12MVO= 12MV with oxypurinol; 18MV= 18 hrs mechanical ventilation; 18MVO= 18MV with oxypurinol. Figure 4-6. Xanthine levels in diaphragm samples from experiment 1. Values are mean SE. *Significantly decreased versus CON (P<0.01). #Significantly decreased versus CONO (P<0.001). Significantly increased versus 12MV (P<0.01). Significantly decreased versus 12MVO (P<0.001). Significantly increased versus 18MV (P<0.001). CON= Control; CONO= Control with oxypurinol; 12MV= 12 hrs mechanical ventilation; 12MVO= 12MV w ith oxypurinol; 18MV= 18 hrs mechanical ventilation; 18MVO= 18MV with oxypurinol. 65

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Figure 4-7. Uric acid levels in diaphragm samp les from experiment 1. Values are mean SE. *Significantly increased versus CON (P<0.05). #Significantly increased versus CONO (P<0.01). Significantly increased versus 12MV (P<0.05). Significantly increased versus 12MVO (P<0.01). Significantly decreased versus 18MV (P<0.01). CON= Control; CONO= Control with oxypurinol; 12M V= 12 hrs mechanical ventilation; 12MVO= 12MV with oxypurinol; 18MV= 18 hrs mechanical ventilation; 18MVO= 18MV with oxypurinol. Figure 4-8. Protein carbonyl levels in diaphragm samples from experiment 1. Values are mean SE. *Significantly increased versus CON (P<0.05). #Significantly increased versus CONO (P<0.05). Significantly increased versus 12MV (P<0.05). Significantly increased versus 12MVO (P<0.01). Significantly decreased versus 18MV (P<0.05). CON= Control; CONO= Control with oxypurinol; 12MV= 12 hrs mechanical ventilation; 12MVO= 12MV with oxypurinol; 18MV= 18 hrs mechanical ventilation; 18MVO= 18MV with oxypurinol. 66

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Figure 4-9. Fold changes (versus contro l) of 4-hydroxynonenal (4-HNE) accumulation in diaphragm samples from experiment 1. Values are mean fold change SE. *Significantly increased versus CON (P<0.05). Significantly decreased versus 12MV (P<0.05). Significantly decreased versus 18M V (P<0.05). CON= Control; CONO= Control with oxypurinol; 12MV= 12 hrs m echanical ventilation; 12MVO= 12MV with oxypurinol; 18MV= 18 hrs mechanical ventilation; 18MVO= 18MV with oxypurinol. Figure 4-10. Total glutathione co ncentrations in diaphragm samp les from experiment 1. Values are mean SE. *Significantly decreased versus CON (P<0.05). #Significantly decreased versus CONO (P<0.01). Significantly decreased versus 12MV (P<0.05). CON= Control; CONO= Control with oxypurinol; 12MV= 12 hrs mechanical ventilation; 12MVO= 12MV with oxypurinol; 18MV= 18 hrs mechanical ventilation; 18MVO= 18MV with oxypurinol. 67

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5 10 15 20 25 3015 Hz 30 Hz 60 Hz 100 Hz 160 Hz Stimulation Frequency (Hz)Specific Force (N/cm2) CON CONO 12MVO 18MVO 12MV 18MV # # Figure 4-11. Diaphragmatic force-frequency response ( in vitro) of diaphragm samples from experiment 1. Values are expressed as mean SE. *12MV, 18MV, 12MVO, and 18MVO significantly decreased versus CON and CONO (P<0.01). #Significantly decreased versus CON and CONO, except 12MVO and 18MVO (P<0.05). 18MV significantly decreased versus 12MVO and 18MVO (P<0.05). CON= Control; CONO= Control with oxypurinol; 12MV= 12 hrs mechanical ventilation; 12MVO= 12MV with oxypurinol; 18MV= 18 hrs mechan ical ventilation; 18MVO= 18MV with oxypurinol. Figure 4-12. Type I fiber cross-sectional area in diaphragm samples from experiment 1. Values are mean SE. *Significantly decreased versus CON (P<0.05). #Significantly decreased versus CONO (P<0.05). CON= Control; CONO= Control with oxypurinol; 12MV= 12 hrs mechanical ventilation; 12MVO= 12MV with oxypurinol; 18MV= 18 hrs mechanical ventilation; 18MVO= 18MV with oxypurinol. 68

