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Neutrophil mediated muscle injury of the diaphragm Samrai, Baljit 1999

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NEUTROPHIL MEDIATED MUSCLE INJURY OF THE DIAPHRAGM By BALJIT SAMRAI B. Sc. The University of British Columbia, 1993 A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF MASTER OF SCIENCE in THE FACULTY OF GRADUATE STUDIES Experimental Medicine We accept this thesis as conforming to the required standard  THE UNIVERSITY OF BRITISH COLUMBIA 1999 © Baljit Samrai, 1999  In  presenting  degree freely  this  at the  thesis  in  partial  University  of  British  available for  copying  of  department publication  this or of  reference  thesis by  this  for  his thesis  and  fulfilment  for  her  the  requirements  Columbia, I agree that the  study.  scholarly  or  of  I further  purposes  may  representatives.  financial  gain  agree  It  shall not  be is  that  C Kfzr<<L\ rv/\evvj-f / y - i -  The University of British C o l u m b i a Vancouver, Canada  Date  DE-6  (2/88)  1U  r^W^,  permission  granted  by  an  \ C i ^ r  advanced  shall make for  the that  allowed without  permission.  Department of  Library  understood be  for  it  extensive  head  of  my  copying  or  my  written  ii ABSTRACT Excessive loading, as seen in chronic respiratory diseases and experimentally produced by tracheal banding (TB) may result in injury to the myofibres of the diaphragm and hypercapnic ventilatory failure. Neutrophils (PMNs) are primary inflammatory cells that migrate to an area of injury in order to phagocytose cellular debris and promote the inflammatory response. During this process, neutrophils may also amplify muscle injury. We hypothesized that neutrophil depletion would reduce diaphragm muscle injury in an animal model where diaphragm injury was induced by tracheal banding. Adult male Sprague-Dawley rats were randomly assigned to one of four groups: 1. control (C), 2. tracheal banded (TB), 3. control plus neutrophil depleted (C+ND), 4. tracheal banded plus neutrophil depleted (TB+ND). In the C+ND and TB+ND groups, neutrophils were depleted by administering anti-rat polymorphonuclear leukocyte antisera, beginning one day pre-surgery. After anaesthesia, the carotid artery was chronically cannulated, esophageal pressures (Pes) were measured and a cuff was tightened around the isolated trachea, until the Pes during tidal breathing (Pesjy) was 20% of the maximal Pes (Pes ) during tracheal Max  occlusion in the TB and TB+ND groups. Arterial blood gas (ABG) samples were taken daily from the awake animal. Three days later, following anaesthesia, in vivo measures of Pes were repeated and then the diaphragm and trachea were excised for histological and immunohistochemical analysis. The following parameters were investigated: (1) the presence of neutrophils in the diaphragm measured by neutrophil specific myeloperoxidase (MPO) activity and point counting H&E diaphragm cross-sections; (2) muscle injury as quantified by point counting H&E stained diaphragm cross-sections; (3) the presence of EDI positive macrophages in the diaphragm by using immunohistochemistry; and (4) ABGs measure daily post-surgery. The TB groups were hypercapnic (PaC0 > 45mmHg; p<0.001) and had a decreased tracheal cross2  sectional area (p<0.001). ND groups had PMN counts <0.5 million/mL. MPO activity tended to increase indicating an increased presence of neutrophils in the diaphragm of the TB group, and C values were similar to those in the TB+ND group. The TB group had the most abnormal and inflamed muscle (p<0.05), whereas, the TB+ND group was similar to C values. EDI positive macrophages per cross-sectional area was greater in the TB group compared to the C group, indicating that the macrophage population increased in response to muscle injury. We conclude that macrophages increase and neutrophils tend to increase in the injured diaphragm induced by tracheal banding. Neutrophil depletion decreased diaphragm injury induced by resistive loading.  IV  T A B L E OF CONTENTS ABSTRACT  ii  TABLE OF CONTENTS  iv  LIST O F T A B L E S  vi  LIST O F F I G U R E S LIST O F ABBREVIATIONS ACKNOWLEDGEMENTS CHAPTER 1  vii ix xii 1  STATEMENT OF THE PROBLEM LITERATURE REVIEW Exertion-Induced Muscle Injury Respiratory Disorders Chronic Respiratory Disorders (CRD) Acute Respiratory Disorders Respiration Respiratory Muscle Dysfunction in Respiratory Conditions Respiratory Muscle Injury Inflammation HYPOTHESIS  1 3 3 5 6 9 13 16 18 25 35  CHAPTER 2 METHODS Animals and Groups Experimental Protocol Neutrophil Depletion White Blood Cell Counts Arterial Blood Gases Measures of Resistive Loading Muscle Histology & Immunohistochemistry Diaminobenzidine Tetrahydrochloride (DAB) Method Point Counting H & E Cross-Sections for Muscle Abnormality and Neutrophils in the Diaphragm Immunohistochemistry Quantification of Macrophages Myeloperoxidase Assay Statistical Analyses  52 52 52 53 55 57 58 58 60 60  CHAPTER 3  73  RESULTS  61 65 67 68 69  73  V  Tracheal Cross-Sectional Area Esophageal Pressures Circulating Absolute Neutrophil Counts Arterial Blood Gases Hemi-Diaphragm and Body Weights Area Fractions of Abnormal Diaphragm Area Fractions of Neutrophils Immunohistochemistry Myeloperoxidase  73 73 74 74 75 76 76 76 77  CHAPTER 4 DISCUSSION CONCLUSION FUTURE DIRECTIONS  109 .109 123 124  REFERENCES  125  APPENDK  136  vi  LIST OF TABLES 1.  Point Counting Categories and Characteristics  2.  Categories and Characteristics for Point Counting H&E Stained Cross-Sections for Neutrophils  62  64  3.  Tracheal Cross-Sectional Area and Esophageal Pressures  89  4.  Circulating Absolute PMN Results  93  5.  Arterial Blood Gas Results  95  6.  Body and Hemi-Diaphragm Weights  97  7.  Area Fractions of Normal, and Abnormal Muscle and Connective Tissue in Diaphragm 99  8.  Area Fractions of PMNs  103  9.  ED1+ Macrophages in the Diaphragm  104  10.  MPO Activity in the Diaphragm  108  vii  LIST OF FIGURES 1.  Hypothetical model of respiratory muscle injury  36  2.  Diaphragm muscle viewed from below  37  3.  Respiratory muscles of the thorax  38  4.  Hypothetical inflammatory response to muscle injury  39  5.  Organization of skeletal muscle  40  6.  Electron microphotograph of skeletal muscle fibres (cross-section)  41  7.  Electron microphotograph of a sarcomere  42  8.  Electron microphotograph illustrating the hexagonal array of myofilaments as seen at the cross-section at an A-band  9.  Electron microphotograph of Z lines cut'en face'  10.  Electron microphotographs illustrating the doublet array of mitochondria along the Zline  43 44  45  11.  Electron microphotograph showing fiber types and mitochondrial distribution  46  12.  Photomicrograph of normal rat diaphragm, cross-section, stained with H&E  47  13.  Photomicrographs of inflamed and damaged muscle fibres in the diaphragm takenfroma tracheal banded rat, stained with H&E  14.  Electron microphotograph of diaphragm from a tracheal banded rat, showing abnormal muscle  15.  49  Photomicrograph of ED 1 + macrophages in the diaphragm cross-section of tracheal banded and neutrophil depleted rat  16.  48  50  Photomicrograph of diaphragm muscle cross-section taken from a tracheal banded rat, stained with DAB and counterstained with Mayer's haemotoxylin solution... 71  viii  17.  Photomicrograph of overstained cross-section of rat diaphragm with Accurate anti-rat neutrophil antibodies, counterstained with Mayer's haemotoxylin solution . . . 72  18.  Bar graph of tracheal cross-sectional areas in control and banded groups  78  19.  Bar graph showing esophageal pressures (PeSry)  79  20.  Scatter graph showing circulating absolute PMN counts on days 0 through to 4 . . . . 80  21.  Bar graphs for arterial blood gas (ABG) data  81  22.  Bar graph of normalized hemi-diaphragm weight  83  23.  Bar graphs showing area fractions of normal and area fractions of abnormal muscle, and connective tissue in the diaphragm  24.  84  Photomicrograph of an abnormal feature in the musclefibrein the diaphragm from a tracheal banded and neutrophil depleted rat, stained with H&E  85  25.  Bar graphs showing numbers of EDI positive macrophages in the diaphragm  86  26.  MPO activity in the diaphragm  88  LIST OF ABBREVIATIONS  A  A  area fraction  ABC-AP  vectastain avidin biotin complex  ABG  arterial blood gases  ABL  acid-base laboratory  AbN  area fraction of abnormal muscle  ARE)  acute respiratory disorders  ATS  american thoracic society  BW  body weight  bv  blood vessel  C  control  C5a  complement fragment 5 a  CD  cluster designation  CFU-GM  colony forming unit - granulocyte, monocyte  COPD  chronic obstructive pulmonary disease  CRD  chronic respiratory disorders  CSA  cross-sectional area  CT  connective tissue  DAB  diaminobenzidine tetrahydrochloride  DIA  diaphragm  DOMS  delayed onset muscle soreness  ECRL  extensor carpi radialis longus  EDI  inflammatory macrophage marker  GM-CSF  granulocyte monocyte-colony stimulating factor  H&E  haemotoxylin and eosin  Herni-Dia  hemi-diaphragm  HTAB  hexadecyltrimethylammonium bromide  ID  internal diameter  IFNy  interferon gamma  IL-1  interleukin 1  im  intra-muscular  Infl-O/L  inflammatory cells in necrotic tissues, no muscle cell outline  Infl+O/L  inflammatory cells in necrotic tissue with muscle cell outline  ip  intra-peritoneal  Mcf)  macrophage  M  molar  mEq/L  milliequimolar per litre  mrtiHg  millimeters of mercury  MPO  myeloperoxidase  Mu  muscle  MV/V  millivolts per volt  NADH  nicotinamide adenine dehydrogenase  ND  neutrophil depleted  No  number  OD  external diameter  O-DMB  (9-dianisidine dihydrochloride  PaC0  partial pressure of carbon dioxide in arterial blood  Pa0  2  partial pressure of oxygen in arterial blood  2  esophageal pressure during tidal ventilation P Max  maximal esophageal pressure  PGE  prostaglandin E  e S  2  2  PMN  neutrophil  PTI  pressure time index  Rd-nucl  inflammatory cell nucleus  SCM  sternocleidomastoid  SD  standard deviation  SE  standard error  Sp-nucl  spindle shaped nucleus  TB  tracheal banded  TB+ND  tracheal banded plus neutrophil depleted  TBS  tris buffered saline  TDMac  tridodecylmethylammonium chloride  Ti  inspiratory time  TNF  tumour necrosis factor  Trach  tracheal  T-Tot  total time taken for one respiratory cycle  WBC  white blood cells  xii  ACKNOWLEDGEMENTS I would like to thank the Department of Experimental Medicine for their support and guidance, especially Ms Anita Keister, I am truly grateful for your help and genuine kindness. I would like to acknowledge and to express my appreciation to the following members of my supervisory committee: Dr. Clive Roberts, Angelo N. Belcastro, and especially to Dr. David C. Walker for his continual support and helpful advice. A very special thanks to Mr. Michael Boyd and his team at the Animal Care Unit. Assistance and suggestions were very much appreciated from Dr. Mark Elliott, Ms. Jenny Hards, Ms. Fanny Chu, and Mr. Stuart Greenefromthe Pulmonary Research Lab at St. Paul's Hospital. I would also like to thank the U.B.C. hospital laboratory for allowing me to use their equipment at my convenience. A heartfelt thanks to all of my friends who have been supportive beyond belief and encouraging to no end. Finally, my gratitude to my mother for providing a roof over my head, food in my stomach, and a shoulder to lean on.  1  CHAPTER 1 STATEMENT OF THE PROBLEM Inspiratory muscles are unique in their requirement to function continuously to keep us alive. They are unlike other skeletal muscles of the body that can reduce activity to rest fatigued muscles. Inspiratory muscles may become vulnerable to fatigue and damage when they work against excessive loads during chronic conditions such as in respiratory diseases (Hards et al., 1990). In severe conditions of respiratory muscle injury, ventilatory failure can occur which could potentially lead to death (Rochester, 1993). In patients suffering from chronic respiratory disorders (CRD), evidence of poor muscle function has been found in respiratory muscles (Begin & Grassino, 1991; Roussos, 1985). As pathophysiological changes occur respiratory muscles become vulnerable to fatigue, weakness and injury. Emphysema and chronic bronchitis are conditions which cause progressive limitation of airflow and increase the work of breathing due to mechanical disadvantage (Rochester, 1993). Respiratory muscle dysfunction in humans has been implicated as a cause of hypercapnic ventilatory failure (Begin & Grassino, 1991; Roussos, 1985). A state of hypercapnia or hypoxia may further impede the force generating ability of a weakened diaphragm (Reid & Dechman, 1995). Inflammation is a natural response that occurs as a result of an insult such as trauma, extreme temperatures, and exercise (Faulkner et al., 1993). Inflammation can occur as a result of diaphragm muscle injury caused by an increased workload. Many inflammatory mediators signal inflammatory cells such as neutrophils, monocytes, and macrophages, to chemotax to the area of injury (Cannon & St. Pierre, 1998). Chemotaxis is the movement of cells along a chemical concentration gradient (Taylor, 1988). Inflammatory cells respond to chemotactic agents released  2  at inflammatory sites, including cytokines, activated complement components, and bacterial products (Taylor, 1988). Phagocytic cells are responsible for degrading and clearing cellular debris, however, during the inflammatory process they may also amplify tissue injury by attracting other inflammatory cells. Neutrophils are one of the first cells to arrive at the site of injury. In the state of neutropenia, fewer circulating neutrophils would be available to respond to injury and less chemoattractants may be produced attracting fewer inflammatory cells, and thus resulting in less injury. Not only may less injury occur after neutrophil depletion (ND) but ND may decrease the influx of macrophages into injured tissue. Tracheal banding (TB) is an animal model where excessive loads are imposed by increased resistive loading. The excessive loading of respiratory muscles through tracheal banding has been shown to result in injury to the myofibres (Reid et al, 1994; Reid, 1993) where maximum injury was observed following three days of tracheal banding (Reid, 1993). TB rats develop ventilatory failure as a result of increased inspiratory load which is evident by hypercapnia (Reid et al., 1992). Ventilatory failure is the inability to ventilate adequately (Reid et al., 1992). A reduction of inflammation by ND may reduce ventilatory failure and improve arterial blood gases. Thereby, this study will show how some of the major inflammatory cells are related to diaphragm injury in an animal model of hypercapnic ventilatory failure.  3  LITERATURE REVIEW Exertion-Induced Muscle Injury Exertion-induced muscle injury is the disruption within the skeletal muscle fibre following unaccustomed activity. It has been well documented as shown in humans (Jones et al., 1986; Hoppeler, 1986; Friden et al., 1983) and in animals (Friden & Lieber, 1992; Armstrong et al., 1983). The activity that requires greater exertion, than to which the muscle is accustomed, results in injury to the vulnerable muscle and delayed onset of muscle soreness (DOMS) (Maclntyre et al., 1995, Armstrong, 1990). DOMS is the discomfort that is experienced following unaccustomed activity, which develops 24 hours post exercise and is at maximal discomfort within two to three days (Clarkson et al., 1992). A bimodal pattern of injury has been described post-exercise. Firstly, the initial event occurs as a result of the muscular contractions (Armstrong, 1990). Eppinger et al. (1995) describes the initial event in reperfusion injury of rat lung as "neutrophil independent". Secondly, the delayed event occurs as a result of inflammation (Armstrong, 1990) or termed as neutrophil mediated (Eppinger et al., 1995). Potential mechanisms of the initiating event, which result in skeletal muscle injury, may be mechanical or metabolic in nature. The mechanical factor could be due to an increase in muscular tension (Armstrong, 1990) stressing the myofibrils and other intramuscular components. Metabolic causations of injury could be due to elevated intramuscular temperature, elevated production of oxygen free radicals, acidosis, or activation of calcium activated proteases such as calpain (Armstrong, 1990). The work by Dr. Robert Armstrong (1990) has received much recognition; he has categorized exertion induced muscle injury into four stages: 1. the initial event; 2. the autogenic stage; 3. the phagocytic stage; and 4. the regenerative stage. As a result of the initial event, a cascade of  4 events proceed and the autogenic stage commences. The autogenic stage begins with selfdegradation via intrinsic proteases and phospholipases of cell membranes and cytoskeletal proteins, which can last for several hours (Armstrong, 1990). The phagocytic stage is announced by the influx of inflammatory cells, beginning as early as four to six hours and lasting up to two to four days (Armstrong, 1990). Neutrophils have been recognized as being one of the earlier cells to arrive at the site of injury during the inflammatory response (Tidball, 1995), hence, the injury has been described as neutrophil-mediated (Eppinger et al., 1995). During this stage, inflammatory cells release their toxic contents and phagocytosis of cellular debris is initiated. This secondary muscle injury, which peaks at three days following the initial event, is associated with the inflammatory response and force loss (Carlson & Faulkner, 1993). Monocytes/macrophages, present later during the inflammatory response (two to five days after the initial event), have primarily been recognized for clearing cellular debris. The final stage is referred to as the regenerative stage where the muscle cells attempt to repair themselves.  The repair process involves the activation of satellite cells to  form neo-myotubes to become muscle fibres (Carlson & Faulkner, 1993). The stage of regeneration peaks after seven days (Kihlstrom et al., 1984). Exertion induced muscle injury occurs as a training response. Muscle injury in exercised limbs can range from mild to severe, however, a medical treatment is usually not required (Maclntyre et al., 1996) and there is usually no long-term functional impairment (Lieber, 1992). Repeated activities with intervals of rest to allow recovery can result in trained and adapted muscles (Clarkson, 1992; Clarkson & Tremblay, 1988; Byrnes et al., 1985). Trained muscles become less vulnerable to injury.  Respiratory Disorders Inspiratory muscles may become vulnerable to fatigue and damage as they are continuously required to work against excessive loads as during chronic or acute respiratory disorders (ARD) (Hards et al., 1990). Exertion-induced skeletal muscle injury can occur following an initiating event which is beyond the normal functioning capacity of the muscle (Armstrong, 1990). Therefore, in conditions where respiratory muscles are subjected to exertion and the load is relentless, respiratory muscle injury is likely. Respiratory disorders can increase the work of breathing due to factors such as increased airway resistance, mechanical disadvantage (Rochester, 1993), and through pathophysiological changes that occurfromchronic respiratory disorders (CRD). CRD brackets many conditions such as chronic obstructive pulmonary disease (COPD), cystic fibrosis, bronchiectasis, and tuberculosis (Professional Guide to Diseases, 1995). ARD includes conditions such as acute respiratory failure, adult respiratory distress syndrome, pulmonary edema, cor pulmonale, atelectasis, respiratory acidosis/alkalosis, pneumothorax, hemothorax, pneumonia, pulmonary embolism and infarction, lung abscess, pleural effusion and empyema (Professional Guide to Diseases, 1995).  Significance The impact of CRD on the health care system is overwhelming. In 1994, approximately 2.3 million patients in the United States were hospitalized for COPD (Integrated Medical Services, 1996). CRD patients spend more time at the physician's office, require greater number of work disability days, and have more frequent short-term hospitalizations in comparison to the general public (Feinleib et al, 1989). COPD is a disease which affects at least 14 million people in the  United States and according to the American Thoracic Society (1995) the figures have increased by 42% since 1982. In comparison to 1966, COPD mortality has increased by 71% whereas a decrease has been observed in cardiovascular (45%) and cerebrovascular diseases (52%) (Integrated Medical Services, 1996). The risk of COPD increases with age and is a leading cause of morbidity and mortality throughout the world (Whittemore et al, 1995). The objective of this section is to make the reader aware of the complexities surroundinng respiratory conditions, their complications, and how they affect the respiratory system. COPD has been described in detail to provide information on pathophysiological changes that can occur during the progession of a debilitating disease and how it can affect the respiratory system. Other chronic and acute respiratory conditions, and associated disorders have been discussed briefly but focussed on their impact on the respiratory system.  Chronic Respiratory Disorders (CRD) Chronic obstructive pulmonary disease (COPD) is a condition where the work of breathing is increased due to increased airway resistance and mechanical disadvantage (Rochester, 1993). Chronic obstructive pulmonary disease is a clinical term, which embraces a group of diseases characterised by airflow limitation during expiration. Diseases included under COPD include emphysema, chronic bronchitis, and sometimes asthma (Reid & Samrai, 1995). Airflow limitation can occur as the result of the following: an abnormal enlargement of the alveoli and a loss of elastic recoil of the lungs, as in emphysema; an increase in sero-mucous glands and an excess of mucus secretion, increased bronchospasm and edematous airway walls, as in chronic bronchitis. In chronic asthma, cardinal changes include mucous plugging, thickening of the basement membrane, infiltration by eosinophils in the bronchial wall and hyper-irritability of airways.  7  Progressive obstructive lung disease causes many devastating consequences for the patient such as dyspnea and dynamic pulmonary hyperinflation (Killian & Campbell, 1983). A patient with hyperinflation presents the following clinical manifestations: barrel shaped chest; larger lung fields; flatter diaphragm; more horizontal rib alignment; upper chest breathing pattern with the recruitment of accessory muscles; increased functional residual capacity, residual volume, total lung capacity (West, 1987) and respiratory rate. The pressure time index (PTI) is used to measure respiratory muscle overload and is defined as the product of [inspiratory pressure during tidal breathing/maximal inspiratory pressure] by [time taken for inspiration/ total time taken for one breath] (Rochester, 1991). The compensatory mechanism to hyperinflation is to reduce the time taken for inspiration resulting in rapid shallow breathing resulting in an increased PTI. Patients with respiratory disorders can have up to a 20 fold increase in their PTI values, which can exceed a fatiguing threshold of a PTI at 0.15 (Rochester, 1991). The pathophysiological response to increased airways resistance is hyperinflation. The benefit of hyperinflation is that the airways become dilated, which decreases resistance (Macklem, 1984) thereby increasing expiratory flow rates and increasing tidal lung volume towards total lung capacity (Derenne et al, 1988). Although hyperinflation may decrease airways resistance, it has a negative impact on the muscles responsible for inspiration, especially the diaphragm (Rochester, 1991; Decramer er al, 1986; Decramer & De Troyer, 1984). Increased lung volumes due to hyperinflation causes the diaphragm to flatten which shortens the length of the muscle and therefore reduces its ability to generate tension (Rochester, 1991; Grassino et al, 1978). Farkas & Roussos (1983) showed that hyperinflation in emphysematous hamsters places the diaphragm at a mechanical disadvantage. They found a loss of sarcomeres which may be an attempt to  8  improve the length tension relationship for muscle contractility of the diaphragm working at a shorter length. Similowski et al (1991) believes that adaptation via absorption of sarcomeres may occur in patients with chronic airflow limitations. Diaphragm adaptation to hyperinflation compromises tension development by altering the costal and crural regions of the muscle from a parallel to a more series orientation (Macklem, 1984; Macklem et al, 1983). As COPD progresses, the action of the diaphragm becomes increasingly limited (Macklem et al, 1983) shifting the action of ventilation to the responsibility of the intercostal and accessory muscles. The recruitment of accessory muscles for quiet breathing results in an increased energy consumption (Fleury et al, 1987), which could lead to weight loss. In severe hyperinflation, the diaphragm may become expiratory in action as seen by the paradoxical movement of the chest wall during inspiration (Goldberg & Roussos, 1990; Zocchi et al, 1987). This may occur due to the reconfiguration of the diaphragm fibres (Zocchi et al, 1987) orfromweakening of the diaphragm. Increased lung volumes affect the compliance of the chest wall by decreasing the outward movement of the chest wall during inspiration and at veiy increased lung volumes, the inspiratory muscles must generate force against the inward recoil of the chest wall (Rochester, 1991; De Troyer et al, 1985). In patients sufferingfromCOPD, evidence of poor muscle function has been found in the respiratory muscles (Begin & Grassino, 1991, Roussos, 1985). In emphysema where there is a loss of elastic recoil of the lungs, the diaphragm and accessory muscles are recruited to work at excessive workloads. Hyperinflation impacts the respiratory system as seen by changes in breathing patterns as mentioned earlier. Cystic fibrosis, an autosomal recessive trait, is a chronic disorder of the exocrine glands. It affects the respiratory system by the production of thickened bronchial secretions resulting from  dysfunctional chloride ion transport, which ultimately leads to blockage of the glandular ducts. Signs and symptoms include noisy breathing, nonproductive violent attacks of coughing, dyspnea, tachypnea, recurring bronchitis and pneumonia. The condition progresses to severe atelectasis and emphysema and results in death (Norris, 1995). Bronchiectasis is the abnormal dilation of the the bronchi and the destruction of the bronchial walls. This disorder occurs from abnormal mucocilliary clearance and repeated damage to the bronchial walls (due to underlying conditions such as cysticfibrosis),which results in a breakdown of supporting tissues around the airways. The patient with bronchiectasis has recurring respiratory tract infections and abscess formations which recruits inflammatory cells and produces fibrous tissue; mucous plugs or abscesses can destroy smaller bronchioles; coarse rales during inspiration; dyspnea; and weight loss could further weaken the individual (Norris, 1995). Tuberculosis is caused by the microorganism Mycobacterium tuberculosis, which resides in and destroys the lung tissue. The inflammatory response to control the infection results in the formation of granulomas in the lungs, which may impede respiratory function. Although the initial infection can be asymptomatic, it can result in fatigue, weakness, weight loss and fever. During reactivation, since the organism is able to lie dormant for many years, the patient may experience a cough, hemoptysis and chest pains (Norris, 1995). The weakened state of the affected individual may cause vulnerability to damage during coughing episodes.  Acute Respiratory Disorders  ARD can increase the work of breathing and reduce the efficiency of the respiratory system. In people with COPD it may further accentuate excessive workloads of their respiratory muscles that were already working at high demands. The chronic condition causes increased vulnerability  10  to acute disorders, which in turn leads to increased work of breathing, a decrease in the efficiency and force of the respiratory muscles (Figure 1). Acute respiratory failure in COPD can be due to any condition causing a further increase in the work of breathing and decreased respiratory drive. In respiratory failure, the patient can present with hypoxemia, hypercapnia or both. Factors causing acute respiratory failure can include the following: respiratory tract infections causing bronchospasms or accumulation of secretions; central nervous system depression resulting from trauma or medication effects; cardiovascular disorders such as myocardial infarction or pulmonary embolism; airway irritants such as smoke; thoracic abnormalities such as trauma, pneumothorax, haemothorax, and flail chest (Norris, 1995). Adult Respiratory Distress Syndrome (ARDS) also called "shock lung" is a form of pulmonary edema causing stiffening of the lungs and impairing ventilation. Some causes of ARDS include sepsis, trauma, drug overdose and near-drowning. Symptoms include rapid shallow breathing, dyspnea and the development of hypoxemia which increases the drive to breathe. Stiffening of the lungs results in the recruitment of accessory muscles for breathing (Norris, 1995). Accessory muscles would be vulnerable to injury since they would be unaccustomed to increased workloads. Pulmonary edema can be an acute or chronic condition and is defined as the build-up of fluid within the lungs. It reduces lung compliance. Pulmonary edema resultsfromleft ventricular failure which occurs due to arteriosclerosis, hypertension, cardiomyopathy or diseases of the heart valves. Symptoms include dyspnea on exertion, episodes of violent coughing at night, orthopnea, and tachypnea. In severe cases, breathing is laboured and rapid, diffused with coughing (Norris, 1995).  11  Cor pulmonale is hypertrophy of the right ventricle which results primarilyfromconditions affecting the lungs, lung vessels, chest wall or the medulla oblongata (the respiratory centre located in the brain stem). Respiratory disorders that result in cor pulmonale include the following: obstructive and restrictive lung diseases; extensive lung reduction surgery; chest wall disorders causing insufficient respiration without primary lung disease such as kyphoscoliosis, muscular dystrophy, and spinal cord lesions; obesity hypoventilation syndrome; and chronic mountain sickness from living in a hypoxic environment (Norris, 1995). Pneumonia is an acute infection of the lung parenchyma that disrupts the exchange of gases, and is the sixth leading cause of death in the United States. Cardinal signs of pneumonia are coughing, sputum production, chest pains, chills and fever. Respiratory complications that arise include hypoxemia, respiratory failure, pleural effusion, empyema, lung abscess and sepsis (Norris, 1995). People with CRDs are more vulnerable to acute lung infections which can over-burden the work of their respiratory muscles.  Other Factors Contributing to Muscle Injury In addition to the many respiratory conditions that can be associated with muscle injury, research has shown that corticosteriod therapy, aging and weight loss can also be contributing factors. These factors are often found in people with respiratory diseases which may further impede their respiratory function. Corticosteroids have been shown to cause muscle weakness and functional impairment (Dekhuijzen & Decramer, 1992). Corticosteriod therapy is often prescribed to patients with respiratory disorders in an attempt to control inflammation, however it can lead to respiratory muscle weakness (Decramer et al., 1994) depending on the dose, therapy delivery and type of  12  steroid (Reid & MacGowan, 1998). High doses of steriod administered over a short period of time can cause acute necrotising muscle pathology (Cumming et al., 1994). Interval steroid therapy was found to be related to weakening of the respiratory muscles (Decramer et al., 1994). Fluorinated steroids have been shown to cause more muscle abnormality than the non-fluorinated steroids (Reid & MacGowan, 1998). Aging is a progressive condition common to all organisms. The process of aging, as seen through animal models, shows more abnormal muscle morphology (Tolep & Kelson, 1993), a higher risk for injury (Watchko et al., 1994), and requires greater time to recover from injury (Brooks & Faulkner, 1990) in the diaphragm of older mice when compared to younger mice. Abnormal muscle morphology includes Z-band streaming and disruption of the cytoskeleton. Toleb and Nelson (1993) have described changes in aged muscle occuring in the myosin heavy chain, atrophied type 2b muscle fibres, and a reduction in capillary density. Effects of aging, which cannot be controlled, may further impede respiratory function in people suffering from respiratory conditions. Weight loss in COPD patients is very common, as the work for breathing increases, respiration requires greater amounts of energy than normal. When energy demands are not met through diet then protein catabolism occurs to be used as energy reserves (Li & Goldberg, 1976). Decreased respiratory muscle strength, in people with CRD, has been shown to be associated with protein catabolism (Ryan et al., 1993) a process that decreases muscle mass and function (Reid & MacGowan, 1998). In a catabolic state respiratory muscles may not be able to demonstrate a training effect to produce an adaptive muscle.  13  Respiration The essential function of respiration is the exchange of gases, oxygenfromthe atmosphere to our tissues and carbon dioxide from our tissues to the atmosphere. Inspiration is the action of breathing in, and expiration is the action of breathing out; these actions renew the air in our lungs. Respiratory muscles function efficiently every few seconds throughout our lives to enable respiration and life. The action of the muscles of inspiration create a negative intrathoracic pressure. As a result of the negative intrathoracic pressure, the lungs can inflate. The control of respiration lies in the respiratory centre located in the medulla oblongata of the brain stem.  Respiratory Muscles. Anatomy and Function: Diaphragm The diaphragm is a primary muscle of inspiration and is responsible for performing 70 to 80% of the work during quiet breathing (Loring & DeTroyer, 1985). Anatomically, the diaphragm is a large dome-shaped structure attached to the circumference of the thoracic cavity and separates the thoracic cavity from that of the peritoneal cavity. It consists of skeletal muscle around the circumference with a large central tendon region, which makes it unique in comparison to limb skeletal muscles. Diaphragm muscle fibers can be classified into three parts: sternal, which originates from the lower end of the sternum; costal, on either side, which originates from the lower six ribs; and crural, located posteriorly, which originate from the upper three lumbar vertebrae (De Troyer & Estenne, 1988) (Figure 2). The diaphragm is innervated by the phrenic nerve originating from the third, fourth andfifthcervical nerve roots (De Troyer & Estenne, 1988). Costal fibres are juxtaposed along the ribcage and run in a cranial orientation. This region of  14  juxtaposition is referred to as the "zone of apposition". In humans, this zone represents about 30% of the total surface area of the ribcage at rest (De Troyer & Estenne, 1988). During inspiration the diaphragm muscle contracts. Although its dome shape is maintained, contraction results in shortened costal muscle fibres, descent of the diaphragm towards the abdomen, and a decrease in the "zone of apposition". Shortened costal muscle fibres pull the ribs in an upward and outward direction, referred to as the "pump handle motion" and the "bucket handle motion", respectively (De Troyer & Estenne, 1988). Ultimately, inspiration enlargens the size of the thoracic cavity. As the chest expands, it creates a negative intrathoracic pressure, below atmospheric pressure, and the lungs expand. It also increases the intraabdominal pressure, which transmitted laterally causes an expansion of the lower ribcage.  Other Primary Muscles of Inspiration Primary muscles of inspiration include the scalenes and the parasternal intercostals (Loring & DeTroyer, 1985) as well as the diaphragm as mentioned above (Figure 3). The scalene muscles originate at the transverse processes of the lower five cervical vertebrae and insert into the upper surface of thefirstand second rib. The respiratory function of the scalene muscles, which are innervated via nerve roots originatingfromC2-C7 (Reid & Dechman, 1995), is to elevate the first two ribs during inspiration (DeTroyer & Estenne, 1988). The parasternal intercostal muscles are attached to the sternum and are intercartilaginous, running in a downward angle laterally from the sternum (Loring & DeTroyer, 1985). During contraction of the parasternal intercostal muscles, the ribcage is lifted and thereby increases the anteroposterior dimension of the thoracic cavity (DeTroyer & Estenne, 1988).  Accessory Muscles of Inspiration Some of the accessory muscles of inspiration include the sternocleidomastoid muscles, the internal and external intercostals, and the abdominal muscles (Figure 3). The sternocleidomastoid muscle originates at the mastoid process and its two heads are attached along the medial part of the clavicle and the ventral surface of the manubrium sterni (DeTroyer & Estenne, 1988). The action of the sternocleidomastoid during inspiration is to elevate the sternum which increases the anteroposterior dimension of the ribcage. The internal intercostals lie in the intercostal space and are attachedfromthe anterior to the posterior upper and lower borders of the ribs (Campbell & Davis, 1970). The external intercostals originate at the tubercles of the ribs and are attached to the costal cartilage where they fuse with the anterior intercostal membrane (Campbell & Davis, 1970). The function of the intercostal and external muscles during respiration is controversial, they may contribute to inspiration and/or expiration depending on lung volume (Taylor, 1960).  Muscles of Expiration Expiration, a passive motion during quiet breathing in healthy humans, occurs naturally by the inward elastic recoil of the lungs pushing the air out upon relaxation of the inspiratory muscles. These actions increase the pressure in the lungs above atmospheric moving air from the lungs to the atmosphere. During active expiration, such as during exercise, childbirth, coughing or sneezing, the abdominal muscles (rectus abdominis, external and obliques, and transverse abdominis) are recruited. Tone or contraction of the abdominal muscles maintains a firm abdominal cavity which also provides a fulcrum for diaphragmatic contractions. Contraction of the abdominal muscles, which attach to the lower ribs, reduces the dimensions of the ribcage and can also extend end-expiration (Loring & DeTroyer, 1985), increasing the time taken for  16  expiration. Therefore, abdominal muscles can act as inspiratory accessory and primary expiratory muscles. The internal intercostal muscles, as described above, are also considered to play a minor role in expiration (Taylor, 1960).  Respiratory Muscle Dysfunction in Respiratory Conditions In patients sufferingfromCRD, evidence of poor respiratory muscle function of the diaphragm has been observed (Begin & Grassino, 1991, Roussos, 1985). Respiratory muscle dysfunction, in humans, has been implicated as a cause of hypercapnic ventilatory failure (Begin & Grassino, 1991; Roussos, 1985), resulting from respiratory muscle fatigue, weakness or injury. Respiratory muscle fatigue is defined as a temporary decrease in the force generating ability of the diaphragm, which is reversible following rest (NHLBI Workshop Group, 1990). In general, muscles may become fatigued following a bout of exercise. However, if the exercised muscle is allowed to rest then it is able to generate normal forces following the period of rest (NHLBI Workshop Group, 1990). If the inspiratory muscles become exhaustively overloaded, as may occur in chronic respiratory diseases, then the respiratory muscles may become fatigued and damaged (Rochester, 1993). Unlike other skeletal muscles of the body that can reduce activity to promote healing, inspiratory muscles need to keep functioning in order to keep us alive. A weakened diaphragm can result in a state of hypercapnia or hypoxia which will further impede the force generating ability of the diaphragm (Reid & Dechman, 1995). In severe conditions of respiratory muscle injury, ventilatory failure can occur which can eventually lead to death (Rochester, 1993). Respiratory muscle weakness is defined as the decreased force generating ability of the diaphragm, which is persistent following rest (NHLBI Workshop Group, 1990). The diaphragm's  17  ability to rest is limited by our necessity to breathe and therefore has the potential for muscle weakness and a weakened muscle is unable to generate normal forces following rest (Reid & MacGowan, 1998). Respiratory muscle injury that manifests as structural abnormalities may be responsible for a decrease in the force generating ability of the diaphragm (Reid & MacGowan, 1998). Dysfunction of the respiratory muscles may resultfrommany contributing factors which can lead to an increased workload, a decreased efficiency and a decreased ability to generate force (Reid & Dechman, 1995; Figure 1). A decrease in the force generating ability of the diaphragm may resultfromthe following conditions: paralysis or neural damage of the nerve roots innervating respiratory muscles; connective tissue disorders; systemic abnormalities which include poor nutrition (Rochester, 1993; Tobin, 1988; Aurora & Rochester, 1982) and abnormal arterial blood gases (Jardim et al., 1981; Juan et al., 1979). The work of breathing for respiratory muscles can increase due to pathological changes within the lung tissue (hyperinflation), structural changes (barrel-chested), and a loss of compliance of the chest wall, placing the diaphragm at a mechanical disadvantage (Druz et al., 1979). Decreased efficiency of the chest wall or components of the respiratory system can increase the load on respiratory muscles. Flail chest resulting from multiple rib fractures can decrease the efficiency of the respiratory muscles, as it moves in paradox to the ribcage during inspiration and expiration. Hyperinflation can also increase the work of the respiratory muscles since respiration would require a greater amount of work to obtain normal amounts of tidal volume and to overcome gas trapping. Hyperinflation causes the diaphragm to perform its pumping action in a more flattened position that results in a shortened working length (Druz et al., 1979). Due to losses of sarcomeres in the diaphragm from working at shortened lengths, as reported by Kelsen et al., (1983) in hamsters, the length-tension  18  relationship may be maintained in humans but the flattened position will decrease its range of motion and contribute to a less than optimal function. Although decreased length may contribute to decreased mass, hypertrophy of fibres may increase mass. The demand to perform the work to maintain adequate ventilation may increase the diaphragm's susceptibility to fatigue, weakness or injury. Several other factors can contribute to poor respiratory muscle function. According to Reid and Dechman (1995), other injury promoting factors, which may contribute to respiratory muscle injury, include: aging, poor nutrition, and immobilization (i.e. if the patient is placed on a ventilator). Patients with COPD experience exercise intolerance, dyspnea, hypercapnic ventilatory failure (Reid & Dechman, 1995) and a decrease in their quality of life (McSweeney & Labuhn, 1990) which may be related to respiratory muscle dysfunction. In severe conditions of respiratory muscle dysfunction and injury, ventilatory failure can occur which can eventually lead to death (Rochester, 1993; Rochester, 1991). Ventilatory failure, which is the inability to ventilate adequately, is defined by the PaC0 value being greater than 45 rnmHg (hypercapnea) 2  and a Pa0 of less than 55 rnmHg (hypoxaemia) (Roussos & Zakynthinos, 1995; Rochester, 2  1993). Hypercapnic COPD patients have an excessive load on their respiratory muscles which leads to muscle fatigue (Reid & Dechman, 1995) and cytoskeletal disruption (Reid & MacGowan, 1998).  