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69 Figure 4-13. Type IIa fiber cr oss-sectional area in diaphrag m samples from experiment 1. Values are mean SE. *Significantly decreased versus CON (P<0.05). #Significantly decreased versus CONO (P<0.05). CON= Control; CONO= Control with oxypurinol; 12MV= 12 hrs mechanical ventilation; 12MVO= 12MV with oxypurinol; 18MV= 18 hrs mechanical ventilation; 18MVO= 18MV with oxypurinol. Figure 4-14. Type IIb/x fiber cross-sectional area in diaphragm samples from experiment 1. Values are mean SE. *Significantly decreased versus CON (P<0.05). #Significantly decreased versus CONO (P<0.05). Significantly decreased versus 12MVO (P<0.05). CON= Control; CONO= Control with oxypurinol; 12MV= 12 hrs mechanical ventilation; 12MVO= 12MV with oxypurinol; 18MV= 18 hrs mechanical ventilation; 18MVO= 18MV with oxypurinol.

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Table 4-3. Maximal isometric twitch force pr oduction in diaphragm strips obtained from control, mechanically ventilated, and mechanically ventilated animals with Trolox. Experimental Group Maximal Isometric Twitch Force, N/cm2 Control 9.55 0.28 12 h of MV 6.73 0.35*# 12 h of MV with Trolox 9.31 0.85 Values are expressed as mean SE. *Significantly decreased ve rsus Control (P<0.05). #Significantly decrease d versus 12 h of MV with Trolox (P<0.05).70

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Table 4-4. Hypoxanthine, xanthine, and uric acid levels in contro l, mechanically ventilated, and mechanically ventilated anima ls with Trolox. Measurement (uM/mg protein) CON 12MV 12MVT Hypoxanthine 10.67 0.36 9.01 0.20* 9.29 0.21* Xanthine 24.43 1.02 19.85 1.09* 19.45 0.47* Uric Acid 24.67 1.15 28.90 1.43* 29.81 0.72* Values are expressed as mean SE. *Significantly different versus CON (P<0.05). CON= Control; 12MV= 12 hrs of mechanical ventilation; 12MVT= 12MV with Trolox.71

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Figure 4-15. Protein carbonyl leve ls in diaphragm samples from experiment 2. Values are mean SE. *Significantly increased versus CON (P<0.05). Significantly decreased versus 12MV (P<0.05). CON= Control; 12MV= 12 hr s mechanical ventilation; 12MVT= 12MV with Trolox. Figure 4-16. Fold changes (versus control) of 4hydroxynonenal (4-HNE) accumulation in diaphragm samples from experiment 2. Values are mean fold change SE. *Significantly increased versus CON (P<0.05). Significantly decreased versus 12MV (P<0.05). CON= Control; 12MV= 12 hrs mechanical ventilation; 12MVT= 12MV with Trolox. 72

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5 10 15 20 25 30 15 Hz 30 Hz 60 Hz 100 Hz 160 Hz Stimulation Frequency (Hz)Specific Force (N/cm2) CON 12MVT 12MV Figure 4-17. Diaphragmatic force-frequency response ( in vitro) of diaphragm samples from experiment 2. Values are expressed as mean SE. *Significantly decreased versus CON and 12MVT (P<0.001). CON= Control; 12 MV= 12 hrs mechanical ventilation; 12MVT= 12MV with Trolox. Figure 4-18. Xanthine dehydrogena se (XDH) activity in diaphragm samples from experiment 2. Values are mean fold change SE. *Significantly decreased versus CON (P<0.05). CON= Control; 12MV= 12 hrs mechanical ventilation; 12MVT= 12MV with Trolox. 73

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Figure 4-19. Xanthine oxidase (XO) activity in diaphragm samples from experiment 2. Values are mean fold change SE. *Significantly increased vers us CON (P<0.05). CON= Control; 12MV= 12 hrs mechanical ve ntilation; 12MVT= 12MV with Trolox. Figure 4-20. Fold changes (versus control) of XOR protein (150 kDa band) content in diaphragm samples from experiment 2. Values are mean fold change SE. CON= Control; 12MV= 12 hrs mechanical ve ntilation; 12MVT= 12MV with Trolox. 74

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Figure 4-21. Fold changes (versus control) of XOR protein (130 kDa band) content in diaphragm samples from experiment 2. Values are mean fold change SE. *Significantly increased versus CON (P <0.01). CON= Control; 12MV= 12 hrs mechanical ventilation; 12MVT= 12MV with Trolox. 75