Respiratory Muscle Injury Muscle Injury: Skeletal muscle has an incredible ability to accommodate and adapt to changes in workload which for the healthy subject means increased activity levels. In general, the cycle of skeletal  19  muscle adaptation leads to a training effect. Various levels of change are experienced during the process of adaptation which include muscle fatigue and /or muscle weakness, recovery, and finally a muscle that is adapted to the workload. Part of this adaptation includes the inflammatory response (Cannon & St. Pierre, 1998) (Figure 4) and delayed onset muscle soreness (Maclntyre et al., 1995) likely experienced from muscle injury. Adaptation may involve the reconfiguration of the normal muscle fibre in order to rebuild or reconstitute a muscle fibre that is able to perform and handle the increased workload (Clarkson et al., 1992). Hence the trained person is able to perform tasks with ease which may have presented a challenge initially, such as running and being able to increase the distance and/or speed. If the training workload is not maintained muscle fibres may become deconditioned to their original state. The cycle of muscle adaptation is reinitiated if the workload was to be increased again in a training pattern of loading and unloading. Injury to skeletal muscle fibres can occur as a result of various forms of insults, such as exercise, trauma, and extreme temperatures (Faulkner et al., 1993). Following exercise, muscle injury has been reported to be localized, which involves a few sarcomeres, or extensive which can include larger regions of muscle (Friden et al., 1983). Muscle injury to skeletal muscles has been reported to occur after three types of muscle activity (Faulkner et al., 1993; van der Meulen et al., 1991; McCully & Faulkner, 1985; Newman et al, 1983) which differ according to the muscle length change during tension development. Firstly, concentric contraction occurs when the muscle shortens during the contraction. Secondly, an isometric contraction occurs when the muscle contracts without changing length. The third is an eccentric contraction which occurs when the muscle lengthens during the contraction. Eccentric activity may generate greater tension per unit area because fewer muscle fibres are recruited. Therefore, more muscle injury is  20  observed following eccentric activity in comparison to concentric and isometric ( Evans & Cannon, 1991, Stauber et al., 1988; McCully & Faulkner, 1985). According to Faulkner et al. (1993), eccentric contractions may result in localized injury, affecting a single sarcomere and adjacent myofilaments. Respiratory musclesfromCOPD patients has shown evidence of muscle damage and cytoskeletal disruption (Reid & MacGowan, 1998). In the diseased state a muscle may experience normal functioning as an increased overactivity and/or overload. Overactivity is described to occur when the firing rate, to induce muscle contraction, exceeds normal parameters and overload occurs when the force required exceeds normal parameters (Stauber & Smith, 1998). In COPD, breathing becomes shallow, rapid, and laboured demonstating both overactivity and overload. To provide a better understanding of the changes that occur in injured muscle, the following section will briefly describe normal skeletal muscle structure.  Normal Skeletal Muscle Structure Skeletal muscle, as in the diaphragm, is composed of striated muscle as defined by the presence of alternating dark and light bands within the musclefibres(Figure 5). Long multinucleated muscle fibres lie in parallel to each other. When the muscle is active, fibres contract along its longitudinal axis. Each single musclefibre,bundle of fibres, and entire muscle is surrounded by connective tissue layers, the endomysium, perimysium and epimysium, respectively. The muscle cell membrane, which surrounds the sarcoplasm is called the sarcolemma and is found beneath the endomysium (McArdle et al., 1991). The inner layer of the sarcolemma is referred to as the plasmalemma and the outer layer is known as the basal lamina.  21  The basal lamina of the muscle consists of loose glycoprotein and a collagen network (Jones, 1995) (Figure 6). Fibroblasts are located between muscle fibres and one of their functions is to produce collagen. The collagen fibres and fibroblasts are major components of the connective tissue that surrounds the various levels of muscle, as mentioned above. Connective tissue serves to provide structure and integrity to the muscle (Copenhaver et al., 1971) and also serves to translate force across and along the longitudinal axis of the muscle. The functional unit of the musclefibreis the sarcomere (Figure 5, Figure 7). In mammalian muscle, the sarcomere is defined as the region from one Z line to another, which is about 2.0 to 2.5 um. Myosin (thick) and actin (thin) are myofilaments that are present in the sarcomere and it is their cross-linking that results in muscular contraction. According to the sliding filament hypothesis, specific sites located on the thick and thin filaments combine and generate tension by pulling thefilamentstowards the centre of the A-band (Huxley & Niedergerke, 1954). As illustrated in Figures 5 and 7, there are regions within the sarcomere that consist of only myosin or of only actin and are labeled as H and I zones, respectively. The region where myosin and actin overlap is referred to as the anisotropic or A-band (McArdle et al., 1991). The Z line, to which the actinfilamentsare associated, appear to bisect the mid-region of the isotropic or I-band (Reith & Ross, 1977) and connects to the sarcolemma providing the muscle structure with stability (McArdle et al., 1991). Proteins that compose the M line appear to bisect the centre of the sarcomere (Copenhaver et al, 1975) and are responsible for maintaining myosin alignment during sliding of the actinfilaments(McArdle et al., 1991). Each thickfilamentis surrounded by an hexagonal array of six thinfilaments(Figure 8). At the Z lines, the thinfilamentsare arranged in a square lattice array (Copenhaver, et al., 1971) (Figure 9). Spiraling projections or myosin heads are found on the portion of the myosin which  22  overlaps the actin filament. In the electron microphotographs, the spiraling projections are not obvious in appearance since they are much thinner than the actin (thin) filaments which are about 50 angstroms in diameter (McArdle et al., 1991). The sarcoplasmic reticulum (SR) is a complex network of interconnecting channels that surrounds each myofibril (Jones, 1995; McArdle et al., 1991). The function of the SR is to store and release calcium. It is the presence of calcium ions that stimulates cross-bridge formation of the myofibrils. Calcium ions are stored in the sarcoplasmic reticulum which is made up of a complex arrangement of tubules and cisternae. The SR is orientated along the long axis of the myofibrils. As the tubules approach the A-I junction, they fuse to form large terminal cisternae (in man). The transverse tubule system or T-system which are invaginations of the sarcolernma is situated in close proximity to the SR. The T-system is an intricate network of channels which makes contact with the sarcomere twice along the length of every myofibril, approximately at the junction of the A- and I-band in man (Jones, 1995). The organization of the T-tubule with the terminal cisternae is known as a triad (Jones, 1995, McArdle et al., 1991). The triad serves as a rapid transportation system that delivers the action potential from the outer regions of the sarcolernma to the inner regions of the terminal cisternae in the muscle cell and hence, calcium release from the SR is initiated (McArdle et al., 1991). The role of mitochondria in cells is to provide the energy source. In muscle, mitochondria are located between the myofibrils in the region adjacent to the I-band or Z line (Rhodin, 1974) and appear as doublets along the Z line (Tomilin et al., 1991) (Figure 10). Clusters of mitochondria are also found at the periphery of the muscle cell at close proximity to the nucleus and in small areas beneath the cell membrane (Tomilin et al, 1991) (Figure 11). Mitochondria have a double membrane where the inner membrane protrudes inwards and forms folds or tubes known as  23  cristae. A cross-sectional view of a mitochondria (Figure 10) reveals that the cristae do not extend all the way across the mitochondria and that they can vary in number and form (Copenhaver et al., 1971). Mitochondrial shape is not consistent; they can be elongated or rounded (Tomilin et al., 1991). Also present in the muscle cells are glycogen particles. Glycogen stores in the muscle provide a source for the energy and in the electron microphotographs, appear as clusters of dense particles due to the lead staining involved in the tissue processing. In active muscle, glycogen stores can be depleted. Lipid droplets are also a source for energy and exist as sarcoplasmic inclusions. A rich capillary network system ensures that an adequate supply of oxygen and nutrients are available to each musclefibre(Copenhaver et al., 1971). Skeletal musclefiberis made up of two types of musclefibers,type 1 and 2 (Figure 11). Fibers are typed according to their contractile and metabolic features (Reid, 1995; McArdle et al., 1991). Type 1fibers,also known as slow twitch or oxidative, contain a large supply of mitochondria which enables thefiberto perform long-term aerobic activity. Type 2fibers,also called fast twitch or fast glycolytic, work anaerobically to deliver energy quickly over a short-term and have few interfibrillar mitochondria. Type 2 fibers can be further categorized as type 2a i.e. fast-oxidative glycolytic and 2b, fast glycolytic (McArdle et al., 1991) and type 2d or 2x (Reid & Dechman, 1995). Normal skeletal muscle of the diaphragm is shown in Figure 12. This photomicrograph of an eight micron thick cross-section taken from the costal region of the diaphragm is typical of normal muscle observed through the light microscope. The haemotoxylin and eosin stain distinguishes nuclei and cytoplasm by different colours. Nucleus appear blue in colour, cytoplasm appears pink, and connective tissue as a pale pink colour. In muscle cross-section, one musclefibremay  24  have up to three nuclei located within the periphery of the fibre beneath the sarcolemma (Cumming et al., 1994). The skeletal muscle cell has a polyhedral shape and its diameter can vary from 10 um (from an occular muscle) to 100 um (from the gastrocnemius muscle) (Copenhaver et al., 1975). Muscle fibres from a healthy individual are thicker thanfromsomeone who is malnourished (Copenhaver et al., 1975). Located within the extracellular matrix are blood vessels and nerves, connective tissue with stellate shaped fibroblast cells which have a spindle shaped nucleus, and residential white blood cells, such as macrophages (St. Pierre & Tidball, 1994).  Muscle injury-Light Microscopic Observations Light microscopic examination of injured diaphragm muscle in animals, where the injury was induced by direct stimulation (Reid, 1988), phrenic nerve stimulation (Dechman et al., 1996), or tracheal banding (Reid et al., 1994; Reid, 1993), has shown damaged muscle fibres of the diaphragm and cellular infiltrates into the muscle tissue. Increased areas of abnormal diaphragm muscle were found by performing point counting on haemotoxylin and eosin stained crosssections. Abnormal muscle is characterized as fibres with a central nucleus, degenerative or nonuniform cytoplasm, and infiltration of inflammatory cells within the muscle fibre or in the interstitium (Reid et al., 1994; Figure 13). Microscopic examination of injured muscle morphology reveals moth-eaten fibres and Z-band streaming or broadening (Hards etal., 1990; Campbell et al., 1980). Muscle nuclei that have moved awayfromthe periphery and appear towards the centre of the fibre may represent a muscle response to a stimulant (Cumming et al., 1994). Changes in the ultrastructure of the muscle seen following post exercise include: cytoskeletal disruption; muscle cells infiltrated by inflammatory cells (Lieber, 1992; Ebelling & Clarkson, 1989); abnormalfibreshapes and sizes (Reid & MacGowan, 1998); increased  25  connective tissue (Reid & MacGowan, 1998; Reid, 1995). Reid (1995), using a tracheal banding model to induce a load on the diaphragm, reported greater muscle abnormality and inflamed muscle after 3 days of banding than after 30 days  Muscle Injury-Electron Microscopic Observations Disorganization of myofilaments and Z line displacement, also known as streaming or broadening, are common findings observed in electron microphotographs of injured muscle tissue (Reid et al, 1994; Faulkner et al., 1993; Friden et al., 1983). In tracheal banded rats, the costal region of the diaphragm showed areas of disrupted myofilaments, zonal lesions, and a loss of distinct A and I bands (Reid et al., 1994) (Figure 14). Zonal lesions refers to areas that show hypercontraction of sarcomeres and adjacent regions where sarcomeres appear stretched. Friden et al. (1983) also showed cytoskeletal dysruption and mitochondrial swelling.  Inflammation Inflammation is a conserved, natural occurrence of the body in response to tissue injury and microbial invasion (Tidball, 1995). The inflammatory response is associated with muscle injury (Figure 4). Cardinal signs of inflammation are: calor (warmth), tumour (swelling), erythema (redness), and pain. Inflammatory cells and mediators contribute to these changes which stimulate an increase in blood flow, and an increase in vascular permeability leading to redness, swelling and pain. Inflammatory cells are signalled to initiate the repair process following an injurious insult. The insult can be brief such as during a bout of exercise, or it can be prolonged such as during chronic respiratory disease. Histologically, the acute stage of inflammation is characterized by the presence of neutrophils, and the chronic stage of inflammation is  26  characterized by the infiltration of monocytes (Armstrong, 1990). Although injury may persist, more often healing and resolution are dominant processes during the chronic phase. Macrophages dominate during the healing and resolution phase (St. Pierre & Tidball, 1994; McLennan, 1993) which may begin within a few hours following injury (Orimo et al., 1991) and can persist for days. Inflammatory mediators There is a multitude of mediators involved in inflammation and tissue repair. Understanding the interrelationships of the huge number of inflammatory mediators that are responsible for stimulating inflammatory cells (such as neutrophils, monocytes, macrophages, endothelial cells, mast cells, etc.) allows one to appreciate the complexity of the inflammatory response. Mediators can act as chemoattractants which attract PMNs and other phagocytic cells which also produce cytokines to further enhance or amplify the response known as the cascade effect. In an attempt to appreciate the complexity of inflammation, a few mediators will be further described. Interleukin 1 (IL-1) and tumour necrosis factor (TNF) are two of the primary inflammatory mediators. Other mediators which also play a major role in inflammation are: C5a, a factor from the complement system, arachidonic acid metabolites, histamine, proteases, and lysosomal enzymes (Cannon & St. Pierre, 1998; Roitt et al., 1989; Ryan, 1970). IL-1 is a potent mediator of inflammation. A few of the properties of LL-1 include being a chemotactic agent for PMNs, lymphocytes and monocytes, and it increases leukocyte adherence to vascular endothelium. IL-1 can also stimulate eosinophil degranulation, fibroblast proliferation, collagenase production, and the arachidonic acid cascade. A large number of cells have the capability to produce IL-1 such as monocytes/macrophages, dendritic cells, B cells, fibroblasts, epithelial cells, and endothelial cells (Roitt et al., 1989). TNF, a cardinal mediator in the inflammatory response, is predominantly produced by  macrophages. JJL-1 and TNF have overlapping biological properties which include: increased hepatic acute phase protein synthesis, activation of endothelium, increasedfibroblastproliferation, and increased synovial cell collagenase and PGE levels (Roitt et al., 1989). 2  The complement system has many functions in the acute inflammatory reaction (Roitt et al., 1989). C5a, a fragment from the complement system, is one of the most important chernotactic peptides for attracting neutrophils and macrophages. In acute inflammation, C5a predominantly attracts neutrophils (Roitt et al., 1989). The inflammatory effects of the complement components includes neutrophil chemotaxis and activation, increased vascular permeability, smooth muscle contraction, mast cell degranulation, and also antibacterial effects. Histamine, a vasoactive amine, is released in response to physical injury, chemical agents and immunological processes. Histamine is liberated by the degranulation of mast cells, basophils and platelets. The action of histamine also induces contraction of smooth muscle and increases vascular permeability (Ryan & Mayno, 1970). Arachidonic acid metabolism (Ryan & Mayno, 1970) is one of the most important systems in the biology of inflammation. Arachidonic acid is found in abundance in an esterified form in membrane phospholipids of a large number of cell types. When activated for example by membrane disruption, arachidonic acid is released by being cleaved from the phospholipids in response to phospholipases. Ultimately, this cleavage leads to the release offreearachidonic acid which can now be acted upon by two metabolic systems: cyclo-oxygenase and lipoxygenase. Cyclo-oxygenase leads to the production of prostenoids which act as chemoattractants for mast cells and PMNs, and also increases the adherence of PMNs and monocytes to vascular endothelium. Lipoxygenase, the other system involved in the arachidonic acid metabolism, leads to the production of hetes and leukotrienes, which also serve as chemoattractants for neutrophils.  28  Acid proteases,fromneutrophil lysosomes, are able to breakdown basement membranes and act as inflammatory mediators (Ryan & Mayno, 1970). Neutral proteases, also released by neutrophils, cause tissue degradation. It is believed that neutral proteases can lead to the production of fragments of C5, thereby increasing the presence of inflammatory mediators. Release of lysosomal enzymes, outside of the cell, can also augment tissue damage and amplify the inflammatory response. Activation of a non-lysosomal calcium activated neutral protease, calpain, is involved in exercise induced muscle injury (Jiang et al., 1998a). It acts as a chemotactic agent by the fragments it creates during protein degradation but also as an agent responsible for cytoskeletal dysruption (Belcastro et al., 1998). Calpain is found in association with membranous structures such as phospholipids, ion transport channels and receptors (Johnson, 1990). Within the sarcomere, calpain is associated with the I band and the Z band. Cytoskeletal proteins such as actin-binding and microtubule associated proteins have been recognized as some of the substrates for calpain degradation (Saido et al., 1994). A loss of calcium homeostasis has been reported with exertion induced muscle injury (Armstrong, 1990) which likely activates calpain during the autogenic stage.  Cellular Response Cell types involved in acute and chronic inflammation tend to be different. Cell types involved in acute inflammation are usually PMN leukocytes, these can be neutrophils, eosinophils or basophils, but macrophages and monocytes may also be involved. Chronic inflammation tends to involve more mononuclear cells which are principally macrophages, T cells, and B cells. Very few neutrophils or other polymorphocytes are usually involved in this stage (Ryan & Mayno, 1970).  