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Figure 4-22. Xanthine dehydrogenase (XDH) activity in C2C12 myotubes from experiment 3. Values are mean fold change SE. CON= Control; H2O2= Hydrogen peroxide; LEU= Leupeptin; H2O2 + LEU= Hydrogen peroxide and leupeptin; Cal-1 siRNA= Calpain-1 siRNA; H2O2 + Cal-1 siRNA= Hydrogen peroxide and Calpain-1 siRNA. 76

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Figure 4-23. Xanthine ox idase (XO) activity in C2C12 myotubes from experiment 3. Values are mean fold change SE. *Significantly increased versus CON (P<0.05). #Significantly decreased versus H2O2 (P<0.05). Significantly increased versus H2O2 + LEU (P<0.001). CON= Control; H2O2= Hydrogen peroxide; LEU= Leupeptin; H2O2 + LEU= Hydrogen peroxide and leupeptin; Cal-1 siRNA= Calpain-1 siRNA; H2O2 + Cal-1 siRNA= Hydrogen peroxide and Calpain-1 siRNA. 77

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Figure 4-24. Fold change (versus contro l) of XOR protein (150 kDa band) in C2C12 myotubes from experiment 3. Values are mean fold change SE. CON= Control; H2O2= Hydrogen peroxide; LEU= Leupeptin; H2O2 + LEU= Hydrogen peroxide and leupeptin; Cal-1 siRNA= Calpain-1 siRNA; H2O2 + Cal-1 siRNA= Hydrogen peroxide and Calpain-1 siRNA. 78

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Figure 4-25. Fold change (versus contro l) of XOR protein (130 kDa band) in C2C12 myotubes from experiment 3. Values are mean fold change SE. *Significantly increased versus CON (P<0.05). #Significantly decreased versus H2O2 (P< 0.05). Significantly decreased versus H2O2 + Cal-1 siRNA (P<0.05) CON= Control; H2O2= Hydrogen peroxide; LEU= Leupeptin; H2O2 + LEU= Hydrogen peroxide and leupeptin; Cal-1 siRNA= Calpain-1 siRNA; H2O2 + Cal-1 siRNA= Hydrogen peroxide and Calpain-1 siRNA. 79

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Figure 4-26. Fold change (versus control) of total calpain-1 protein in C2C12 myotubes from experiment 3. Values are mean fold change SE. *Significantly decreased versus CON (P<0.05). CON= Control; H2O2= Hydrogen peroxide; LEU= Leupeptin; H2O2 + LEU= Hydrogen peroxide and leupeptin; Cal-1 siRNA= Calpain-1 siRNA; H2O2 + Cal-1 siRNA= Hydrogen peroxide and Calpain-1 siRNA. 80

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Figure 4-27. Fold change (versus contro l) of cleaved calpain-1 protein in C2C12 myotubes from experiment 3. Values are mean fold change SE. *Significantly differe nt versus CON (P<0.05). #Significantly decreased versus H2O2 (P<0.05). CON= Control; H2O2= Hydrogen peroxide; LEU= Leupeptin; H2O2 + LEU= Hydrogen peroxide and leupeptin; Cal-1 siRNA= Calpain-1 siRNA; H2O2 + Cal-1 siRNA= Hydrogen peroxide and Calpain-1 siRNA. 81

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Figure 4-28. Fold change (versus control) of total calpain-2 protein in C2C12 myotubes from experiment 3. Values are mean fold change SE. CON= Control; H2O2= Hydrogen peroxide; LEU= Leupeptin; H2O2 + LEU= Hydrogen peroxide and leupeptin; Cal-1 siRNA= Calpain-1 siRNA; H2O2 + Cal-1 siRNA= Hydrogen peroxide and Calpain-1 siRNA. 82

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Figure 4-29. Fold change (versus control) of total -II spectrin protein (250 kDa) in C2C12 myotubes from experiment 3. Values are m ean fold change SE. CON= Control; H2O2= Hydrogen peroxide; LEU= Leupeptin; H2O2 + LEU= Hydrogen peroxide and leupeptin; Cal-1 siRNA= Calpain-1 siRNA; H2O2 + Cal-1 siRNA= Hydrogen peroxide and Calpain-1 siRNA. 83

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84 Figure 4-30. Fold change (versus control) of calpain-specific cleaved -II spectrin protein (145 kDa) in C2C12 myotubes from experiment 3. Values are mean fold change SE. *Significantly increased versus CON (P<0.01). #Significantly decreased versus H2O2 (P<0.01). CON= Control; H2O2= Hydrogen peroxide; LEU= Leupeptin; H2O2 + LEU= Hydrogen peroxide and leupeptin; Cal-1 siRNA= Calpain-1 siRNA; H2O2 + Cal-1 siRNA= Hydrogen peroxide and Calpain-1 siRNA.

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CHAPTER 5 DISCUSSION Overview of Principal Findings These experiments provide new and important information regarding the impact of the XO pathway on oxidant production in the diaphr agm during prolonged MV. We tested the hypothesis that XO inhibition would attenuate MV -induced diaphragmatic oxidative stress and contractile dysfunction. Our findings support this postulate as MV resulted in the up-regulation of diaphragmatic XO activity and XO inhibition attenuated some of the MV-induced oxidative stress and contractile dysfunction that is normally observed in the diaphragm. We also predicted that the administration of an exogenous antioxida nt during MV would main tain redox balance in the diaphragm and retard XO activation via the re versible conversion of XDH to XO. However, our results did not support this postulate as Trolox administrati on failed to a ttenuate the MVinduced increase in XO activity and uric acid formation in the diaphragm. Finally, to investigate the activation mechanism(s) of XO in skeletal muscle, we utilized a myogenic cell line to examine the effect of a ROS challenge on XO act ivation in skeletal muscle myotubes. Our results indicate that hydrogen peroxide treatment of C2C12s results in the irreversible proteolytic conversion of XDH to XO. Furthermore, calpain-1 specifically does not play a role in the irreversible proteolytic convers ion of XDH to XO in skeletal muscle myotubes. A detailed discussion providing an interpreta tion of our experiments follows in the subsequent sections. Mechanical Ventilation-induced Induction of XO Activity The xanthine oxidoreductase (XOR) family, which consists of xanthine dehydrogenase (XDH) and xanthine oxidase (XO), is present in almost all mammalian tissues. The enzyme catalyzes the breakdown of hypoxanthi ne and xanthine to urate. Importantly, the oxidase form of XOR, xanthine oxidase, produces superoxide in the catabolism of the purine substrates to uric 85

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acid. Therefore, the XO pathway is one of the major sources of free radicals in biological systems. To examine the XO pathway during MV we measured both XDH and XO activities in the diaphragm. Our findings show that diap hragmatic XO activity is increased during MV as shown by the increase in XO enzy matic activity and XO protein e xpression (130 kDa band) in the diaphragm. Moreover, hypoxanthine and xanthine, substrates for XO, were depleted in the diaphragms of our ventilated animals while uric acid, an end product of enzymatic activity, was significantly elevated. Interestingly, prolonged (18 hrs) MV resulted in a pronounc ed increase in the 150 kDa band of XOR along with a concomitant increase in total XOR activity in the diaphragm. This may have been the result of an increase in XOR protein synthesis and/ or a decrease in XOR protein degradation during prolonged MV. Howeve r, these experiments do not reveal the exact mechanism for increased XOR prot ein expression during prolonged MV. Pharmacological inhibition of XO via oxypurinol administration attenuated the MVinduced induction of diaphragmatic XO activity and uric acid formation. As expected, oxypurinol administration had no ef fect on the protein levels of XO (130 kDa band of XOR) in the diaphragm but interestingly increased the induction of the 150 kDa band of XOR following ventilation. Consequently, XO inhibition during MV may produce a feedback system in which the diaphragm muscle produces more XOR prot ein to overcompensate for the pharmacologicalinduced inhibition of enzymatic activity. Overa ll, the MV-induced induc tion of diaphragmatic XO activity was abolished with oxypurinol administration. 86