29  Neutrophils Neutrophils make up 90% of the circulating PMN granulocytes, the remaining 10% is made up of eosinophils and basophils. Granulocytes are produced in the bone marrow at a rate of 80 million per minute, compared to eight hundred thousand lymphocytes per minute. Neutrophils only survive for two to three days (Roitt et al., 1989). As the name suggests, PMN granulocytes have a multi-lobed nucleus and have many granules. The neutrophil is produced in the bone marrow and is 10 to 20 um in diameter (Roitt et al., 1989). Neutrophils are the primary cells in an acute inflammatory response at sites of injury. During the inflammatory response, the main role of neutrophils is chemotaxis to sites of antigen entry or complement activation, and degradation and phagocytosis of damaged tissue. At maturation, neutrophils enter the bloodstream where they exist in circulating and marginated pools (Cartwright et al., 1964). Margination is the adhesion to endothelial cells (Roitt et al, 1989). The number of neutrophils in circulation can increase drastically by decreasing those present in the marginated pools and those stored in the bone marrow. Some of the stimulants that can increase circulating neutrophils include an increase in blood flow (Schmid-Schoenbein et al., 1980), exercise and inflammatory mediators (Golde, 1990). An increase in neutrophils is achieved by several factors such as: decreased maturation time, decreased time spent in the bone marrow by mature neutrophils, stimulation of immature and mature neutrophils to enter the bloodstream (Cronkite, 1988), and also stimulation of the marginated pool. During circulation, neutrophils respond to inflammatory mediators by migrating along a chemotactic gradient. As mentioned earlier there are many activating agents of neutrophils which include: cytokines, complement fragments i.e. C5a, endotoxin, arachadonic acid metabolites (Territo, 1981). It has been shown that there is an increased presence of circulating neutrophils in  30  the plasma (Camus et al., 1992) and skeletal muscle (Fielding et al., 1993) following downhill running either immediately (Camus et al., 1992) or 45 minutes post-exercise (Fielding et al., 1993). Evans et al. (1991) has shown an increase in circulating neutrophils during and for several hours post-exercise. Neutrophils have been found to contibute to myocardial injury after reperfusion of ischaemic tissue (Eppinger et al., 1995, Entman et al, 1991, Lucchesi et al, 1989).  Myeloperoxidase The enzyme myeloperoxidase is the most abundant protein present in the neutrophil and is contained in the primary or azurophilic granules. During phagocytosis, activated neutrophils show metabolic changes that increases oxygen consumption (Ryan & Mayno, 1970). This occurs to support a primary function of defense by neutrophils to produce oxygen radicals which are cytotoxic to cellular membranes. The respiratory burst system produces hydrogen peroxide and oxygen intermediates. Activation of glucose-6-phosphate and NADH oxidase, another enzyme in the respiratory burst pathway, produces unstable oxygen radicals. MPO acts as a catalyst during the cleavage of hydrogen peroxide to form a highly reactive oxygen radical which in turn is responsible for the production of hypochlorous acid. Hydrogen peroxide can be broken down to form hydroxyl radicals and superoxide, which are highly reactive oxygen intermediates. If a halide group, such as chloride, is added to the hydrogen peroxide in the presence of MPO, this results in the production of hypochlorite which has degradative capabilities and an antimicrobial function (Ryan & Mayno, 1970). Quantification of the MPO protein provides a relative marker for neutrophils i.e. indicative of the presence or absence of neutrophils according to the amount of reactive MPO. An increase of MPO in the muscle following exercise has been shown in downhill running (Fielding et al, 1993)  31  and strenuous swimming (Morozov, et al, 1990). Other cells that contain the enzyme peroxidase include granulocytes (eosinophils and basophils) and monocytes/macrophages (Roitt et al., 1989)  Macrophages Macrophages play a more dominant role in chronic inflammation. Macrophages were believed to be primarily scavengers for debrisfromdamaged tissue (Cline, 1968) but are now also considered to be modulators of the inflammatory response (Canon & St. Pierre, 1998; Dijkstra et al, 1985). Tissue macrophages originate from monocytes, which are derivedfrompluripotent stem cells in the bone marrow and are 10 to 18 um in diameter. Macrophages are mobile, phagocytic cells which contain a large variety of enzymes and secrete products that act as mediators in the inflammatory reaction. In addition to their role in the inflammatory response, macrophages provide ongoing removal of cellular debris as a result of cell apoptosis (Roitt et al, 1989). Macrophage sub-populations have been investigated in skeletal limb muscles to reveal an association with myonecrosis and muscle regeneration following muscle injury (Tidball, 1995; Orimo et al, 1991). Other studies performed on injured nerve tissue supports the critical role of macrophages with respect to the degenerative and regenerative processes following injury (Venzie et al, 1995, Dahlin, 1995). Dahlin (1995) used a milipore chamber to prevent the invasion of macrophages in nerve grafts to demonstrate that if macrophages are preventedfrominfiltrating an area of injury, it impairs the regenerative process. It is apparent that in order to increase the healing resolution in the injured muscle, the cellular debrisfromdamaged fibers needs to be removed. Some of the mediators of macrophages are similar to those discussed for neutrophils.  32  However, unlike neutrophils, macrophages are not primary inflammatory cells at a site of injury. Some of the inflammatory mediators for macrophages are EL-l, TNF, and fragments of collagen and fibronectin (Roitt et al., 1989).  Phagocyte Function A property of all phagocytic cells is chemotaxis. In fact, even tumour cells have chemotactic properties; they actually metastasize to a site where there is an inflammatory stimulus (Bevilacqua & Nelson, 1993; Dertlagopian et al., 1978). Chemotaxis is the locomotion of a cell along an increasing concentration gradient of chemotactic substances detected by receptors on leukocytes. Phagocytes arrive at a site of inflammation by following a concentration gradient of chemotactic substances. Chemotactic substances can be derived from many sources such as the complement cascade (eg. C5a), peptides (eg. N-formyl peptide), kinins, and arachidonic acid metabolites. Monocytes and PMNs are normally in the vascular compartment and in order to get to a site of injury or infection they first have to adhere to the vascular endothelium (Figure 4). To adhere neutrophils first roll, then tether, finally forming a firm adhesion by adhesion molecules. Once these cells adherefirmlyto the endothelial cells, they become ameboid and squeeze through the spaces between the endothelial cells. The process of transmigration across the endothelium is known as diapedesis, which enables neutrophils to migrate into the tissue towards the site of injury (Roitt et al., 1989; Ryan & Mayno, 1970) (Figure 4). The ingestion phase follows after the engulfment and internalization of the particle to be degraded by creating a phagosome. The phagocytic activity of neutrophils is to release cytotoxic agents such as myeloperoxidase (MPO) (Marquez et al., 1990; Miyasaki et al., 1991; Morozov et al., 1990; Nauseef, 1990), elastase and collagenase (Camus et al., 1992; Evans & Cannon, 1991) into the phagosome. These cytotoxic  33  agents are able to degrade phagocytosed materials. However, adjacent healthy tissue may also be degraded when the phagocytic cells ruptures or degranulates before the phagosome has completely formed, thereby releasing lysosomal enzymes outside of the cell (Klebanoff, 1971).  Identification of Neutrophils and Macrophages Neutrophils can be identified by the function of the enzyme MPO. MPO is found in the azurophilic granules within the neutrophil. The enzyme is able to cleave the substrate hydrogen peroxide resulting in the production of a superoxide radical, which reacts with the trapping agent diaminobenzidine. As a result of the chemical reaction a dark precipitate can be distinguished under a light microscope. Populations of macrophages can be distinguished using monoclonal antibodies. EDI, ED2, la (McLennan, 1996; St Pierre & Tidball, 1994; Dijkstra, 1985), and ED3 are monoclonal antibodies which specifically recognize cell markers of rat tissue macrophages (Dijkstra, 1985). The monoclonal antibody EDI is used widely as a marker to recognize the inflammatory macrophage which is associated with muscle necrosis (Figure 15). The antigen is localized on cytoplasmic granules like phagolysosomes, as well as on the cell surface. Phagocytic activity of the ED1 cell +  has been correlated to the increased expression of this antigen (Damoiseaux et al., 1994). ED2 and ED3 antibodies recognize and bind to membrane markers on the resident tissue macrophages, which are associated with muscle regeneration (McLennan, 1996; Dijkstra, 1985). Macrophages expressing the class II major histocompatibility antigen la (OX-17), are found in the connective tissue in adult rat muscle (Honda et al., 1992). Muscle injury induced by eccentric contractions has shown the infiltration of Ia macrophages into the myofibres (Stauber et al., +  1988) and this macrophage subpopulation is known to be phagocytic (Beller & Unanue, 1981).  34  Acute Phase Response The acute phase response is a cytokine dependent event that involves the liver, pituitary, adrenal glands, and the cells that make the cytokines in the first instance (Roitt et al., 1989). Acute phase response occurs during a systemic inflammatory response. PMNs, macrophages, and other cells are activated and produce IL-1 and TNF in response to such noxious insults as trauma, microbial invasion, or immunological stimuli. These mediators act on hepatocytes in the liver. Hepatocyte stimulating factor, which consists of JL-1, TNF and EL-6, stimulates liver cells to produce acute phase proteins. Acute phase proteins serve to promote inflammation and also serve to downregulate inflammation. At the same time, during the acute phase response, these cytokines, in particular EL-l and TNF, go to the central nervous system where they stimulate the pituitary to release the adrenocorticotrophic hormone (ACTH). The primary role of ACTH is to stimulate the adrenal glands to release a corticosteroid. Steroids play a prominent role in host response to: trauma, injury, stress, inflammation, and infection by reducing shock (Ryan & Mayno, 1970).  35  HYPOTHESIS Neutrophils mediate and may amplify some of the diaphragm muscle injury observed after resistive loading induced by tracheal banding. We hypothesize that neutrophil depletion will reduce diaphragm muscle injury in tracheal banded rats, and reduce the infiltration of inflammatory macrophages.  Specific Aims: The specific aims of this study were: 1. to examine the amount of diaphragm muscle injury in the tracheal banded plus neutrophil depleted rats compared to the tracheal banded, control, or control plus neutrophil depleted rats; 2. to measure arterial blood gases daily to determine the hypercapnic state of the tracheal banded rats compared to the control rats; 3. to determine the number of neutrophils and inflammatory macrophages present in the costal region of the diaphragm in the tracheal banded plus neutrophil depleted groups versus the tracheal banded, control, and control plus neutrophil depleted groups; 4. to determine the amount of inspiratory resistive loading induced by tracheal banding by measuring tracheal cross-sectional areas and esophageal pressures; 5. to examine the changes in circulating neutrophils in all animal groups to ensure neutrophil depletion induced by the administration of antisera was adequate.  36  tWork  ^Strength Neuromuscular  j Efficiency  Airways  Disorders  Flail Chest  Obstruction  Systemic  Chest Wall  Abnormalities  Compliance  Hyperinflation  Respiratory Muscle Overload  I Respiratory Muscle Fatigue  s  Recovery<  \.  ^Respiratory Muscle Injury  Adaptation  Figure 1: Hypothetical model of respiratory muscle injury. Reprinted with permission (Reid & Dechman, 1995).  37  Sternum  Sternal Region Lower Costal Border 8M  Central Tendon Inferior Vena Cava  ^=1— Costal Region Esophagus Aorta  ,yr-^ } s  cwalRegion  Lumbar Vertebrae  Figure 2: Diaphragm muscle viewed from below. Reprinted with permission (Reid & Dechman, 1995).  38  Expiratory Muscles  Inspiratory Muscles itemoeleldomastoid Scalenus •Trapezius  Internal Intercostals  Parasternal Intercostals  External Intercostals  iaphragm Rectus Abdominis External Oblique  Internal Oblique Transversus Abdominis  Figure 3: Respiratory muscles of the thorax. Reprinted with permission (Reid & Dechman, 1995).  39  Figure 4: Hypothetical inflammatory response to muscle injury. Reprinted and modified with permission (Maclntyre et al., 1995)  Musdefibeni--'  light T>ark I band Aband A I band band " ° ' . *jn~  H  zone  , w  Actin thin filament  .V.*.  v.-.v  .W.  Hto  Myosin thick filament  Myofilaments (cross section)  Figure 5: Organization of skeletal muscle. Adapted from Vander (1985).  Figure 6: Electron microphotograph of skeletal muscle fibres (cross-section). The muscle fibre is surrounded by the sarcolernma, which consists of the inner layer, plasmalemma (PL), and the outer layer, the basal lamina (BL). Muscle fibres are multi-nucleated (N). (x66 000).  Figure 7: Electron microphotograph of a sarcomere, which is defined as the region from one Z-line (Z) to another. The H zone (H) consists of only myosin myofilaments and the I band (I) consists of only actin filaments. The M line (M) bisects the centre of the sarcomere. The region where the actin and myosin overlap is labelled as the A band. (x63 000).  Figure 8: Electron microphotograph illustrating the hexagonal array of myofilaments as seen at the cross-section of an A band. Each myosinfilament(My) is surrounded by six actin filaments (A). In the upper right portion of the micrograph is a cross-sectional view of actin filaments cut at the I band region. The sarcoplasmic reticulum (SR), which is a complex network of interconnecting channels, is also visible. (x59 500).  Figure 9: Electron microphotograph of Z lines cut 'en face'. Note the square lattice array of the Z line shown at a high magnification (x85 000).  Figure 10: Electron microphotographs illustrating the doublet array of mitochondria (m) along the Z line (x36 000). Inset: Cross-sectional view of a mitochondria. Note the double membrane and the folded appearance of cristae labelled c (xl05 000).  Figure 11: Electron microphotograph showing fibre types and mitochondrial distribution. Fibre type 1 (I) has numerous mitochondria (m), fibre type 2a (Ila) has fewer, and fibre type 2b (lib) has an intermediate amount present. In the micrograph an interfibrillar blood vessel (bv) is present and also lipid droplets (L) are visible within the muscle fibres. (x6 250)  47  Figure 12: Photomicrograph of normal rat diaphragm stained with H&E, staining cytoplasm pink and nuclei blue. Muscle fibres have a polygonal shape and are multinucleated with nuclei located in the periphery of the muscle fibre (arrow with white arrowhead). Each muscle fibre is surrounded by connective tissue made by fibroblasts with spindle shaped nucleus (black arrow). A network of capillaries (black arrowhead) supplies the muscle fibres with oxygen and nutrients. Scale bar =10 urn.  48  F i g u r e 1 3 : P h o t o m i c r o g r a p h s of inflamed a n d d a m a g e d m u s c l e fibres in the d i a p h r a g m t a k e n f r o m a t r a c h e a l b a n d e d rat s t a i n e d with H & E . F i g u r e A illustrates a necrotic m u s c l e fibre without a cell outline. T h e m u s c l e fibre h a s b e e n d e g r a d e d w h e r e only r e m n a n t s of c y t o p l a s m (black a r r o w h e a d ) a r e d i s t i n g u i s h a b l e a m o n g s t inflammatory c e l l s a n d e f f u s i o n . F i g u r e B s h o w s f e a t u r e s of a b n o r m a l i t y , s u c h a s m u s c l e fibres with internal n u c l e i (arrow) a n d i n f l a m m a t i o n , s u c h a s i n c r e a s e d interstitial s p a c e (white a r r o w h e a d ) , a n d a n influx of n u m e r o u s r o u n d e d n u c l e i (arrow with white a r r o w h e a d ) . S c a l e b a r = 1 0 jam.  Figure 14: Electron microphotograph of diaphragm from a tracheal banded rat, showing abnormal muscle. Note the myofibrils lacking parallel alignment, presence of vacuoles (V), and Z-line streaming (Z). (xl8 000).  50 Figure 15: Photomicrographs of ED1+ macrophages in the diaphragm cross-section of tracheal banded and neutrophil depleted rats. Macrophages have been stained with fuschin red using immunohistochemistry and the cross-sections were counter-stained with Mayer's haemotoxylin solution. Panel A shows an overview of the muscle cross-section and the overall distribution of ED1+ macrophages at a 40X magnification. Scale bar =25 um. Panel B is a 100X magnification with oil immersion of the boxed areafromfigure 4A. The EDI antibody is primarily a cell surface marker, which targets inflammatory macrophages (black arrow). Other inflammatory cells (arrow with white arrowhead) such as monocytes or neutrophils may also be present in the interstitium and have not been identified by the EDI antibody. Scale bar =10 pm  51  52  CHAPTER 2 METHODS Animals and Groups Adult (9-10 weeks) male Sprague Dawley rats, weighing approximately 350 g were randomly assigned to the following groups: control (C), n=T3; control and neutrophil depleted (C+ND), n=12; tracheal banded (TB), n=12; and tracheal banded and neutrophil depleted (TB+ND), n=12. The numbers of animals per group, as mentioned above, completed the protocol and met the criteria of the intervention. The animals were housed individually and fed a diet of Purina Laboratory Chow and water ad libitum. The care and use of the animals were according to the guidelinesfromthe "Guide to the Care and Use of Experimental Animals" (Canadian Council on Animal Care, 1980). The inclusion criteria for the rats were as follows: 1. TB groups had to have a PaC0 greater 2  than 45 rnmHg; and 2. ND groups had to have an absolute PMN count less than 0.5 million/mL. Animals that did not meet the criteria were excluded. In addition to the numbers that had completed the protocol, the following rats did not meet the inclusion criteria. Three animals were excludedfromthe TB group whose PaC0 was less 2  than 45 rnmHg. Seven animals were excludedfromthe ND group because their PMN counts were greater than 0.5 million/mL. 14 animals died prematurely because the animal either stopped breathing during esophageal pressure (Pes) measurements and we were unable to revive them (n=5), or were found dead in their cage (n=7), or due to surgical complications, such as during carotid artery cannulation (n=2).  53  Experimental Protocol One day pre-op, both ND groups (C+ND and TB+ND) received intra-peritoneal (i.p.) injections of anti-rat neutrophil antisera. On day 1, animals were anaesthetized with the following anaesthetic regimen: valium (1.5 mL/Kg) -intra-muscular (i.m.), ketamine (0.8 mL/Kg) - i.m., and glycopyrolate (0.375 mL/Kg) - i.p. injection. Animals were then prepared for surgery by shaving the site of incision and then cleansing it by applying Betadine, a 10% povidone-iodine topical solution (Purdue Frederick Inc, Toronto, Canada). The surgical protocol consisted of firstly, a carotid artery cannulation procedure; secondly, measurements of esophageal pressures; and thirdly, tracheal isolation for all groups and tracheal banding for the TB groups. The midline incision over the trachea was sutured closed with Ethicon 3-0 silk ( Ethicon Inc., Somerville, NT) and the animals were then monitored until they were fully recoveredfromanaesthesia. The site of incision was protected by a spray dressing, OpSite (Smith & Nephew Inc. Quebec, Canada). Following the surgical protocol, the rats were administered a prophylactic i.m. injection of ampicillin (0.144 mL/Kg) to minimize risk of infection, and a subcutaneous injection of 10 mL of 0.9% normal saline to replenish fluid loss during surgery. On a daily basis, the animals were treated as follows: 1. givenfreshwater with tetracycline hydrochloride (8 g/L); 2. had blood samples taken for white blood cells (WBC) counts, circulating PMN counts, and arterial blood gas analysis; and 3. antisera was administered to the ND groups. On day 4, following anaesthesia, final Pes measurements were taken and the animals were euthanized. The diaphragm, sternocleidomastoid (SCM), extensor carpus radialis longlus (ECRL), gastrocnemius, and trachea were excised for further analysis. The left hemidiaphragm was weighed and then quick frozen by submersion into liquid nitrogen for biochemical analysis to quantitate the MPO activity and biopsies were taken from the right hemidiaphragm for  54  histology. The ECRL and gastrocnemius were removed to serve as non-respiratory control muscles and the SCM to serve as another inspiratory muscle. Biopsies were mounted in gum tragacanth (Sigma), and quick frozen in isopentane cooled in liquid nitrogen for histochemical and immunohistochemical analysis. All muscle was stored at -70°C until further analysis was performed.  Carotid Artery Cannulation  The carotid artery was exposed through a midline incision over the trachea. The carotid artery was clamped with a 6 mm straight arterial dietrich clamp to restrict bloodflow to the artery to allow cannulation. The proximal end of the carotid artery was tied with a silk thread to prevent blood flow from the superior region prior to the arterial incision. Once the carotid artery was cut, the chevroned end of the sterile carotid artery cannula (see Appendix A for carotid artery cannula preparation) was threaded into the artery to the insertion point marked on the cannula. The cannula was then tied in place with silk threads. The other end of the cannulae was subcutaneously threaded, exposed and stabilized at the back of the neck. The carotid cannula was madefroma polyvinyl tubing (I.D. 0.58 mm, O.D. 0.99 mm) coated with a 7% TDMac Heparin Complex (Polysciences Inc. Warrington, PA) which provided an anticoagulant surface. Carotid artery cannulation was performed on all of the rats to obtain arterial blood gas samples on each post-operative day in the awake rat. Antisera was administered and blood samples for white blood cell counts were also obtained via this cannula.  Esophageal Pressures and Tracheal Banding  The TB groups had their tracheas isolated and then banded. Esophageal pressure was  55  monitored by inserting a water-filled catheter into the lower one third of the esophagus (Reid et al., 1994; Reid et al., 1992). The water-filled catheter was attached to a differential pressure transducer (model 267BC; Hewlett Packard, Waltham, MA). The pressure transducer was connected to a calibrated recorder (see Appendix B for an example of an esophageal pressure (Pes) print out and calculations to determine Pes).  PeSjy  is the esophageal pressure during tidal  volume and was determined by measuring the height of one breathfrombaseline and then multiplied by the Gain factor of xlO. The Gain (MV/V) amplified the readingsfromthe differential pressure transducer. Maximal esophageal pressure (Pes,^) was obtained by occluding the trachea with forceps and then taking the maximal Pes which was defined as the highest value following a plateau of Pes during a minimum of three breaths. Pressure time index (PTI) was definedfrom(Pesxv/Pes,^) x (T/T ), where T is the inspiratory time and T is the total time tot  (  tot  taken for one respiratory cycle (Rochester, 1991). A tracheal band was made from a polyethylene tube (I.D.: 1.8mm; width: 2 mm) with a central slit. This band was placed around the isolated trachea and then tightened with two silk threads. The band was then gradually tightened in small increments until the post-banded Pes^, was approximately 10 cmH 0. Control rats also had their 2  trachea isolated, and Pes™ and Pes,^ measured. On day 4, following anaesthesia, all Pes measurements were taken, and the animals were euthanized.  Neutrophil Depletion Initially we investigated two different techniques to deplete neutrophils. First we attempted to produce our own antisera by injecting rabbits with rat serum, and secondly, by administering the drug cyclophosphamide. Injecting rat serum into rabbits resulted in antisera which lacked specificity to the rat neutrophil, therefore neutrophil depletion could not be achieved. The second  56  attempt to deplete neutrophils was to administer cyclophosphamide, a cytotoxic drug. Cyclophosphamide is a potent immunosuppressant which interferes with rapidly proliferating cells and can cause neutropenia (Norris, 1995). However, cyclophosphamide is non-selective and is therefore able to reduce the numbers of phagocytes, lymphocytes, B and T cells. Its cytotoxic effect on the rats left them in a very weakened state which also resulted in vomiting and weight loss. In humans, cyclophosphamide may be used for the treatment of certain autoimmune disorders such as systemic lupus erythematosus (Norris, 1995). Finally, neutrophil depletion was achieved by administering rabbit anti-rat polymorphonuclear leukocyte antisera (A1-A51140, Accurate Chemical & Scientific Corporation, Westbury, NY) daily. Different doses were tried in order to establish a regime for the administration of the antisera that would maintain neutrophil depletion. Initially 1.0 mL of antisera was administered and differential WBCs were counted to determine when circulating neutrophils begin to increase, usually seen by day 2 or 3. However, we found that different batches of the antisera had slightly different potencies. 1.0 mL of antisera from one batch might only be effective for one day of neutrophil depletion, therefore requiring a large (1.0 mL) subsequent dose the following day, whereas antisera from another batch migh be effective in maintaining low circulating PMN counts with one dose lasting for at least two days. Monitoring differential WBC counts was crucial for maintaining neutrophil depletion and determining the dose of antisera to be administered. Also, administering the antisera via the carotid cannula (post-surgery), rather than i.p.(pre-surgery), was more effective in depleting neutrophils, required smaller subsequent doses (0.5 mL) of antisera to maintain neutrophil depletion, and was more cost effective. The initial dose given to animals in the ND groups commenced one day prior to surgery via an i.p. injection. Following surgery, the antisera was delivered intra-arterially via the carotid cannula.  57  The initial dose was 1 mL of antisera which was usually sufficient to deplete neutrophils to less than or equal to 0.5 million/mL. Subsequent doses varied between 0.5 to 1.0 mL depending on the level of differential WBC counts in the morning blood sample takenfromeach animal. If the WBC counts were very low then a smaller dose of antisera was administered to maintain the neutrophil depletion level, and if the WBC count was closer to 0.5 million/mL then a larger dose of antisera was administered.  White Blood Cell Counts WBC counts were performed daily commencing one day prior to surgery. Prior to the carotid cannulization, blood samples were taken via the tail vein. The animal was stabilized in a plastic mould and its tail swabbed with heparin. The very tip of the rat's tail was nipped with a sharp scalpel and a blood sample volume of 2 uL was drawn up in a heparin rinsed pipette tip. The blood volume was then immediately mixed in 38 uL of counting solution for absolute WBC counts (Appendix C). The counting solution consisted of 49 mL of deionized water, 1 mL of glacial acetic acid and a pinch of methylene blue. Absolute WBCs were counted by placing 10 uL of the mixed solution onto a haemocytometer and viewed through a Nikon light microscope. WBCs were counted at a magnification of 400x in the four quadrants of the WBC counting chamber of the haemocytometer and corrected for the dilution factor which provided a value in millions/mL (Appendix C). Differential WBC counts were performed on blood smear slides that had been stained with Wright's stain. A drop of blood was placed on a glass slide and smeared with the edge of another slide to feather out the droplet. The blood cells were fixed in ethanol and allowed to air dry. The slide was then placed in an autostainer (Hema-Tek 2000, Miles Inc., Elkhart, IN) which was  58  loaded with a Hema-Tek Modified Wright's-Giemsa Stain Pak (Bayer Corp., Elkhart, IN), and coverslipped using permount. Differential WBC counts were performed while viewing slides at a magnification of 400x using a Nikon light microscope. A minimum of 100 cells were counted and white blood cells were categorized as polymorphonuclear (PMN) or non-PMN.  Arterial Blood Gases Daily arterial blood gases were taken to determine the acid-base status of the animal. An arterial blood sample volume of 0.3 mL drawn in a heparin coated syringe was taken from the carotid cannula in the awake animal. After the blood was drawn into the syringe, it was rotated back and forth between the hands to ensure mixing and then immediately injected into an Acidbase Laboratory (ABL) gas analyzer (Corning, Medfield, MA) to be analyzed. The blood volume removed was replenished with an equal volume of 0.9% saline, and the volume of the cannula was replaced with heparin 1000 Units/mL (Organon Teknika Inc, Toronto, Ontario) (Reid et al., 1994; Reid etal., 1992).  Measures of Resistive Loading Esophageal pressures and the internal cross-sectional area (CSA) of the trachea were used as indices of resistive loading. Pes^, Pes^/Pes^, and PTI values, taken immediately after the banding or sham procedure on day 1, and again on day 4, were used as estimates of loading. The trachea was excisedfromall rats and the internal CSA was measured and used as an index of resistive loading. The segments werefixedin 2.5% glutaraldehyde buffered with 0.1 M sodium cacodylate and stored in this fixative until further processing. Tracheal segments, trimmed to an appropriate size tofitthe embedding mould, were then washed for three 15 minute  59  washes in 0.1 M cacodylate buffer (pH 7.3). The tracheal segments were then dehydrated for ten minutes in 30%, 50%, 70%, and 90% ethanol followed by a total of three washes in 100% ethanol. Tracheas were infiltrated overnight on a rotator with catalyzed JB4 solution A (catalyzed solution A = 1:100 ratio of catalyst, an organic peroxide; to JB4 solution A, respectively, mixed well for 15 minutes). At this point, the polyvinyl tracheal band was carefully dissected free from the tracheal segment. The JB4 embedding solution (JB4 embedding solution B and catalyzed solution A, at a ratio of 1:25, respectively) was used tofillthe plastic embedding mould before placing the trachea upright in the middle of the mould. The trachea identification label was then placed alongside the mould wall and a mounting stub placed on top allowing some embedding solution to overflow. Polymerization occurred within one hour. Using a rotary microtome (DuPont Instruments Sorvall JB-4 Microtome), two micron thick serial crosssections were cutfromthe narrowest section of the embedded trachea and then stained with a 1% solution of Toluidine Blue O (Fisher Scientific, Fair Lawn, NJ). The internal cross-sectional area was measured using a light microscope (Nikon, MicrophotFX, Tokyo, Japan) connected to a multi-purpose image analysis system (Bioview Infrascan, Richmond, B.C.). The microscope was connected to a video camera (25.4 mm Vidicon, 60 Hz resolution, 81 series, DAGE-MTI Omo, Michigan City, IN) which projected the image of the trachea at a magnification of 25x (eyepiece lOx, optivar 1.25x, objective 2x) on a 20 inch colour monitor (1024 x 1024 pixels, 24 bit) (Sony Multiscan HG, GDill-1936m, Tokyo, Japan). Using the Bioview program, the vertical and horizontal axes of the video image was calibrated using a stage micrometer (Zeiss) delineated in microns. CSA of several serial tracheal cross-sections (which had been cutfromthe narrowest region of the rat trachea) were measured by outlining the internal circumference of the epithelial border using the mouse. The smallest value was taken as  60  the CSA for the trachea.  Muscle Histology & Immunohistochemistry Transverse sections of the muscle biopsies were cut to 10 and 8 um thickness using a cryostat-microtome (Reichart-Jung). Ten urn cross-sections were stained with haematoxylin and eosin stain and eight um cross-sections were processed for staining with DAB to identify PMNs, or processed using immunohistochemical techniques to identify PMNs or macrophages, respectively, then stained with haematoxylin. The sections were then quantified using either the point counting method or a counting frame to count cells. All slides were coded prior to analysis and the observer was blinded to the identity of the slide.  Diaminobenzidine Tetrahydrochloride (DAB) Method DAB method was used to determine the presence and location of neutrophils in the diaphragm muscle biopsies. An 8 p.m thick cross-sections of frozen diaphragm muscle was placed on a glass slide coated with Fro-Tissuer (Zymed Lab Inc), an agent used to adhere tissue sections onto the slide. On each cross-section 8 tolO drops of DAB substrate/chromogen was applied. The DAB substrate/chromogen kit (Dimension Labs Inc, Mississauga, On) was consisted of a 1:1 ratio of solution A, the chromogen solution which consisted of 3,3'-Diaminobenzidine, and solution B, the substrate solution consisting of hydrogen peroxide in buffer. The slides were incubated for 60 minutes in the dark for the development of the calorimetric reaction. The reaction was terminated by rinsing the slides with distilled water. The slides were counterstained in Meyer's haernotoxylin, dehydrated in ethanol, cleared in Hemo-De, and then coverslipped with a mounting medium. The presence of a brown precipitate, when examined under a light microscope, illustrated the presence  61  of the MPO enzyme. The DAB stain was used to determine the presence or absence of the enzyme MPO which is abundant in the neutrophil cytoplasm. The presence of a brown coloured precipitate is a positive result for the presence of MPO. DAB stained sections were not counted due to several reasons. Firstly, the precipitate was not localized and hence some cells were very light. Figure 16 shows a DAB stained muscle section takenfromthe diaphragm of a TB rat, DAB positive cells, indicated by an arrow, are lightly stained and difficult to distinguish. Secondly, it was difficult to determine if the precipitate represented a single cell or more than one cell. Lastly, the DAB stain underestimated the number of neutrophils in the muscle tissue by about 15 % compared to the H&E stained cross-sections.  Point Counting H&E Cross-Sections for Muscle Abnormality and Neutrophils in the Diaphragm The point counting method was employed to quantitate the area fraction (A^) of: 1. normal and abnormal muscle, and 2. PMNsfromH&E stained cross-sections. The setup consisted of a light microscope (Microphot-FX, Nikon, Tokyo, Japan) equipped with a camera lucida (Labophot, Nikon, Tokyo, Japan) and a computer with a stereology software package (The Gridder, WillRich Technologies, American Megatrends Inc.) used for point counting to determine A . A point grid from the computer monitor was projected via an angled side arm mirror (camera A  lucida) onto the light microscope field of view of the diaphragm cross-section. The points within the grid flashed in sequence until they had been designated a number by pressing one of the number keysfromthe key pad. Key pad numbers were pre-assigned using the Gridder program to different categories of morphologic features. Points that fell onto empty space or artifact were  62  keyed as zero and were automatically excludedfromthe total count. Point counting H&E sections for muscle morphology A 63 (7 x 9 rectangle) point grid was used for point counting muscle morphology viewed at a magnification of x400. Points located on the image of the muscle cross-section were assigned as follows (details of characteristics given in Table 1): 1. normal muscle fibre (Figure 12); 2. abnormal musclefibre(Figure 13); 3. inflammatory cell(s) in necrotic tissue - no cell outline (Figure 13); 4. inflammatory cell (s) in necrotic tissue - with cell outline; 5. connective tissue nucleus; 6. inflammatory cell nucleus (Figure 13); 7. effusion; 8. collagen and fat; 9. blood vessel and nerve; 0. no count. Once all the points had been assigned a number, the program stored the data onto a disk. Categories were then grouped as either normal = category 1, or abnormal and connective tissue = categories 2-9. The connective tissue category was grouped into the second category to include possible abnormalities present within the connective tissue, such as swelling of the intercellular space. The A was determined by taking the total number of points from each A  grouped category then dividing by the total number of points for all groups and then a percentage was calculated. TABLE 1; POINT COUNTING CATEGORIES AND CHARACTERISTICS Category  Characteristics  1. Normal muscle fibre  - polygonal shaped - consistentfibrediameter - closely packed muscle fibres - eachfibreand fasicle is surrounded by an endomysium and a thicker perimysium, respectively - uniform cytoplasmic staining - musclefibrenuclei located at the periphery of the fibre  63  2. Viable but abnormal fibre  - rounded and/or enlarged musclefibre(ie. >25% of the normal diameter) - small muscle fibres (ie <25% of the normal fibre diameter) - angulated muscle fibres - cytoplasmic staining appears homogeneous or non-homogeneous - nucleus located awayfromthe periphery  3. Inflammatory cell(s) in necrotic tissue- no cell outline  - muscle cell outline is absent - inflammatory cells including PMN and/or mononuclear cells - infiltration of inflammatory cells into the muscle fibre - non-homogeneous cytoplasm - remnants of cytoplasm  4. Inflammatory cell(s) in necrotic tissue- with cell outline  - cell outline present - inflammatory cells including PMN and/or mononuclear cells - infiltration of inflammatory cells into the muscle fibre - 3 or more rounded nuclei within or encroaching into the muscle cell - non-homogeneous staining of cytoplasm - cytoplasm of musclefibrelooks disorganized  5. Connective tissue nucleus  - spindle shaped - looks like a nucleus within a fibroblast - located in the interstitium  6. Inflammatory cell nucleus  - round, oval shaped, enlarged nucleus - looks like a nucleus within a monocyte or macrophage - located in the interstitium  7. Effusion  - proteinaceous substance stained pale pink - absent of fibroblasts - amorphous  8. Collagen and fat  - appears pale pink - present around muscle fibres, fasicles, and blood vessels - collagen apparent by the presence of fibroblasts and fibrillar appearance - fat cells appear as clear regions with defined outline approximately 25-200 microns in diameter  9. Blood vessel and nerve  - blood cells present within the lumen - vessels surrounded by collagenous material - nerves appears as bundles within an encapsulated cell  64  0. No count  - point falls on a clear space, or artifact - point has fallen out of thefieldof view  Inter and intra-observer reliability had been performed prior to commencing data collection. Inter-observer reliability was performed between myself and Dr. W. Darlene Reid. An interobserver reliability of 0.96 for normal muscle (category 1) and 0.84 for abnormal and connective tissue (CT) (categories 2-9) was found. Intra-observer reliability was 0.99 for normal and abnormal and CT categories on 12 occasions.  Point counting H&E sections for PMNs Point counting of H&E stained cross-sections to quantitate PMNs was performed as an alternative technique to quantitate PMNs in the diaphragm. Problems arose identifying PMNs using immunohistochemical techniques in the neutrophil depleted tissue and thus we could not accurately quantitate neutrophils using this technique. A point grid consisting of 196 points (14 x 14) was used for point counting PMNs viewed at a magnification of 1500x (eyepiece lOx; optivar 1.5x; objective lOOx), using oil immersion. Points located on the muscle cross-section were assigned as shown in Table 2.  TABLE 2: CATEGORIES AND CHARACTERISTICS FOR POINT COUNTING H&E SECTIONS FOR PMNs Category  Characteristics  1. Normal Muscle  - see Table 1, category 1  2. Other  - see Table 1, categories 2-9  3. PMNs  - cells with multilobed nuclei - located in interstitium or within the muscle fibre  65  4. Questionable  - cells that are difficult to distinguish clearly as PMNs - all mononuclear cells could be fibroblasts, macrophages, lymphocytes or muscle progenitor cells  Immunohistochemistry Identification  of Macrophages:  Macrophages were identified by the mouse monoclonal anti-rat  antibody, EDI (Cedarlane Laboratories Ltd, Ontario, Canada). The EDI antibody is directed against a macrophage subpopulation associated with inflammation and muscle atrophy (McLennan, 1996; St. Pierre & Tidball, 1994). The antibody is targeted against a single chain glycoprotein that is expressed mainly on a macrophage lysosomal membrane and with weak cell surface expression on macrophages and peripheral blood granulocytes (McLennan, 1996). Eight um thick sections, were placed on 3-aminopropyltriethoxysilane coated slides and air dried for 30 minutes. Sections werefixedin acetone for 10 minutes and then allowed to air dry for another 30 minutes. Prior to the slides being loaded onto the Dako Autostainer (Dako Corp, Mississauga, On), a paraffin marker pen (Dako) was used to encircle the sections to retain the solutions applied onto the sections. As soon as all of the solutions, except for the substrate, used for immunohistochemistry were prepared the Dako Autostainer was started. Solutions were positioned in the rack according to the map provided by the autostainer. The New Fuchsin Naphthol AS-B1 phosphate substrate was prepared immediately before it was needed as indicated at the appropriate time by the autostainer. The Dako Autostainer had been programmed according to the following detailed procedure (Cordell et al., 1984). The sections were rinsed in Tris buffered saline containing 0.25% Tween 20 (TBS), pH 7.6 then incubated with 100 uL of 5% normal goat serum, which was used as a protein block. Following the protein block, the primary antibody, EDI at a predetermined dilution of 1/250, was applied and allowed to incubate  66  for 180 mins at room temperature. The sections were then again rinsed in TBS, followed by another incubation with a 1/250 dilution in TBS of biotinylated goat anti-mouse immunoglobulin G (Vector Laboratories) and 2% rat serum for 90 minutes. Sections were then rinsed with TBS prior to the application of alkaline phosphatase linked to Vectastain Avidin-Biotin Complex (ABC-AP) (Vector Laboratories) for 30 minutes. Following a TBS rinse, the sections were incubated for 20 minutes with freshly made New Fuschin and Naphthol AS-B1 Phosphate, a substrate for the enzyme phosphatase. The calorimetric reaction was stopped by rinsing the sections with TBS followed by counter staining with Mayers' haematoxylin for 1 minute and then rinsed with water. Sections were then dehydrated in gradated ascending concentrations of ethanol, cleared in xylol to remove the paraffin mark around the section on the slide, and then cleared with Hemo-de (Fisher). Lastly, the slides were coverslipped in Entellan permanent mounting medium (BDH). As a negative control, the primary antibody was replaced by mouse IgG at the same concentration as the EDI antibody.  