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XO Inhibition during MV Attenuates Diaphr agmatic Oxidative Stress and Contractile Dysfunction Previous work from our laboratory has re vealed that MV promotes diaphragmatic oxidative injury and contractile dy sfunction (12, 20, 31, 80, 81, 98, 108, 110, 122, 132). Therefore, we postulated that if diaphragmatic XO activ ity was induced during MV, XOmediated oxidant production would contribute to the MV-induced diaphrag matic oxidative stress and contractile dysfunction that is normally observed. First, we confirmed our earlier reports by showing that our MV animals exhibited an increased level of oxida nt production in the diaphragm when compared with control as both protein oxidation a nd lipid peroxidation increased with MV. However, for the first tim e we have shown that XO inhibition significantly reduces protein oxidation and lip id peroxidation in the diaphr agm during MV. While oxypurinol administration did not completely attenuate MV -induced protein oxidation in the diaphragm, oxypurinol significantly blunted the appearance of 4-hydroxynonena l modified proteins. The failure of oxypurinol to completely prevent oxidative stress in the diaphragm during MV does not appear to be due to an ineffectiv e dose of oxypurinol. Inde ed, it has been reported that, oral oxypurinol treatment of 50 mg/kg/day for 2 days, pr ovides an extracellular fluid concentration of 10 M oxypurinol and that this concentration is sufficient to cause a >80% inhibition of XO activity without a scavenging effect (134). Thus our findings indicate that our dose of oxypurinol effectively inhi bited purine substrate binding and prevented the enzyme from catalyzing the conversion of the s ubstrates into superoxide and ur ic acid. The observation that oxypurinol only inhibited some of the oxidative stress in the dia phragm during MV supports our hypothesis that XO plays a role in the producti on of ROS in the diaphragm during MV but suggests that XO is not the only source of ROS in the diaphragm. 87

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Based upon our finding that XO i nhibition was associated with decreased oxidative stress in the diaphragm, we anticipated that XO inhib ition would significantly a ttenuate the contractile dysfunction that is normally obser ved with MV. Our results suppor t this notion as XO inhibition attenuated the decrease in diaphr agmatic specific force at stimula tion frequencies above 60 hertz. This result is in agreement with Matuszczak and colleagues who found that XO inhibition via allopurinol lessened contractile dysfunction of skeletal musc le caused by prolonged unloading (78). The fact that oxypurinol administration dur ing MV attenuated both diaphragmatic XO activity levels and improved contr actile function suggests that free radical damage contributed to the contractile dysfunction. In particular, oxidative modification of myosin and/or actin is a potential cause of mechanical dys function given their critical role in the contractile machinery. The observation that oxypurinol therapy improved isometric twitch force and shifted the forcefrequency curve toward control suggests a direct effect of oxypurinol in attenuating contractile protein oxidation. A similar finding was noted recently by Kogler et al. who found that oxypurinol enhanced cardiac muscle twitch tension in spontaneously hypertensive, heart failureprone rats without alterations in intr acellular calcium amplitudes (59). While XO inhibition prevented some of the d ecrease in diaphragmatic dysfunction, it did not completely restore contractile function to co ntrol. The partial protection of contractile function with XO inhibition may be due to ROS generation from additional pathways. In our MV model, XO may work in conjunction with other oxidant producing pa thways (i.e., hemeoxygenase-1 and mitochondria) in the diaphragm to generate free radi cals. This theory is logical given that oxypurinol only inhibits XO activity and does not i nhibit ROS production from other oxidant producing pathways and the fact th at the MV-induced oxidative stress was not 88

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completely restored with XO inhibition. Accordingly, ROS generation from another pathway could theoretically contribute to both the MV-induced oxidative stress and contractile dysfunction that is observed in the diaphrag m that is not restored with XO inhibition. Antioxidant Administration during MV Fails to Attenuate XO Activation Mammalian XORs (XDH & XO) catalyze the hydroxylation of hypoxanthine and xanthine at the molybdenum cente r of the enzyme and reducing e quivalents thus introduced into the enzymes are transferred via two iron sulfur centers to flavin adenine dinucleotide (FAD), where the reduction of NAD+ or mo lecular oxygen occurs (60). These enzymes are synthesized as the dehydrogenase form (XDH) but can be readily converted to th e oxidase form (XO) reversibly by oxidation of sulfhydryl residues or irreversibly by proteolysis (2, 22, 91, 114). During the reversible XDH to XO conversion, oxidation of cysteines residues on XDH results in the displacement of an active loop around FAD. The displaceme nt of this loop blocks the approach of the pyridine ring of the NAD+ substrate to the isoa lloxazine ring of the enzymes cofactor and changes the electr ostatic environment around FAD. Thus the movement of the active loop causes the reversible conversion of XDH to XO (27). Therefore, the reversible conversion of XDH to XO via the oxidation of cysteine residues on XOR changes enzymatic function but does not change the enzymes molecula r weight. Hence, it is feasible that during MV XO activity may be up-regulated via the ox idation of XDH to XO. Once again, pathways other than XO may generate ROS during MV and the oxidants from one of those pathways may be responsible for the reve rsible activation of XO. In an attempt to understand the reversible conversion mechanism by which XO activity maybe increased in the diaphragm during MV, we used the pharmacological agent Trolox to maintain redox balance. Trolox is a water so luble Vitamin E analog that has been shown to 89