Identification of PMNs. Two antibodies were tested to label rat PMNs in muscle cross-sections. Both rabbit anti-rat polymorphonuclear leukocyte antisera - a polyclonal hyperimmune antisera (A1-A51140, Accurate Chemical & Scientific Corporation), and a rabbit anti-human MPO antibody (Dako Corp.) were used to target neutropils but with little success. Eight um thick sections were placed on 3-aminopropyltriethoxysilane coated slides and air dried for 30 minutes. Sections were fixed in either acetone or formalin for 10 minutes and then allowed to air dry for another 30 minutes. Prior to the slides being loaded onto the Dakoautostainer, a paraffin marker was used to encircle the sections to contain the solutions applied onto the sections. As soon as all of the solutions for immunohistochemistry were ready, with the exception of the substrate, the  \  67  autostainer was started. Various concentrations of the Accurate antisera and rabbit anti-human MPO antibody failed to yield an appropriate dilution because overstaining persisted even at the weakest dilution of one in one million in the diaphragm of the C+ND and TB+ND rats (Figure 17). Overstaining occurred in the C+ND and TB+ND tissue as a result of the secondary antibody recognizing and binding to the rabbit component of the Accurate rabbit anti-rat PMN antibody. The Accurate antibody had been successfully used to deplete circulating neutrophils in the peripheral blood in these rats. Residual antibody however, from prior injections, may have still been present in the tissues. Therefore, the rabbit anti-human MPO antibody, which cross reacts with rat MPO also resulted in overstaining because of the rabbit host in which the antibody was made. In an attempt to resolve the overstaining of the tissue different concentrations of the protein block, 5% and 10% normal goat serum, was used as well as trying tofixthe tissue with formalin instead of acetone to block the residual antibody in the diaphragm of the C+ND and TB+ND rats. The problem was not resolved and overstaining persisted in the ND tissue. Immunohistochemistry for PMNs was not continued since a commercial antibody was not available that had not used a rabbit host.  Quantification of Macrophages  EDI positive cells were quantified by using the Bioview image analysis system, as described above (see section titled Measures of Resistive Loading). A large unbiased countingframewas superimposed on the image of the muscle cross-section viewed at a magnification of x200 (eyepiece lOx, optivar lx, objective 20x) displayed on the computer monitor. A stage micrometer (Zeiss) was used to check the dimensions of the countingframeprior to quantification. The counting frame consisted of two sides for inclusion which appeared as dashed lines, and  68  two sides for exclusion which were solid lines. EDI positive cells were counted if they were within the frame or touched the inclusion sides, but were excluded if they touched the exclusion sides of the frame. The same criteria was applied while counting muscle fibres. The number of macrophages was expressed as either the number of EDI positive cells per number of muscle fibres or as the number of EDI positive cells per muscle cross-sectional area. The total sum of the cross-sectional area was calculated by multiplying the number of fields viewed by the area of the counting frame which was 0.26 mm . 2  Myeloperoxidase Assay 10 |ig of frozen diaphragm was placed in an iced test-tube containing 70 n L of iced hexadecyltrimethylammonium bromide (HTAB). Next, enough HTAB was added to reach a dilution of 10 pL per mg of muscle. The muscle sample was kept as cold as possible to minimize enzyme activation. The tube was kept on ice while the muscle was homogenized using an UltraTurrax homogenizer (IKA Laboratories-model TR-10) for two 15 sec intervals. The homogenized muscle was then centrifiiged (Hermle HM-22059V05 Rotor, Germany) at 2900 rpm for 12 minutes at a temperature of 0°C. Following centrifugation, the sample was twice frozen in liquid nitrogen and then allowed to thaw. After the second thaw, the sample was recentrifiiged at the same settings as mentioned. It has been reported by Smith et al. (1989) that thefreezingand thawing procedure yields a supernatant with maximal MPO enzyme and minimal haemoglobin contaminants. MPO activity was quantified using a spectrophotometer (Shimadzu UV-160, Japan) set at 480 nm wavelength (Belcastro et al., 1996). The spectrophotometer was maintained at 37°C by  69  employing a circulating water bath. The spectrophotometer was zeroed with the following solution mixture as background: 500uL of 0.03% hydrogen peroxide; lmL of 10 mM potassium phosphate buffer (pH 6.0); 50uL of the substrate 0.4 mM O-dianisidine dihydrochloride (ODMB); and 450uL of HTAB. MPO in the muscle sample was quantified by substituting the HTAB with the supernatant from the sample. The reaction was followed for a minimum of 150 seconds atfivesecond intervals. Maximal activity was calculated by curve-fitting the data onto a first order rate constant (GraFit version 3.0, Erithacus Software, Staines, UK)(Belcastro et al., 1996). MPO activity was expressed in Units/g of wet weight of tissue, where 1 unit of MPO activity was defined as 0.1 absorbance change at 480 run.  Statistical Analyses  Multivariate analysis of variance (ANOVA) was used to test for differences between the four animal groups for related parameters: 1. ABGs (Pa0 , PaC0 , pH, and HC0 "). Univariate 2  2  3  ANOVAs were used to test for differences between groups of animals for the following: 1. tracheal CSA; 2. diaphragm weights; 3. abnormal and normal muscle; 4. number of circulating PMNs; 5. number of macrophages in the diaphragm. A two way ANOVA was performed on circulating PMN counts to examine differences between groups and days. If the ANOVA indicated a significant difference between the groups then the Tukey HSD post hoc test was performed to determine the specific difference between groups. Differences were determined to be significant at a p-value less than 0.05. A two-way repeated measures ANOVA, with time as the repeated measure, was performed examining for differences between Pes , Max  PeSyy/PesMax  Pesyy  and PTI amongst the four animal groups. If the ANOVA indicated a significant  difference between groups then a Dunnett's test, which adjusts for non-normality of data and only  compares each experimental group to the control group, was performed. Data for the C+ND group on day 1 was excluded since this group had an n of 3 for esophageal data and did not meet the requirements for ANOVA.  71  F i g u r e 16: P h o t o m i c r o g r a p h of d i a p h r a g m m u s c l e c r o s s - s e c t i o n t a k e n f r o m a T B rat s t a i n e d with D A B a n d c o u n t e r - s t a i n e d with M a y e r ' s h a e m o t o x y l i n . D A B positive (arrows) neutrophils w e r e u n d e r - r e p r e s e n t e d by this stain a n d w e r e not c l e a r l y d i s t i n g u i s h a b l e , (x 4 0 0 )  72  Figure 17: Photomicrograph of over-stained cross-section of rat diaphragm with Accurate anti-rat neutrophil antibodies counter-stained with Mayers haemotoxylin solution. Over-staining of the interstitial space (arrows) surrounding the muscle fibre (arrow head) occurred due to the secondary antibody recognizing the primary antibody, which had initially been injected for neutrophil depletion, and then used for immunohistochemistry. Attempts to block the residual anti-rat neutrophil antibody failed and over-staining persisted. Scale bar = 25 urn.  73  CHAPTER 3 RESULTS Tracheal Cross-Sectional Area Tracheal CSA was used as an index of resistive loading and was significantly (p<0.001) reduced by an average of 65% for both TB groups (Figure 18) compared to the C and C+ND groups. The mean tracheal CSA for TB and TB+ND groups was 0.89 mm ± 0.34 and 1.11 mm 2  2  ± 0.21 (respectively), a reduction of 68.77% and 61.05% (respectively, as shown in Table 3 and Figure 18). There was no difference between the two C groups or between the two T B groups.  Esophageal Pressures Results for esophageal pressures are presented in Table 3. Data is missing because of complications that occurred during Pes measurements. Complications included respiratory arrest, and airway spasms which resulted from a difficult insertion of the esophageal catheter. Comparing the difference at each time interval for each group we found: Pes^ for the TB and TB+ND groups had significantly (p<0.05, p<0.01, respectively) increased postbanding on day 1 compared to prebanding on day 1 (Table 3). Comparing groups, postbanding Pes^ in the TB and TB+ND group was significantly (p<0.05) higher than the C group on day 1 (Table 3; Figure 19). PeSxy/PesM^  was also significantly higher (p<0.05) for the TB and TB+ND groups when  compared to the C group on day 1, although Pes  Max  alone did not show any significant difference  (Table 3). Differences were not found between all four groups for the following: P e s T y / P e s ^ on day 4; PTI on day 1 or day 4; Pes  Max  on day 1 or 4, or Pes^ on day 4. Significant differences  were not found between the TB group compared to the TB+ND group for prebanding, and postbanding Pes^ on day 1 and day 4.  74  Circulating Absolute Neutrophil Counts  Circulating absolute PMN counts, when compared to the C and TB groups, were significantly lower in the C+ND and TB+ND groups on days 1, 2, 3 and 4 (Table 4; Figure 20) (day 1 to day 3, p<0.001; day 4, p<0.05). No differences were shown between the C and TB groups or between the C+ND and TB+ND for each of the experiment days, or between the four groups on day 0, prior to the commencement of administration of the anti-rat PMN antisera. Comparing the differences at each time interval for each group showed two main results. Firstly, absolute PMN counts increased in the C and TB groups post-operatively on day 2 compared to their day 0 and day 1 values (p<0.001) (Table 4), an increase of 189% and 106%, respectively. Secondly, the absolute PMN counts decreased in C+ND and TB+ND groups (89% and 92%, respectively) after administration of the anti-rat PMN antiserafromday 0 to day 1 (p<0.001). A significant difference was not shown between the following: day 0 to day 1 and day 3 to day 4 for the C and TB groups; and between day 1 to day 4 for the C+ND and TB+ND groups. In summary, neutrophil numbers were significantly reduced in the neutrophil depleted groups (C+ND and TB+ND) compared to the C and TB groups.  Arterial Blood Gases  The results for the arterial blood gases are presented in Table 5 and Figure 21. PaC0 : TB and TB+ND groups were hypercapnic (p<0.001, p<0.05, respectively) with a 2  PaC0 of 57.21 mmHg ± 11.31 and 50.17 mmHg ± 9.28, on day 4 respectively compared to the 2  C and C+ND groups. There was no difference in PaC0 between the C and C+ND groups (mean 2  values were 40.69 mmHg ± 3.01 and 41.48 mmHg ± 1.74, respectively) or between the TB and TB+ND groups.  75  pH: Arterial blood was more acidic in the TB and TB+ND groups compared to C values. The pH in the TB group was significantly (p<0.001) lower by 22% compared to the C group (pH 7.46 ± 0.02) and 18% compared to the C+ND group (pH 7.44 ± 0.03). The pH was significantly (p<0.05) lower in the TB+ND group by 13% when compared to the C, and by 9% when compared to the C+ND group. The TB group had a lower pH, a difference of 11% than the TB+ND group (p<0.02). A significant difference was not found in the pH of arterial blood between the C and C+ND,groups. Pa0 : The Pa0 was significantly lower in the TB group by 15% compared to the C group 2  2  (p<0.05). The TB group tended (p<0.219) to be 10% lower in comparison to the C+ND group. In summary the TB group was hypoxemic. HCOf: HC0 " was higher in the TB group (31.57 + 3.92) when compared to the C+ND group 3  (27.79 ± 1.59) (p<0.05). No significant differences were shown between the C, C+ND and TB+ND groups. In summary, tracheal banding had the effect of increasing PaC0 and lowering Pa0 2  2  Hemi-Diaphragm and Body Weights Hemi-diaphragm weight (normalized as a percentage of the body weight on day 4) was higher by 18% in the TB+ND group compared to the C group (p<0.005) (Figure 22, Table 6) and tended to be higher compared to C+ND group (p=0.068). There were no significant differences for hemi-diaphragm weights between the C, C+ND and TB groups. Animal body weight loss was about 10 to 11% and was not significantly different between groups.  76  Area Fractions of Abnormal Diaphragm  The TB group had significantly (p<0.05) 33.5 % more abnormal muscle and connective tissue (16.46 % ± 1.30 S.E.) when compared to the C+ND group (12.34 % ± 0.70 S.E.), as presented in Table 7 and Figure 23, and 22% more when compared to the TB+ND group (12.83 % ± 0.74 S.E.; p<0.05). The TB group also had the smallest A of normal muscle (83.54 % ± 1.30 S.E.) A  compared to the C+ND group (87.66 % ± 0.70 S.E.; p<0.05) and compared to the TB+ND group (87.17 % ± 0.74 S.E.; p<0.05). An interesting abnormal featurefroma TB+ND rat, shown in Figure 24, shows an enlarged cross-section of a muscle fibre completely surrounding another muscle fibre. The fibre appears to be separated by connective tissue. This abnormal feaiture was located in the midst of the muscle cross-section, ie. in the same plane of section which appeared normal, therefore reducing the possibility of an artifact.  Area Fractions of Neutrophils  A of PMNs was performed on a total of 12 cross-sections (3 animals/group). This data was A  collected to further examine whether neutrophil depletion had occurred in the ND groups. Table 8 shows that the area fraction of PMNs in the diaphragm cross-section tended to be lower in the ND groups. The TB group almost had a four-fold increase in the PMN A compared to the C A  group.  Immunohistochemistry  EDI  The data for immunohistochemistry using an EDI antibody directed against inflammatory macrophages is presented in Table 9. Fields of view were counted over the whole cross-section  77  of the costal muscle biopsy which ranged between 31 and 63 according to the size and quality of the muscle cross-section. Two outliers were excluded from the C and C+ND groups' data (one from each group) because their ED1+ counts were extremely high due to overstaining. It is likely that overstaining may have occurred due to technical errors such as excess delivery of the antibody or by some contamination of the muscle section which increased the immunostaining. EDI positive cells per musclefiberwere significantly higher in the TB (0.0402 ± 0.0071 S.E. EDI/muscle fiber, p<0.01) and TB+ND groups (0.0354 ± 0.0054 S.E. EDI/musclefiber,p<0.05, Figure 25) compared to the C group (0.0164 ± 0.0024 S.E. EDI/muscle fiber). The TB group was also significantly (p<0.05) higher than the C+ND group ( 0.0215 ± 0.0036 S.E. EDI/muscle fiber). A significant difference was not present between the two control groups, C and C+ND. ED1+ cells per cross-sectional area were higher in the TB group (8.12 ± 1.41 S.E.; p<0.02) and tended to be higher in the TB+ND group (6.90 ± 1.08 S.E.; p=0.085) compared to the C group (Figure 25).  MPO  MPO activity results from the costal diaphragm are presented in Table 10 and Figure 26. MPO activity tended to be lower in C+ND (n=2) (1.75 units/g ± 0.38) and TB+ND (n=8)(1.45 units/g ± 1.30) groups than in the C (n=9) and TB (n=6) groups. The TB group tended to show the greatest value for MPO activity (4.58 units/g ± 1.86).  78  Figure 18: Bar graph of tracheal cross-sectional areas in control and banded groups. Means + SD are shown. * indicates significant difference from C and C+ND at p<0.00L  79  15  R  Day 1  Day 4  Figure 19: Bar graph showing esophageal pressures (Pes-rv). Means ± SD are shown. * indicates a significant differencefromC at p<0.05  80  10  r  9  E "5-  o  0)< * •—  8  h  7  -  6  -  5  -  4  "  J3 O  3  <  2  c C+ND TIB  TB+ND  1  -fi  0  Day 0  Day 1  Day 2  Day 3  Day 4  Figure 20: Scatter graph showing circulating absolute PMN counts on days 0 through to 4. Means ± SD are shown. * indicates a significant difference between C+ND and TB+ND compared to C and TB groups at p<0.001. ** indicates a significant difference between C+ND and TB+ND compared to C and TB groups at p<0.03. t indicates a significant difference (p< 0.05) compared to day 0, 1, 3 and 4 for the C and TB group.  81  Figure 21:  Bar Graphs for arterial blood gas (ABG) data. Means ± SD shown. Top left - PaC0 * indicates a significant differencefromC and C+ND at p<0.001 ** indicates a significant differencefromC and C+ND at p<0.05 2  Top right - pH * indicates a significant differencefromC and C+ND at p<0.001 ** indicates a significant differencefromC and C+ND at p<0.05 ["I indicates a significant difference between groups at p<0.02 Bottom left -  HCO3"  * indicates a significant differencefromC+ND at p<0.001 Bottom right - Pa0 ** indicates a significant differencefromC at p<0.05 2  Figure 22: Bar graph of normalized hemi-diaphragm weight. Means ± SD shown. * indicates significant differencefromC at p<0.005.  % of Total o  u c  •a  0>  cd  *-  <U  .S  o  s 1 52  _  >  I  •s  g  C  ed  u  <  O  13 C CO  O  Ui  3  O  Ui  m +l Ui  C  ca  CO  CO  O  6fl  c 15  « -  ©  J =  ©  SP v <U J = ~  13 C cd  §s  "c5  Ml  a. co  g p  ^ <i= — 0)  <  <  o E c £ cd  Ui  e o  o  PQ  2H  c O  E i_  Q . CO Q .  c  o £  t3 ,2  ° "°  J a  CO  - -a  a> CO  g  60  > "tn O co *2 M  o  J3  2 %  a. &o 60 — CO co 60 J=  u a. as CO  CO  CQ '-a  (S  *  8 3  F i g u r e 2 4 : P h o t o m i c r o g r a p h of a n a b n o r m a l feature in the m u s c l e fibre from the d i a p h r a g m t a k e n from a t r a c h e a l b a n d e d + n e u t r o p h i l d e p l e t e d rat, s t a i n e d with H & E . N o t e the r o u n d e d s h a p e d m u s c l e fibre c o m p l e t e l y s u r r o u n d e d by a n o t h e r m u s c l e fibre. S c a l e b a r = 1 0 u m .  86  Figure 25:  Bar graphs showing the number of EDI positive macrophages. Means ± SE are shown. Top Panel - Number of ED1+ cells per cross-sectional area (CSA) of diaphragm.  * indicates a significant differencefromC at p<0.02 Lower Panel - number of ED1+ cells per muscle fiber (xlO ) 2  ** indicates a significant differencefromC at p<0.01 + indicates a significant differencefromC at p<0.05 |~1 indicates a significant difference between TB and C+ND groups at p=0.05 Note: SE bars for C and C+ND groups are not visible because of small SE values.  87  88  Figure 26: MPO activity in the diaphragm. 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 w M  i—1 T f C N CO C N  o o © ©  CN  > O CA  T f  0.26 0.18  CN CO CN  o  0  o o o  © Tf  © o  0.41 0.89 0.14 0.32  a 5  0  TJ-  0.07  0  hH  o  1—1 co  0.32  * * PH  o\  0.61  1i  3  —1  ~  0.91  9  H  (N  0.08  >  3 S cu PH  co  0.17  PH  vo m  W> O CN CN  CN —'  H  > O  in1 in  s  8  5 I  3  S3  ro  o  O O r~ O 0 0 0 VO  00  t~~  in ©  ©  0  o  o  o o  o o  *ocor^vOcooorfTTO^ITION^OONOOONCNCO'—> O ^ r-.' O O O ~ -1  VO  3  •  M  0  ^. *l 00 CN CN CO CN  o  in  o  00 00  o  CN  CN  To 2  CO  o  o  in  in  CN  vd  00 CN ©* CN ^H • •  Tf  o o CO © ON <i CM  »H  ©  CO  m  a.  o  Q iZ CQ H  C N CO  (A  92  TABLE 4: Circulating Absolute PMN Counts GROUP Animal Absolute Circulating PMNs no. Day: 0 1 2 1 C 0.90 0.90 3.37 • 2 1.33 1.33 4.00 3 3.15 3.15 3.51 4 1.04 1.04 10.79 5 1.52 1.52 3.92 6 3.76 3.76 10.75 7 2.12 2.12 7.89 8 0.30 0.30 1.27 9 2.34 2.34 5.83 10 1.40 1.40 1.55 11 1.49 1.49 3.30 12 1.68 1.68 4.63 Mean 1.75 1.75 5.07 SD 0.96 0.96 3.18 1 C+ND 0.40 0.35 0.14 2 1.92 0.5 0.09 3 2.22 0.06 0.04 4 0.24 0.03 1.10 5 0.93 0.17 0 6 2.96 0.40 0.42 7 5.03 0.08 0.12 8 3.93 0.08 0.13 9 2.53 0.24 0.36 10 3.11 0.36 0.081 11 4.11 0.47 0.18 12 2.00 0.40 0.39 Mean 2.52 0.28 0.17 SD 1.39 0.16 0.15 A  1B  1B  (million/mL) 3 4 1.61 0.78 1.54 2.35 2.24 2.02 2.96 2.15 0.53 1.46 1.83 2.06 3.65 7.02 0.55 0.38 5.50 1.11 2.45 2.36 0.83 1.39 1.43 2.58 1.73 2.50 0.96 1.90 0.005 0.22 0.08 0.27 0.01 0 0 0.12 0.12 0.04 0.12 0.69 0.04 0.18 0.11 0 0.08 0.34 0 0.08 0.08 0.25 0.51 0.13 0.10 0.19 0.14 0.19 1B  2B  1 = Significant difference (p<0.001) compared to C and TB groups; 2= significant difference (p<0.03) compared to C and C+ND groups. A = significant difference (p<0.05) compared to day 0, 1, 3, and 4; B = significant difference (p<0.05) compared to day 0.  TABLE 4: Circulating Absolute PMN Counts (Continued) GROUP Animal Absolute Circulating PMNs (million/mL) no. Day: 0 1 2 3 4 TB 1 1.58 1.58 5.57 2.20 1.33 2 3 4 5 6 7 8 9  Mean SD TB+ND  Mean SD  1 2 3 4 5 6 7 8 9 10 11 12  1.44 1.69 3.01 3.34 1.47 3.20 3.07 3.59  1.44 1.69 3.01 3.34 1.47 3.20 3.07 3.59  4.15 5.62 4.10 2.11 5.35 9.91 5.31 4.05  0.51 1.23 1.01 1.37 4.17 3.21 1.56 3.34  0.70 1.11 2.00 1.06 2.77 0.69 1.24 2.00  2.49 0.91  2.49 0.91  5.13 2.11  1.97 1.26  1.53 0.73  2.46 3.65 1.41 2.62 2.88 3.14 4.90 2.50 4.25 2.41 1.64 2.11  0.11 0.13 0.05 0.15 0.53 0.32 0 0.11 1.08 0.12 0.19 0.08  0.08 0.25 0.02 0.04 0.26 0.05 0.24 0.10 0.40 0.05 0.18 0.02  0.24 0 0.04 0 0.11 0.09 0.08 0.03 0.12 0 0.07 0  0.25 0.15 0 0.62 0.11 0 0 0.08 0 0 0.02 0  2.83 1.02  0.24 0.30  0.14 0.12  0.07 0.07  0.10 0.18  ,B  A  1B  1B  2B  1 = Significant difference (p<0.001) compared to C and TB groups; 2= significant difference (p<0.03) compared to C and C+ND groups. A = significant difference (p<0.05) compared to day 0, 1,3, and 4; B = significant difference (p<0.05) compared to day 0.  T A B L E 5: Arterial Blood Gas Results Group Animal pH PaC0 (mmHg) C 1 7.48 34.44 2 7.48 36.60 3 7.48 41.80 4 7.46 38.40 5 7.43 41.86 8 7.43 41.22 9 7.43 43.87 10 7.47 41.54 11 7.46 41.01 12 7.45 43.87 13 7.44 43.02 Mean 7.46 40.69 SD 0.02 3.01 C+ND 1 7.47 39.70 2 7.46 41.50 3 7.43 40.00 4 7.38 44.60 5 7.46 38.57 6 7.45 42.50 7 7.44 42.92 8 7.41 42.50 9 7.45 40.16 10 7.41 40.48 11 7.46 43.13 12 7.43 41.65 Mean 7.44 ' 41.48 ' SD 0.03 1.74 2  2 3  2 5  Pa0 HC0 (mmHg) (mEq/1) 112.85 25.31 100.40 27.10 83.10 30.50 83.70 27.10 107.20 27.68 97.49 27.19 85.15 28.87 100.08 30.16 97.38 28.79 95.22 30.00 89.79 28.87 95.67 28.32 9.61 1.60 94.20 28.30 84.70 29.40 99.30 26.40 71.60 26.30 107.66 27.02 81.58 29.78 84.40 29.07 82.99 26.72 101.49 27.86 99.00 25.09 83.31 30.21 93.70 27.27 90.33 27.79 10.46 1.59 2  3  7  1 = significant difference (p<0.001) compared to C group; 2 = significant difference (p<0.001) for the TB group compared to C+ND group; 3 = significant difference (p<0.03) for the TB+ND group compared to C+ND and TB groups; 4 = significant difference (p<0.03) for TB+ND group compared to C group; 5 = significant difference (p<0.05) for TB+ND group compared to C+ND group; 6 = significant difference (p<0.02) for TB group compared to C group; 7 = significant difference (p<0.03) for TB group compared to C+ND group.  