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reduce MV-induced oxidative stress and contrac tile dysfunction (12, 80, 81). So initially, we confirmed our earlier reports by showing that Trolox administration effectively attenuated both protein oxidation and lipid peroxidation in the diaphragm during MV. Second, we demonstrated the ability of Trolox to attenuate all of the cont ractile dysfunction that is normally observed with 12 hours of MV. As a result, our use of Trolox as a means to restore redox balance in the diaphragm was achieved in this experiment. Because Trolox administration attenuated the MV-induced oxidative stress in the diaphragm, we hypothesized that the MV-induced induction of XO activity would be blunted by the elimination of the revers ible conversion of XDH to XO vi a oxidation. However, Trolox administration during MV failed to attenuate either the increase in XO activity in the diaphragm during MV or the increase in uric acid form ation. Furthermore neither the levels of hypoxanthine or xanthine were changed in th e animals treated with Trolox. Hence, the restoration of redox balance with Trolox does not affect the increas e in diaphragmatic XO activity during MV. It is important to note that our MV duration may not have been adequate to examine the reversible conversion of XDH to XO in the diaphr agm. Specifically, the ir reversible proteolytic processing of XDH results in the cleavage of a 15-20 kDa fragment from the enzyme resulting in the appearance of the active 130 kDa band of XO R (22, 130). During this conversion, the active loop around FAD is moved resulting in an electrostatic environment that favors the use of only molecular oxygen in the reduction of the purine subs trates. Importantly, th is proteolytic process is believed to occur subsequent to the reversible conversion (14, 83). Because the irreversible proteolytic conversion of XDH to XO is present in the diaphrag m during MV, as demonstrated by the increase in diaphragmatic XO activity al ong with the induction of the 130 kDa band of 90

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XOR, we may have missed the re versible conversion of XDH to XO in our model because our ventilation was too long. Perh aps the reversible conversion of XDH to XO in the diaphragm would be adequately observed following 3 or 6 hours of MV. Hydrogen Peroxide Activates XO in Myotubes Oxidant challenges (H2O2) to C2C12 myotubes can induce oxidativ e stress and proteolysis (74, 116). In addition, hydrogen peroxide has been shown to modulate the reversible conversion of XDH to XO in non-muscle cell lines (85). Therefore, as a means of providing a controlled in vitro environment to study the activati on mechanism(s) of XO with a H2O2 challenge, we included a myogenic cell culture model in our experiments. Specifically, we tested the hypothesis that exposure of skel etal muscle myotubes to hydr ogen peroxide (100 M) would increase XO activity via the oxidative modification of XDH to XO. Our results indicate that hydrogen peroxide increases the proteolytic c onversion of XDH to XO via the induction of the active form of XO protein (130 kDa band) in co njunction with increases in XO activity. These results are similar to our in vivo results whereby XO activity was increased in the diaphragms of our MV animals. While indicating that the irreversible process is increased in our in vitro model, these results fail to specifically identify the pr otease responsible for th e induction of the 130 kDa band of XOR. Knowing that hydrogen peroxide in creases calpain activity in our C2C12 cell line, we inhibited calpain activity in our cell culture experiments using both leupeptin and short interference RNA (siRNA) to focus solely on the oxidative challenge presented by hydrogen peroxide treatment. Leupeptin is an inhibitor of lysosomal thio l proteases and calcium-activated proteases (77). Specifically, leupe ptin inhibits papain, cathepsin B, endoproteinase Lys-C, and calpain. Interestingly, we found that when cells were treated with both hydrogen peroxide and 91