TABLE 5: Arterial Blood Gas Results (Continued) Group Animal pH PaC0 Pa0 HC0 fmmHgl fmmHgt (mEq/h TB 1 7.35 50.76 70.76 27.50 2 7.37 51.61 86.56 29.55 3 7.30 68.04 81.80 32.62 4 7.38 51.50 86.56 29.90 5 7.36 57.65 82.99 32.30 6 7.36 54.58 78.44 30.36 7 7.45 46.50 99.10 31.74 8 7.37 56.59 77.90 32.02 9 7.40 47.69 90.56 29.20 10 7.31 58.90 81.50 29.20 11 7.30 88.20 52.30 43.00 12 7.38 54.50 89.90 31.50 Mean 7.36 ' 57.21 81.53 31.57 SD 0.04 11.31 11.70 3.92 TB+ND 1 7.35 53.30 60.10 29.30 2 7.44 41.60 102.00 27.60 3 7.40 53.73 89.05 32.90 4 7.43 50.30 99.00 33.20 5 7.39 46.80 95.10 28.20 6 7.40 49.50 82.00 30.00 7 7.43 41.40 96.20 26.90 8 7.39 48.60 92.70 29.10 9 7.42 41.80 92.90 26.50 10 7.43 44.80 93.60 29.20 11 7.38 54.70 78.30 31.80 12 7.36 75.50 76.30 41.90 Mean 7.40 ' 50.17 ' 88.10 30.55 SD 0.03 9.28 11.90 4.19 2  1 2  1,2  1 3  4 5  2  6  3  7  1 = significant difference (p<0.001) compared to C group; 2 = significant difference (p<0.001) for the TB group compared to C+ND group; 3 = significant difference (p<0.03) for the TB+ND group compared to C+ND and TB groups; 4 = significant difference (p<0.03) for TB+ND group compared to C group; 5 = significant difference (p<0.05) for TB+ND group compared to C+ND group; 6 = significant difference (p<0.02) for TB group compared to C group; 7 = significant difference (p<0.03) for TB group compared to C+ND group.  97  TABLE 6: Body and Hemi-Diaphragm Weights Group Animal No. C  Mean SD C+ND  Mean SD  1 2 3 4 5 6 7 8 9 10 11 12  1 2 3 4 5 6 7 8 9 10 11 12  Body Weight (g) Day 0 Day 4 403 375 402 352 515 426 498 385 503 460 489 446 502 402 404 388 404 380 381 315 381 363 384 354 439 387 56 41 463 443 449 442 464 437 408 372 386 338 365 324 422 397 413 380 387 356 416 376 392 348 395 361 413 381 32 41  Hemi-Dia (g) Day 4 0.540 0.397 0.460 0.575 0.468 0.550 0.496 0.466 0.428 0.436 0.494 0.471 0.482 0.052 0.601 0.512 0.563 0.478  Hemi-Dia BW (%) 0.144 0.113 0.108 0.149 0.102 0.123 0.123 0.120 0.113 0.138 0.136 0.133 0.125 0.015 0.136 0.116 0.129 0.128  0.469 0.558 0.444 0.494 0.535 0.455 0.473 0.507 0.051  0.145 0.141 0.117 0.139 0.142 0.131 0.131 0.132 0.010  1  2  Weight Loss (g) 28 50 89 113 43 43 100 16 24 66 18 30 52 33 20 7 27 36 48 41 25 33 31 40 44 34 32 11  % Weight Loss 7 12 17 23 9 9 20 4 6 17 5 8 11 6 4 2 6 9 12 11 6 8 8 10 11 9 8 3  ABBREVIATIONS: Hemi-Dia = hemi-diaphragm, BW = body weight on day 4. 1 = significant difference (p<0.005) compared to the TB+ND group; 2 = trend (p= 0.068) for TB+ND group compared to the C+ND group.  TABLE 6: Body and Hemi-Diaphragm Weight (Cont'd) Group Animal No. TB  Mean SD TB+ND  Mean SD  1 2 3 4 5 6 7 8 9 10 11 12  1 2 3 4 5 6 7 8 9 10 11 12  Body Weight Day 0 530 498 397 422 405 443 430 457 433 382 438 493 444 44 397 410 401 420 502 495 427 407 418 432 399 433 428 35  (  Day 4 485 431 333 374 384 412 358 425 399 315 346 439 392 50 365 331 336 390 436 439 384 362 374 390 385 352 379 34  Hemi-Dia (g) Day 4 0.545 0.566 0.422 0.470 0.520 0.585 0.561 0.610 0.551 0.464 0.470 0.630 0.533 0.064 0.424 0.595 0.417 0.520 0.740 0.679 0.565 0.451 0.615 0.606 0.576 0.541 0.561 0.098  Hemi-Dia BW (%) 0.112 0.131 0.127 0.126 0.135 0.142 0.157 0.143 0.138 0.147 0.136 0.144 0.137 0.012 0.116 0.180 0.124 0.133 0.170 0.155 0.147 0.124 0.165 0.155 0.150 0.154 0.148 0.020 1,2  Weight Loss (g) 45 67 64 48 21 31 72 32 34 67 92 54 52 21 32 79 65 30 66 56 43 45 44 42 14 81 50 20  % Weight Loss 8 13 16 11 5 7 17 7 8 18 21 11 12 5 8 19 16 7 13 11 10 11 11 10 4 19 12 5  ABBREVIATIONS: Hemi-Dia = hemi-diaphragm, B W = body weight on day 4. 1 = significant difference (p<0.005) compared to the C group; 2 = trend (p= 0.068) for TB+ND group compared to the C+ND group.  rs  U  86.61 1.04 2.05 0.38  CO  0.39 0.19 0.86 0.27 0.70 0.07  0.49 0.07  0.05 0.02  7.98 0.60  0.88 0.11  Effusion Collagen Nerve + bv 0.18 6.56 0.40 0.10 9.36 1.11 0.00 5.83 0.60 0.03 7.70 0.70 0.00 6.84 0.53 0.04 5.23 0.94 0.19 11.35 1.70 0.00 1.03 6.65 0.00 1.09 7.60 0.00 0.61 10.99 0.00 1.25 7.19 0.08 0.57 10.52  =J  84.28 84.27 91.67 90.16 86.11 90.13 79.00 89.87 83.35 85.81 87.97 86.64  01 es u JS Q 08  Rd-nucI 0.90 0.52 0.44 0.19 0.28 0.31 0.86 0.45 0.27 0.72 0.42 0.49  00  Infl -O/L Infl +0/L Sp-nucl 2.33 0.81 0.49 0.43 0.72 0.79 0.20 0.16 0.44 0.03 0.03 0.44 0.25 2.01 0.77 0.12 0.59 0.75 0.78 2.64 0.56 0.06 0.19 0.78 0.18 2.34 1.12 0.22 0.28 0.61 0.00 0.47 0.56 0.04 0.08 1.10  os  AbN 4.04 2.72 0.67 0.72 3.22 1.89 2.92 0.97 4.05 0.75 2.14 0.49  13.39 1.04  9.87 21.00 10.13 16.65 14.19 12.03 13.36  AbN + CT 15.72 15.73 8.33 9.84  A  Group Animal  OS  2w i  w  ii  60  eej  ed  c  00  E u ©  B  IS E i. o 93  Z «*.  o  <  W  CO  2 s  cu _> '-3  B O  s  U •a  B 93  CU  CO  93  CN  <  TJ S  i—I  0.12 0.06  0.69 0.12  41  2.00 0.22  JS  0.44 1.48 0.64 0.13 1.28 0.70 0.94 0.15 0.45 0.75 0.31 0.97  9i.3 JS a cs  0.00 0.74 0.00 0.00 0.00 0.07 0.06 0.00 0.14 0.06 0.31 0.07  e  0.69 2.35 2.04 1.48 2.78 1.63 3.22 1.36 0.95 2.45 2.70 2.34  o  0.44 0.43 0.52 0.47 0.36 0.73 0.42 0.44 0.68 0.36 0.64 0.55  0.50 0.04  0.72 0.43 1.05 0.47 0.53 0.97 0.50 0.65 1.13 0.89 0.61 0.73  0.72 0.07  0.06 0.02  0.06 0.25 0.00 0.13 0.00 0.00 0.03 0.00 0.14 0.06 0.06 0.00  0.89 0.43 0.70 1.62 0.61 1.07 1.06 0.95 0.72 0.95 0.31 0.68  0.83 0.10  6.81 9.95 4.08 11.73 6.70 5.41 7.89 5.86 6.39 7.84 8.80 7.46  7.41 0.60  Rd-nucl Effusion Collagen Nerve + bv  a  Z  89.94 83.93 90.97 83.96 87.74 89.42 85.89 90.59 89.40 86.66 86.27 87.19 87.66 0.70 1  "3 00  Infl-O/L Infl+O/L Sp-nucl  "SI  AbN  o\  Group Animal C+ND  1  AbN + CT 10.06 16.07 9.03 16.04 12.26 10.58 14.11 9.41 10.60 13.34 13.73 12.81 12.34 0.70  CN  100  3 CN oo  •K « »  00  O  a>  2 w  B  60  B fi O  03 eu  "3  u  o B  es *«  £  CM  O  <  <  w  PQ S IT)  eu  CU  eu  Tf  u  •a  B  f)  tn S  £  mO  «N  i-H  S5 0.17 0.20 0.45 0.04 0.06 0.00 0.06 2.09 1.20 0.09  0.41 0.18  3.17 2.62 6.22 2.97 3.00 0.93 7.02 1.83 1.85  3.27 0.50  03  2.98  mm O  1.31 0.35  0.57  0.56  3.68  0.55  2.69  0.19  3.24  1.14  0.72  1.39  so  0.83  eu  0.45  o  3.07  es  0.17  5 t-  0.14  01  3.61  E  86.13 81.75 81.71 85.57 88.56 80.21 89.19 85.12 88.60 73.86 81.68 80.05  o  0.25 0.64 0.78 0.44 0.47 0.36 0.39 0.74 0.55 0.60 0.42 0.31  0.50 0.05  0.77 1.01 0.72 0.31 0.80 0.44 0.80 0.44 0.86 0.56 0.68  0.67 0.06  eu  0.61  0.50 0.55  10.82 9.69 8.58 5.76 8.48 6.25  7.10 8.73 10.70 11.79 15.11  9.27 0.75  0.00 0.09 0.00 0.11 0.07 0.00 0.00 0.03 0.20 0.08 0.00  0.05 0.02  0.98 0.18  1.34  1.89  0.99  0.12  0.58  0.58  0.60  2.17  1.66  8.28  0.00  0.81  Rd-nucl Effusion Collagen Nerve +bv  s  Infl-O/L Infl+O/L Sp-nucl  00  AbN  e>  Group Animal TB  AbN + CT 13.87 18.25 18.29 14.43 11.44 19.79 10.81 14.88 11.40 26.14 18.32 19.95  OS  101  x> -  oo  B  CS  mj mmf  M 00  -a 00  B  %r S  00  — o  9  s  eu > u eu  E o s .o  <  es "es  B  E o  CM  z,  •• t--  O  «5  W  03 •*•»  C  •o  es cu  B  "3 m 3  "3  •o  Z 0.05  0.30 0.23  1.07  0.65 0.06  0.74 0.05  «s 0.18  rt  2.74  1/3  85.76  c o  ?  W  86.00  cu  82.80  es  86.65  5  88.07  es  85.55  u JS  89.77  01  87.54  s 6.84 0.48  0.07 0.03  0.07  0.55  Collagen Nerve +bv 8.83 0.58 7.83 0.66 5.99 0.49 3.48 0.28 5.07 0.45 7.17 0.31 7.64 0.77 5.94 0.15 8.22 0.72 7.98 0.87 8.53 0.82 5.40 0.46  e  92.71  o  Effusion 0.09 0.00 0.00 0.00 0.12 0.08 0.02 0.35 0.03 0.06 0.00 0.05  • mm  88.99  H 0 0  87.29  a  Rd-nucl 0.64 0.89 0.70 0.50 0.45 0.31 0.51 0.79 0.53 0.84 0.58 1.01  S  84.88  cu  Infl-O/L Infl+O/L Sp-nucl 0.46 0.35 0.72 0.28 1.22 0.61 0.00 1.03 0.81 0.06 0.78 0.50 0.20 1.10 0.78 0.14 0.03 0.61 0.14 1.19 0.77 0.00 0.50 0.89 0.00 0.45 0.59 0.46 2.26 0.64 0 07 1.16 0.88 0.32 2.84 1.05  CU  AbN 3.45 1.22 2.00 1.68 4.29 1.59 3.40 3.32 2.81 4.09 1.97 3.11  •o s  Group Animal TB+ND  ON  14.24  14.00  17.20  13.35  11.93  14.45  10.23  12.46  7.29  11.01  12.71  15.12  AbN + CT  102  ro rt  °°. ^. rs ©  CU  ~3 CN  03  co  00  So °  B es  60  TABLE 8: Area Fractions of PMNs Group C  Animal  Normal  Other  PMN  Questionable  1  89.86 90.03 88.50  9.90 9.27 11.26  0.07 0.19 0.05  0.17 0.51 0.19  89.47 0.48  10.14 0.59  0.10 0.04  0.29 0.11  2 3  Mean SE C+ND  Mean SE TB  Mean SE TB+ND  Mean SE  1 2 3  1 2 3  1 2 3  89.91 89.47 91.34  9.86 10.12 8.50  0.07 0.03 0.02  0.15 0,37 0.14  90.24 0.57  9.50 0.50  0.04 0.02  0.22 0.08  68.95 88.89 87.36  29.40 9.59 11.39  0.41 0.48 0.41  1.24 .1.04 0.83  81.73 6.41  16.80 6.33  0.43 0.02  1.04 0.12  86.03 89.63 89.12  13.54 10.12 10.49  0.03 0.03 0.02  0.39 0.22 0.37  88.26 1.13  11.38 1.09  0.03 0.00  0.33 0.05  eS J3 D O OJ  u es  u.  + Q  ca  Mu Fibre 0.0293 0.0092 0.0274 0.0063 0.0222 0.0117 0.0080 0.0202 0.0212 0.0104 0.0150 1  S  U E ^ E S  -eS  +  U 3.46 0.54  M eu N"*  4.78 0.18  3.91 4.49 5.59 4.30 5.81 4.46 4.96 4.33 5.41 4.75 4.53  rj  7.49 2.06 4.91 1.46 3.83 2.63 1.61 4.66 3.92 2.18 3.32  IZ)  2323 3036 2150 2235 2068 2816 2259 2772 2123 2415 2596  # of fields Mu Fibres  etf  Animal  a  Group  104  ^  3  w  n T  VO O O °. ©  Tt  «N  eS  «N  T  o  a oooooNTj-vofnoovommoN  UJ  VO  Ov  00  B  M  M m  CN  3  <2 o  es u JS  es  Q  3  O o s_  w es  Q  W ca  es  CU  1.66  0.0171 0.0173 0.0195 0.0240 0.0211 0.0139 0.0162 0.0215  52.64 9.84  2344 3121 2313 2706 2702 2295 2223 2416.09 110.36  0.0036  1.01  4.87  2.93  3.42  4.54  5.94  4.32  3.89  3.57  6.28  3.48  14.07  0.17  4.59  5.52  4.07  4.64  4.05  4.50  4.45  4.78  5.43  4.98  3.61  4.40  rC^  2  0.0340  1821  w  0.0173  +  2250  -©E  0.0508  cu  2738  s s  1.10  < ^  0.0048  u "  2064  CU  41.09  •a Mu Fibre  u  C+ND  E  # of fields Mu Fibres  +3 B O  Animal  s G  Group  105  <J3 "o  w  W  B  EJ -  00  CO _ l  00  Animal 3  u ^ a  +  o u s s  w  -eS  2 2 ^ 2 : « "  4.93 0.15  4.85 4.91 4.98 4.82 5.32 5.72 4.17 5.09 4.47  S  9.14 11.63 14.69 7.30 9.76 7.71 9.40 2.07 1.38  *  2484 126  md) Mu Fibre 0.0443 0.0571 0.0732 0.0352 0.0519 0.0441 0.0392 0.0105 0.0061 i-\  U  2751 2609 2089 2214 1907 2379 3012 2465 2930  # of fields Mu Fibres in  TB  Group  106  < *©  00  © © © •t °. © ©  VO VO OS i-H  4>  O O O O N O O O — ' t - - r ~ a \  m T j - f O T j - c n m T j - n - T } -  s  03  1  M  r-l  Animal  a 6.90 1.08  U S  10.89 3.02 1.93 3.89 3.37 8.16 6.48 7.49 10.05 4.87 14.60 8.03  Vi  Mu Fibre 0.0620 0.0130 0.0083 0.0237 0.0214 0.0365 0.0384 0.0380 0.0488 0.0226 0.0704 0.0419  rj  «s  w  -eE 5.14 0.20  5.69 4.31 4.31 6.08 6.34 4.47 5.92 5.08 4.86 4.64 4.82 5.22  M  2534 2230 1921 2241 2107 2331 2346 2523 2089 2301 2713 3223  # of fields Mu Fibres Vi  TB+ND  Group  107  -H  "3* £ ©  +  oo *n  Q W 00  £  cu  •5 S £  °  o -a  i-l  © 00 ON tn es  4>  s  ^?  M  «5 CN  TABLE 10; MPO Activity in the Diaphragm Group C  Animal No MPO (U/g) 1 2 3 4 5 6 8  1.15 2.05  3.22 2.10 1  1 3 4 5 6  7.25 5.01 3.56 6.03 3.35 2.28  4.58 1.86 1  2 3 4 5 6  7 8  Mean SD  2.30 1.21  1.75 0.38 2  Mean SD TB+ND  1.46 2.43 6.54 3.48  1.27  2 Mean SD TB  4.29  7 9 Mean SD C+ND  6.34  0.05 0.41 0.74 2.86 3.65 2.10 1.39 0.40  1.45 1.30  109  CHAPTER 4 DISCUSSION We found that neutrophil depletion decreased muscle injury and neutrophil-mediated damage caused an influx of macrophages into the diaphragm when loading was induced by tracheal banding. These observations are consistent with the hypothesis that neutrophils may mediate and even amplify muscle injury under conditions where the muscle is overloaded. Diaphragm loading, induced by tracheal banding, resulted in diaphragm muscle injury and an increased presence of inflammatory macrophages. Neutrophil depletion, however, decreased muscle injury and showed that neutrophils are involved in the process of attracting macrophages in to the diaphragm. The decreased tracheal cross-sectional area (by an average of 65%), increased arterial PaC0 (>45 2  mmHg), and increased esophageal pressure during tidal ventilation after banding in the TB and TB+ND groups indicates that the tracheal banding succeeded in loading the diaphragm. The decrease in the numbers of absolute circulating PMN to <0.5 million/mL in the C+ND and TB+ND groups demonstrates that we were able to significantly reduce neutrophil participation in the response. This study is unique in showing reduced diaphragm injury using a neutrophil depletion intervention, where diaphragm loading was induced by tracheal banding. Circulating neutrophils have been shown to become activated (Pyne, 1994) and to increase in number following exercise (Camus et al., 1992; Smith & Telford, 1990) due to increased cardiac output (Foster et al, 1986), muscle injury, or increased cytokines (Evans & Cannon, 1991). It is not known how much cardiac output increases during tracheal banding but cytokine release occurs during increased respiratory muscle loading in humans (Vassilakopoulos et al., 1999). Cytokine release may trigger a sequence of events; activated neutrophils and monocytes increase cell surface expression of adhesion molecules, such as CD1 lb/CD18. A variety of adhesion  110  molecules mediate leukocyte adherence to endothelial surfaces, transmigration, and chemotaxis towards interstitial inflammatory stimuli or tissue damage. Neutrophils may release a variety of cytotoxic chemicals that may affect and disrupt neighbouring cells and extracellular matrix. A chronic stimulus, such as tracheal banding, may further amplify their cytotoxic response. It has been described that neutrophils can be activated by cytokines such as interferon gamma (IFNY) d tumour necrosis factor (TNF), resulting in the a n  production of more superoxide anions (Roitt et al., 1989). If treated with both cytokines the response has been shown to be even greater than when treated by only one cytokine. Neutrophils pretreated with granulocyte monocyte-colony stimulating factor (GM-CSF) were able to kill parasites more quickly than without pretreatment. Similarly, the cytotoxic action of neutrophil may be accentuated by the mediator release due to ongoing muscle injury in the tracheal banded model. Cytokine release induced by diaphragm injury may promote activation and cell maturation of myeloid cells. Myeloid cells (monocytes and granulocytes) originatefroma pluripotent haemopoietic stem cell which can differentiate into one of five pathways (Roitt et al., 1989), depending on the cytokines that stimulate the stem cell. If a stem cell is stimulated by a granulocyte monocyte colony stimulating factor (GM-CSF) it becomes a more specialized stem cell called a colony forming unit - granulocyte, monocyte (CFU-GM) (Roitt et al., 1989; Metcalfe, 1986; Wong et al., 1985). Further stimulation by cytokines such as interleukin 3 (IL-3) and monocyte-CSF (M-CSF) promotes specialization of CFU-GM to monocytes (Roitt et al., 1989; Wong et al, 1987), or IL-3 and G-CSF promotes specialization to neutrophils (Nicola, 1989; Roitt et al., 1989; Nagata et al., 1986). In the absence of PMNs we saw less disruption of the muscle cells. Reduced tissue injury has  Ill  been found when the adherence of migrating neutrophils was blocked by monoclonal antibodies directed against adhesion molecules (Folkesson et al., 1997). The expression of CD1 lb has been shown to increase when neutrophils are activated (Pizza et al.,1996). Monoclonal antibodies directed against CDllb (Pizza et al., 1996) and CD18 both reduce the infiltration of neutrophils (Akimoto et al., 1996) and hence neutrophil-mediated reperfusion injury was reduced in the heart (Byrne et al., 1993) and in skeletal muscle. The purpose of this study was not to address the molecular mechanism of neutrophil migration in response to muscle injury but rather to demonstrate whether or not neutrophils play a role in muscle damage by investigating the effects of neutrophil depletion on muscle injury. In studies investigating the effects of neutropenia, less injury was observed following the injury inducing insult. Eppinger et al., (1995) demonstrated the effects of neutrophil depletion on lung reperfusion injury using a rat lung model with an occlusion of the left pulmonary artery for 90 minutes. They described a bimodal pattern of injury of neutrophil-independent and neutrophilmediated following reperfusion of the lung. Neutrophil depletion was achieved by the administeration of an anti-rat neutrophil serum or preimmune serum. Thirty minutes of reperfusion, described as neutrophil-independent, did not reduce lung injury, however, injury was significantly reduced after 4 hours of reperfusion. Studies investigating neutrophil depletion effects on tissue injury by myocardial ischaemia (Entman et al., 1991; Lucchessi et al., 1989), complement activated lung injury (Till et al., 1982), and cerebral ischaemia (Yenari et al., 1998) have all demonstrated reduced tissue injury. This study is novel in that we demonstrated for the first time that neutrophil depletion may reduce diaphragm injury produced by loading, which was induced by tracheal banding. This study found areas of normality and abnormality within the diaphragm of the C group  112  (Table 7). A significant difference was not detected (p=0.133) when the area fraction of AbN+CT of the TB group was compared to the C group. This result indicates two points. Firstly, the impact of the surgical procedure on the control animals. The surgical procedure involved isolating the trachea and measuring esophageal pressures under a general anaesthetic. Measurement of esophageal pressure and post operative atelectasis may have caused some loading of the diaphragm and diaphragm injury in the control group. Pes ^ measured to Max  determine the maximal effort of inspiration was obtained by tracheal occlusion, requiring a maximal effort of the diaphragm. It has been reported that unaccustomed activity, in this case, the measurement of Pes  MaXi  could possible lead to muscle injury. Alternately, the abnormal muscle in  the control diaphragm may represent changes that occur normally when a muscle is continually going through change. Less area fraction of AbN+CT was found in the C+ND group compared to the TB group (p<0.05). Our study showed similar diaphragm injury in the TB group compared to Reid et al. (1994) in terms of the area fractions of normal (83.54; 80.8 respectively) versus abnormal muscle and connective tissue (16.46; 18.2, respectively). Both studies used the tracheal banding model to load the diaphragm. Abnormal muscle categories were grouped with connective tissue categories (Table 1, categories 2-9) to incorporate the changes in the interstitium such as increased inflammatory cells and intercellular swelling. Road & Jiang (1998) reported widening of the interstitium and increased numbers of inflammatory cells in the rabbit diaphragm following resistive loading. In addition to similar quantitative changes, we found similar qualitative changes. Abnormal morphology present in the diaphragm in previous studies included features such as necrotic fibers, disruption of the normal cytoskeleton and staining, increased infiltration of inflammatory cells  113  (Lieber, 1992), angulated fibers, abnormal shapes and sizes of muscle fibers, and internalized nuclei (Reid & MacGowan, 1998). The TB group had increased (p<0.05) abnormal muscle and connective tissue compared to the C+ND and TB+ND groups. Interestingly, the TB+ND group had an A of abnormal muscle and A  connective tissue to be similar to the C values. This suggests to us that the difference in abnormality in the TB group compared to the TB+ND group after 3 days of tracheal banding is due to the neutrophil depletion intervention and therefore to the effect of the neutrophils on the costal muscle of the diaphragm. Less normal muscle features (p<0.05), therefore more abnormal, (described in Table 1 and in Chapter 1) were present in the TB group than for the C+ND and the TB+ND groups. The TB+ND group had A of normal muscle similar to C values. These results A  support the hypothesis that neutrophils do mediate and amplify muscle injury under a continuous load for at least a period of three days. Previous studies using increased resistive loading in different species have also found similar abnormal muscle features and connective tissue in the costal regions of the diaphragm (Jiang et al, 1998; Reid et al, 1994). Our study found the sum area fractions of abnormal muscle and connective tissue to be 16.46% ± 1.3 in the TB rats, while Jiang et. al. (1998) reported 8.0% ± 0.6 for abnormal muscle and 7.3% ± 0.6 for connective tissue, a sum area fraction of 15.3%. In another study by Jiang et. al. (1998a) they reported 8.7 ± 2.6 and 7.9 ± 0.5, respectively, a sum of 16.6%. Both studies were performed on the rabbit diaphragm after high inspiratory resistive loading. Inspiratory resistive loading was applied by an endotracheal tube connected to a 2 way valve with an adjustable needle connected to the inspiratory port (Jiang et al, 1998, 1998a). Reid et al (1994) used the tracheal banding model on hamsters for a period of 6 days and reported abnormal muscle and connective tissue in the TB group to be 13.3% ± 0.3 and 4.9% ± 0.7, a sum  114  area fraction of 18.2%. Overall, resultsfromthis study are comparable to previous studies, however some variations may be due to different loading regimens and different species:; hence different species may have somewhat different load tolerances due to fibre type characteristics or variable respiratory rates to produce muscle injury. ND was achieved by administering anti-rat PMN antisera. Figure 4 provides a summary of the results for the circulating absolute PMN counts on day 0 to day 4, showing ND to have been maintained (PMN<0.5 million/mL) during the experiment in the ND groups (p<0.001). The TB and C groups both had significantly (p<0.05) increased circulating PMN counts on day 2 of the experiment. This response is likely due to the effects of the surgical intervention. Circulating PMN counts returned to baseline values on day 3 for both the TB and C groups. Examination of the gastrocnemius muscle, a non-respiratory control muscle used in this study, lacked any evidence of muscle injury. However, the sternocleidomastoid, an inspiratory muscle did show evidence of muscle injury from the TB group. Although an extensive examination was not performed, muscle abnormality was not apparent in the SCM in the two animals from the C group. Muscle injury was not quantified for the SCM, however, due to the presence of injury in the SCM of the TB group warrants further investigation of other inspiratory muscles. Consistent with our hypothesis, there was a trend for less (15%) EDI positive macrophages in the costal diaphragm (per cross-sectional area) of the TB+ND group compared to the TB group. The TB and the TB+ND groups had more ED1+ macrophages per fibre compared to the C group (p<0.01, p<0.05, respectively). These macrophages most likely camefromcirculating monocytes responding to chemotactic agents and migrating to the sites of injury, since the EDI antibody specifically targets inflammatory macrophages (St. Pierre & Tidball, 1994; McLennan, 1996, 1993).  115  We found both TB and TB+ND groups to have increased ED1+ macrophages per crosssectional area compared to the C group by 135% (p<0.02) and 99% (p=0.085), respectively. These observations indicate an apparent trend of a dampened effect in the TB+ND group compared to the TB group. Neutrophil depletion may have blocked potential mechanisms which attract macrophages ie. specific cytokines released by neutrophils, such as IL-1 which attract macrophages, or aspects of injury that may attract macrophages independent of neutrophils but would be present to a greater extent in the TB rats because of the greater injury in the TB diaphragm versus the TB+ND diaphragm. The ED1+ macrophage subpopulation was investigated because the greatest amount of change was expected to occur within this subpopulation, i.e. an early response and an association with degeneration (McLennan, 1996; St. Pierre & Tidball, 1994). The number of macrophages present in the diaphragm are not comparable to other studies, the trend however, regarding an influx of the inflammatory macrophage subpopulation following loading was similar. We found in our loaded diaphragm 8.12+1.41 (SE) and 6.90 ± 1.08 (SE) ED1+ macrophages/mm in the TB 2  and in the TB+ND groups, respectively, and 3.46 ± 0.54 (SE) and 4.87 ± 1.01 (SE) ED1+ macrophages/mm in the C and C+ND groups, respectively. St. Pierre & Tidball (1994) observed 2  an increase (p<0.05) in the ED1+ macrophages following ten days of post-hind limb suspension and 2 days of muscle loading in the soleus muscle, 350 ED1+ macrophages/mm for the 2  experimental group, compared to about 15 ED1+ macrophages/mm for the control group. 2  McLennan (1996) examined macrophage subpopulations in freeze-lesioned skeletal muscle, tibialis anterior, and although his data was not quantified, McLennan used a five point scale to describe macrophage populations. He reported very dense concentrations of EDI + macrophages dispersed throughout the cell membrane of skeletal muscle during early stages of phagocytosis  116  (McLennan, 1996). Differences in the number of macrophages in control and injured muscle in our study compared to others (St. Pierre & Tidball, 1994) may be due to several reasons, such as the characteristics of the diaphragm, i.e. a muscle required to work continuously, the nature of the load used to induce injury, and different muscle groups examined. Unloading and then reloading skeletal muscles, such as in hind limb suspension, induces greater muscle injury than increasing the load on a muscle required to work continuously. Hind limb suspension exposes the muscle to a double impact of insult. Firstly, the muscle experiences atrophy for lack of use followed secondly, by a training effect as the muscle experiences fatigue and muscle injury due to an overload when the hind limb is recruited. The hind limb muscles may have atrophied during suspension and once reloaded would experience an excessive load and result in greater injury. Hence the results from St. Pierre & Tidball have been amplified due to the effects of loading plus muscle atrophy. The diaphragm, a continuously working muscle, in the tracheal banding model did not undergo atrophy before loading. When comparing the response of a limb injury to a diaphragm injury, one may have to consider the rest period following the event that resulted in the injury. For example, in studies where eccentric activity has been performed to induce muscle injury, the subject would most likely rest the limb to overcome fatigue, this is not possible for the diaphragm. Perhaps the resting period with a relatively low blood flow allows for a greater influx of inflammatory cells. In limb muscles, the exercise stimulus causing injury is associated with an elevated cardiac output, increased regional blood flow and possibly ischaemia during high activity. In contrast, during the rest period the cardiac output would return to baseline levels and regional blood flow would greatly diminish, making it easier for inflammatory cells to adhere to the endothelial walls of  117  vessels closest to the sites of injury. In the diaphragm however, the dynamics of blood flow would be very different. The metabolic demands of tracheal banding may increase blood flow but the absence of a rest period would not allow a "low-flow" state. Thus the differing blood flow dynamics in limb muscle loading versus the diaphragm post tracheal banding may provide some explanation for the EDI values in the injured muscle. Measures of resistive loading were used as indices to determine the degree of overload induced on the diaphragm, which included the internal cross-sectional area of the trachea and esophageal pressures. Hypercapnia as determinedfromthe ABG results was another determination of the outcome of the load. A reduction in the internal cross-sectional area of the trachea by an average of 65% as shown in this study for the TB and TB+ND groups was comparable to a 66% reduction as shown by Reid et al (1994). The work of breathing was increased due to the increased resistive loading as evident by the reduction of the internal crosssectional area of the trachea. Esophageal pressures, the second index used to indicate increased resistive loading, is not the most reliable method but does provide another estimate. Comparisons performed on the same day between pre- and post-banding measurements would provide some consistency in the variables (i.e. anaesthetic dose, room temperature effects, animal state prior to surgery). However, high degree of variability between subjects within groups may well be due to pressures being monitored while the animals are under variable levels of anaesthetic and their response to loads under anaesthetic. Other factors such as tracheal secretions and spasm could affect Pes. Limitation of the PTI, shown as being greater than 0.15 for the four groups in this study, which in humans is the fatigue threshold, may have been an inaccurate indication for the rat's fatigue threshold. Elevated PTI values, in all groups, may have been due to a reduced Pes^, a  118  factor used in the calculation of the PTI. Pes  Max  presented a weakened response likely due to the  effect of the anaesthetic. In humans, normal transdiaphragmatic pressure is between 80 to 100 cmH 0, when compared to the Pes 2  Max  of rats would be a poor comparison. According to  literature review, transdiaphragmatic pressure for rats has not been determined, therefore a comparative value was not available. Another possibility, again resultingfromthe effect of anaesthesia, would be increased inspiratory time, which would increase the PTI value. A trend was observed for higher PTI values on day 1 in both of the TB groups than both of the C groups, indicating an increased effort in breathing as a result of the tracheal banding intervention. However, on day 4, the TB+ND group had a PTI value similar to the C groups. The ND intervention which showed less muscle abnormality in the diaphragm of the TB+ND group, may have better functioning capability than the diaphragm in the TB group. However, invivo or in-vitro functional testing on the diaphragm muscle may verify the assumption that due to less abnormality being evident in the muscle of the TB+ND group, the diaphragm performance of this group would be closer to the C values.  PeSyy  had significantly increased post-banding on day  1 compared to pre-banding for the TB and TB+ND groups (p<0.05, p<0.01, respectively) indicating an increased resistive loading had been applied by tracheal banding. Day 1 postbanding Pesyy values had not changed significantly when compared to day 4 values, indicating that the inspiratory load had been consistent throughout the experiment. ABGs, the third index used to assess resistive loading, showed evidence of respiratory acidosis and hypoxaemia and were reflective of the increased resistive loading and the induction of ventilatory failure. We found similar results for ABGs as shown by Reid et al (1994) where PaC0 and HCO" values had increased and Pa0 and pH had decreased. Although respiratory 2  3  2  failure was evidentfromthis study (PaC0 >45 mmHg), our results did not show the same extent 2  119  of hypercapnia as shown by Reid et al (1994) (PaC0 =97.9 Torr ± 29.6), which could be due to 2  the different species used. Hamster C groups also had high PaC0 values (51.6 Torr) (Reid et al, 2  1994) which is normal for this species. Hamsters are burrowers and thus may have a blunted hypercapnic response in comparison to rats. Respiratory acidosis was present in the TB and TB+ND groups even though there was less muscle injury in the TB+ND group. Although a significant difference was not found in the PaC0  2  between the TB (PaC0 = 57.21 mmHg ± 11.31) and TB+ND (PaC0 = 50.17 mmHg ± 9.28) 2  2  (p=0.121) groups, a significant difference was indicated in the pH of the arterial blood gases (p<0.02) between the tracheal banded groups, indicating that the state of acidosis was significantly less in the TB+ND group, which coincides with less diaphragm injury. Acidosis occurs as a physiological compensation to an acid-base disturbance. Contributing effects to acidosis for the tracheal banded groups was that they required excessive exertion to breathe for the period of 3 days of banding. Acidosis can occur due to many causes such as drug induced, toxin induced, either a secondary metabolic response (where the primary cause could be respiratory disturbance) or as primary metabolic response (Robin & Knowles, 1970). Starvation can also result in acidosis, as can renal failure (Welt, 1970). Attempts to quantify PMNs in the muscle tissue proved to be a challenge. Three methods were employed to quantify and to locate PMNs histologically in the tissue, DAB, immunohistochemistry, and determining area fractions using point-counting on H&E stained cross-sections. Initially DAB was used to distinguish the MPO enzyme found in abundance in the PMNs. The MPO enzyme cleaves the substrate hydrogen peroxide, of which the product reacts with the DAB molecule resulting in a black brown precipitate. The coloured precipitate distinguishes the presence of the MPO enzyme. We found that the DAB technique did not label  120  all neutrophils when compared to H&E stained cross-sections. The second method used an antibody directed against rat neutrophils. The same antibody had been used to target and deplete neutrophils in the circulating blood stream. However, attempts to block the secondary antibody (biotinylated anti-rabbit) from binding to the residual primary antibody (rabbit anti-rat PMN) in the tissue did not succeed. Over-staining persisted due to the secondary antibody binding to the 1. the primary antibody that had been placed on the muscle tissue for immunohistochemistry; and 2. to the residual primary antibody in the tissuefromprior injections for neutrophil depletion. Over-staining occurred even at the weakest dilution of the secondary antibody. Following the immunohistochemical attempt thefinalmethod to quantify PMNs was to determine areafractionsof muscle cross-sections stained with H&E and results are presented in Table 8. Due to morphological features and stereo interpretations it is not feasible to determine the identification of specific cells. PMNs are mobile cells and when only a portion of its multilobed nucleus is apparent in a cross-section it becomes difficult to distinguish the cell specificity without bias. The questionable category was included to distinguish cells that did not look like typical multi-lobed PMNs, howeverfibroblasts,mast cells and macrophages may also be included in this category. The degree of confidence that this investigator places on the PMN results is low. Current treatment for arthritis includes using anti-inflammatories and immunosuppressants. These have positive effects in depressing the inflammation within the joints and provides the patient with relief of pain and discomfort. However, the patient is left with anxiety and concern with regards to the high risk to personal health by opportunistic organisms (personal communication), which includes the common cold through to the devastating effects of improperly cooked foods. If a specific adherence proteinfromneutrophils is targeted to suppress the neutrophil population, this would not only provide a degree of relieffrominjury from  121  overworked muscles/tissue but also reduce the vulnerability to disease and infections by the presence of other WBCs, such as macrophages, mast cells, eosinophils, etc., which are capable of combating pathogenic organisms. From this study, we have shown that neutrophils do contribute to exertion-induced muscle injury. This study would have overcome one of its limitations if muscle histologyfromthe costal region of the diaphragm had been examined in a C group, which had not been exposed to any interventions. Baseline data for area fractions of muscle morphology and PMNs takenfroma group that had not experienced the surgical component of the experimental protocol may have provided a more normal and a better comparative group. However, due to the nature of data collection, such as measuring arterial blood gases and esophageal pressure measurements, required anaesthetizing the C animals to obtain their values. Arterial blood was obtainedfromthe carotid artery, which was cannulated during the surgical procedure. Measuring esophageal pressures included passing a water filled catheter into the esophagus which sometimes resulted in irritating the airways, and measuring the maximal esophageal pressure involved isolating the trachea in order to occlude it. These techniques could only be performed while the animal was under anaesthetic. The benefit of including a group which had not been exposed to the experimental protocol would have enabled us to compare the effects of anaesthesia and the surgical component of the experiment. Effects of anaesthesia may have included hypoventilation and post-op atelectasis which may have affected muscle morphology, although this is unlikely in such a short period. Measuring maximal esophageal pressures, however, would have required the diaphragm, of the C group, to perform greater exertion than to which the muscle is accustomed, resulting in exertion induced muscle injury.  122  Standard error was used in portraying the data for areafractionsof muscle morphology (Table 7), for area fractions of PMNs (Table 8), and for Edl+ macrophages in the diaphragm (Table 9) because the mean of the total number of fields viewed represented one animal. Standard error was then calculated to represent the distribution of the mean values for each group. Datafromthis study indicates that not only do neutrophils contribute towards muscle injury when the muscle is overloaded, but may also be involved in the process of attracting ED1+ inflammatory macrophage cells. The importance of the work described in this thesis is not so much in the provision of an exact, mechanistic explanation of how neutrophils injure the muscle fibres as it is in the discovery of their contribution in muscle injury and the revelation of the complexity surrounding the inflammatory response.  CONCLUSION From the data presented in this thesis I propose that neutrophils mediate muscle injury when excessive loads are placed on the diaphragm, and that neutrophil-mediated damage causes an increase in ED1+ macrophages. One can further conclude that neutrophils are in some way involved in the process of attracting macrophages into the diaphragm during excessing loading. Further experimentation is necessary to elucidate the exact mode of attraction of neutrophils to the site of injury and to measure functional capacity of the overloaded muscle under neutrophil depleted conditions. It will be interesting to see what future experiments will tell us about the effects of neutrophils on muscle injury.  FUTURE DIRECTIONS The scope of research in thefieldof muscle injury and the inflammatory response leads to many questions. We have seen evidence from this study that there is less muscle injury in the ND tissue. 1. This study found less muscle abnormality in the TB+ND rats, future direction would be to perform functional tests on the diaphragm and compare results between the TB and the TB+ND group. 2. Examine other muscles of inspiration from this study for injury, since preliminary examination of the SCM indicated signs of muscle injury in the TB groups. Muscle abnormality was not apparent in the SCM of the C group, although only a small portion of animals were examined. 3. Markers of muscle injury have recently included measuring the structural protein troponin in the blood of patients experiencing chest pains to evaluate the degree of cardiac muscle injury due to myocardial infarction. Creatine kinase, the standard marker for muscle injury, has been found not to be as specific for muscle injury as cell markers. Antibodies directed against troponin C (present in skeletal muscle) may provide some evidence of respiratory muscle breakdown. Although this marker would not specifically target changes in the diaphragm, it may provide information for muscle injury in the respiratory muscles experiencing excessive loads as evident in the tracheal banded animal model. 4. Further investigation of resident macrophage subpopulations following ND may add to the information provided by this study. We saw a trend towards less inflammatory ED1+ macrophages in the TB+ND group. 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Effect of hyperinflation and equalization of abdominal pressure on diaphragmatic action. J Appl Physiol. 1987;62:1655-1664.  APPENDIX A: C A R O T I D A R T E R Y C A N N U L A B: PES PRINTOUT A N D C A L C U L A T I O N S C: W B C H A E M O C Y T O M E T E R A N D C A L C U L A T I O N S  137  APPENDIX A: CATHETER PREPARATION FOR C A R O T D 3 CANNULATION Solutions:  Tubing:  2 mL Toluene 2 mL Tridodecylmethylammonium chloride (TDMAC) Heparin Complex 7% WAV (Polysciences) Saline 0.9% Heparin 1000 U/mL Intravenous medical vinyl tubing, size V3; ID: 0.58; OD: 0.99 mm.  Procedure: Measure tubing roughly 15 cm for 1 catheter. Mix 2 mL of T D M A C heparin complex and 2 mL of toluene in a glass beaker. Insert a blunted 22 gauge needle into the vinyl tubing and force the mixed solution into the tubing using a disposable 5 cc syringe. Do not allow air bubbles to enter. Allow the tubing filled with the solution to sit for 4 minutes. After 4 minutes blow dry the tubing byflushingwith air. Next flush tubing with heparin-saline solution to clear the TDMAC-toluene mix. Flush again with dry air and allow to dry for at least 4 hours. Cut and mark the tubing according to the diagram below: Diagram of Carotid cannula  Insertion Bevelled  3O  Region  Point  ~Exposed  Edge  mm  7  0  mm  2  5  mm  APPENDIX B: CALCUATION OF ESOPHAGEAL PRESSURE  ~TT#t  a. + b  =  c.  APPENDIX C : M I C R O D E T E R M I N A T I O N O F W B C C O U N T S  1. Absolute WBC Counts Working solution: 49 mL dH20 1 mL glacial acetic acid pinch of methylene blue Procedure: 1. Mix all ingredients in a vial and dispense 38 uL into a microcentrifuge tube. 2. Swab rat tail with heparin. 3. Rinse pipette tip with heparin. 4. Cut the very tip of the rat's tail, swab the tail again with heparin, discarding the initial drop of blood to remove any tissue or hair particles. Draw 2 uL of blood into the pipette tip and immediately place in the vial containing 38 uL of counting solution. The tail may need milking if it does not bleed adequately. 5. Mix the vial well by tapping the vial with your finger. 6. Inject 10 uL into the haemocytometer countingframe(see figure below) which had a coverslip upon it. 7. Count 16 squares from each quadrant of the counting chamber. 8. The following equation was used to determine absolute WBC million/mL: Count in 64 squares x dilution factor = million/mL (Dilution factor = blood volume X total volume)Differential WBC Counts Haemocytometer  1  mm  140  2. Absolute Circulating PMN counts 1. Place a drop of blood directlyfromthe rats tail, which the tip has been cut with a sharp blade, onto a glass slide and using the edge of a second glass slide smear the drop of blood 2. Fix the smeared slide by dipping the slide in methanol and then allow to air dry. 3. The slide is then loaded onto an autostainer and is stained with Wrights solution. Wrights' stain stains the cell nuclei blue and the cell cytoplasm a pinkish colour. 4. The blood smear slide is then coverslipped. 5. The cells were counted at a 400x magnification which allows cells to be differentiated. 6. Cells were counted according to two categories: PMN or non-PMN. Cells were counted until 100 cells had been counted from either category. 7. %PMNs were calculated by determining: number of PMNs total number of cells counted x 100 8. Absolute PMNs was determined by calculating: %PMNs x Absolute WBC = millions/mL  


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