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leupeptin, the hydrogen peroxide-i nduced increases in both XO activity and protein expression were significantly attenuated, sugge sting that the irreversible pr oteolytic conversion of XDH to XO in skeletal muscle myotubes with a ROS challenge (H2O2) involves either a lysosomal or calpain protease. Short interference RNA (siRNA) for calpain-1 was also used to inhibit the hydrogen peroxide-induced increase in calpain activity in our C2C12 cell line. siRNA is used to interfere with the expression of the calpain-1 gene. We found that ce lls treated with both hydrogen peroxide and siRNA for calpain-1 exhibited the same increase in the 130 kDa band of XOR in concomitant with an increase in XO activity as cells treated with only hydrogen peroxide, proving that the irreversible conversion of XDH to XO via a ROS challenge does not involve calpain-1 activation. Our cell culture results coin cide with work by McNally et al. who found that H2O2 markedly enhances the irreversible conversion of XDH to XO in endothelial cells (85). In addition, with respect to MV, our cell culture experiments shed light on the observation that leupeptin can inhibit ventilator -induced diaphragm dysfunction in rats (77). Perhaps leupeptin prevents the induction of XO activity in the in vivo MV model and prevents XO-mediated ROS production. However, our results do not ag ree with a study performed on human liver by Saksela et al (102). While these researchers s uggest that excess intracellu lar calcium is involved in the XDH to XO conversion in human liver, they predict that a mitochondrial protease is responsible for the proteolytic cl eavage of XDH to XO. In an ischemia-reperfusion model, they theorized that an intracellular calcium in flux produced by reperfusion would damage the mitochondria resulting in the opening of the mi tochondrial permeability transition pore and result in the release of m itochondrial proteins into the cytoplasm (102). While these researchers failed 92

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to identify the specific mitochondrial protease, a study by Sitte et al. recently characterized a mitochondrial intermembrane protease isolated fro m rat liver that is si gnificantly inhibited by leupeptin (111). Additional work is required to determine if a mitochondrial protease is responsible for activation of XO in myotubes exposed to a ROS challenge. Conclusions and Future Directions This study provides the first in vivo evidence that diaphragmatic XO activity contributes to MV-induced oxidative stress and contractile dysf unction. Specifically, thes e results demonstrate that XO inhibition attenuates MV-i nduced oxidative stress and elimin ates some of the contractile dysfunction that is normally obs erved with MV. Furthermore, XO is up-regulated in the diaphragm by the irreversible proteolytic cl eavage of XDH to XO during MV. Finally, the irreversible conversion of XDH to XO in skeletal muscle myotubes induced by a ROS challenge (H2O2) is a calpain-1-independent process. Coll ectively, these are novel and important findings. In fact, prior to the current experiments, the question of whether XO contributes to MV-induced oxidative damage in the diaphragm remained unknown. Our results clearly indicate that XOmediated production of superoxi de is involved in MV-induced diaphragmatic oxidative stress and contractile dysfunction. Moreover, our findi ngs suggest that pharmac ological inhibition of XO activity could be a potential therapeutic strategy to retard MV-induced oxidative stress and contractile dysfunction in the diaphragm during prolonged MV. This is clinically significant because MV-induced diaphragmatic weakness plays an important role in weaning difficulties following MV. Future studies should focus on elucidating the protease that is responsible for the proteolytic cleavage of XDH to XO in skeletal muscle and effectively determine if the reversible conversion of XDH to XO via oxidati on is an essential first step in the activation of XO activity in skeletal muscle under conditions of disuse. A complete understanding of these two important 93

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94 issues is essential to fully understand the XO pathway in skeletal muscle. Specifically, the complete understanding of the XO pathway in the diaphragm during MV will be beneficial in the development of effective and safe countermeasures for preventing respiratory muscle weakness during prolonged MV. The development of a therapeutic strategy to retard MV-induced diaphragmatic oxidative damage and weakness would likely result in the maintenance of adequate inspiratory muscle function a nd result in higher weaning success rates.

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BIOGRAPHICAL SKETCH Melissa A. (Deering) Whidden was born in Cortland, New York. She earned a Bachelor of Science degree in exercise science from the University at Buffalo, The State University of New York. Following graduation, she pursued a masters degree in applied physiology and graduated from the University at Buffalo, The State University of New York in 2003. Deciding to focus her career in basic scien ce, Melissa began her doctoral work at the University of Florida in 2003 under the direction of Scott K. Powers. Melissa focused her studies on oxidative stress and proteolysis of the diaphragm muscle during prolonged mech anical ventilation. She received her PhD in 2008. 106