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Calcium activated neutral protease (calpain) and the neutrophil: their relationship and association with.. Raj, Daniel Adelbert 1997-03-10

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CALCIUM ACTIVATED NEUTRAL PROTEASE (CALPAIN) AND THE NEUTROPHIL: THEIR RELATIONSHIP AND ASSOCIATION WITH THE ACUTE INFLAMMATORY RESPONSE TO EXERCISE by DANIEL ADELBERT RAJ B.H.K. The University of British Columbia, 1994 A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF MASTER OF SCIENCE in THE FACULTY OF GRADUATE STUDIES School of Human Kinetics We accept this thesis as confonmng to the required standard THE UNIVERSITY OF BRITISH COLUMBIA March, 1997 © Daniel Adelbert Raj, 1997 In presenting this thesis in partial fulfilment of the requirements for an advanced degree at the University of British Columbia, I agree that the Library shall make it freely available for reference and study. I further agree that permission for extensive copying of this thesis for scholarly purposes may be granted by the head of my department or by his or her representatives. It is understood that copying or publication of this thesis for financial gain shall not be allowed without my written permission. Department of VWvncxn KAoeA^CS, The University of British Columbia Vancouver, Canada Date 2-^-1^ MexccrW IH^? DE-6 (2/88) ABSTRACT The purpose of this study was to determine whether exercise-induced neutrophil accumulation into striated muscle could be assisted by locally produced myogenic factors. It is hypothesized that a relationship exists between the processes of calpain-mediated muscle injury and the recruitment of neutrophils to the injured sites. To test this hypothesis, male Sprague-Dawley rats (-325 g) were randomly assigned to either a control (n = 10) or an experimental (n = 10) group, where the experimental protocol involved 1 hour of -14% grade running at 25 m/min on a motorized treadmill. Neutrophil chemotaxis assays were conducted on extracts of whole tissue samples of the heart, soleus, and plantaris; processed in phosphate buffered saline for the membrane soluble and cytosolic protein fractions. Chemotaxis was quantified in triplicate across all conditions and normalized to the maximal chemotactic response at 10"7M/MLP. In the membrane soluble fraction, neutrophil chemotaxis was increased in the skeletal muscles following exercise: Condition Control (% max chemotaxis) Exercise (% max chemotaxis) Significance Soleus 9.03 ± 3.03 46.70 ±1.91 p < 0.05 Plantaris 9.31 ± 1.25 22.31 ±4.06 p < 0.05 Heart 7.74 ± 2.73 4.28 ± 1.83 p > 0.05 However, all muscle extracts from the cytosolic fraction reflected a significant decrease in neutrophil chemotaxis following exercise: Condition Control (% max chemotaxis) Exercise (% max chemotaxis) Significance Soleus 31.79 ±14.41 9.26 ±4.63 p < 0.05 Plantaris 16.16 ±4.68 4.97 ± 2.49 p < 0.05 Heart 47.14 ±13.69 10.92 ± 4.74 p < 0.05 ii These results indicate that there exists a neutrophil chemoattract(s) in muscle. It is plausible that this chemoattactant(s), initially contained within the cytoplasm, could feasibly leak out into the interstitium/circulation mainstream during exercise. To investigate a relationship between calpain-like activity (Ca2+-stimulated proteolysis) and neutrophil accumulation as measured by myeloperoxidase (MPO) activity, these activites were compared in cardiac and plantaris muscles from male Wistar rats (n = 10) completing 1 hour of running exercise (25 m/min). Exercise promoted increases (p < 0.05) in both calpain-like and MPO activities; ranging from 2.79 to 58.9 U/g wet wt and 0.03 to 4.88 U/g wet wt respectively. Pearson's correlational analysis (r) on calpain-like and MPO activities for cardiac and plantaris data were 0.97 (p < 0.001) and 0.68 (p < 0.05) respectively; with a combined r = 0.83 (p < 0.001) for both muscles across all conditions. To further investigate the extent to which calpain-like activity may promote neutrophil accumulation, another exercise group (n = 5) was pre-injected with the cysteine protease inhibitor, E64c, 1 hour before exercise. Administration of E64c lowered calpain-like and MPO activities by 66% and 56% respectively (average from cardiac and plantaris muscles). From these results it is concluded that: 1) a relationship exists between Ca2+-stimulated proteolysis and neutrophil accumulation into striated muscle with exercise; and 2) the calpain system is involved in localizing the neutrophilic response associated with exercise. iii TABLE OF CONTENTS ABSTRACT TABLE OF CONTENTS LIST OF TABLES LIST OF FIGURES ACKNOWLEDGMENT INTRODUCTION OBJECTIVES.... HYPOTHESES LITERATURE REVIEW Eccentric Exercise and Muscle Damage Acute Inflammatory Response Neutrophils Chemotaxis and Chemotactic Agents Myeloperoxidase Calpain RESEARCH METHODS Animal Care and Training Exercise Protocol — Chemotaxis Muscle Preparation Chemotaxis Research Design Isolation of Neutrophils Chemotaxis Protocol Exercise Protocol - Correlation Calpain Assay Myeloperoxidase Assay STATISTICAL ANALYSES Correlation Analysis RESULTS Neutrophil Chemotaxis Relationship: Calpain and Myeloperoxidase DISCUSSION CONCLUSION FUTURE RECOMMENDATIONS BIBLKXJRAPHY 72 APPENDK 81 Polymorphonuclear Cell Isolation 82 Reagents 3 Protocol for Neuro Probe™48 well chemotaxis chamber :. 84 Microplate Assay for Calpastatin-inhibited Calcium-dependent Proteolysis - Calpain-like Activity 89 Myeloperoxidase Assay 93 v LIST OF TABLES Table 1. Control vs exercise muscle tissue neutrophil chemotaxis — done in triplicate.... 41 Table 2. Summary of 2 (Condition) x 3 (Muscle-type) 2-way ANOVA 49 Table 3. Calpain substrates (adapted from Saido et al., 1994) 81 Table 4. Some of the proposed physiological functions of the calpain system (adapted from Goll et al., 1992) 8Table 5. Results: individual sample means for neutrophil chemotaxis 94 LIST OF FIGURES Figure 1. Proposed sequence of events following exercise induced muscle damage 4 Figure 2. Schematic representation of calpain. The primary amino acid sequence(s) of neutrophil chemotactic regions are shown 28 Figure 3. Schematic representation of muscle tissue processing 40 Figure 4. Schematic representation of modified 48-well Boyden chamber set-up 43 Figure 5. Bar-plot representing neutrophil chemotactic potential in the cytosolic fraction of muscle extracts. Neutrophil chemotaxis as normalized and plotted to a percentage of maximum (100%) of 10"7M/MLP. Results are presented as means ± S.D.; * denotes significant difference from control at p < 0.05 52 Figure 6. Bar-plot representing neutrophil chemotactic potential in the soluble fraction of muscle extracts. Neutrophil chemotaxis as normalized and plotted to a percentage of maximum (100%) of 10"7M/MLP. Results are presented as means ± S.D.; * denotes significant difference from control at p < 0.05. 53 Figure 7. Pearson's correlational analysis (SPSS for Windows; version 6.1) illustrating the: Relationship between myeloperoxidase and calpain-like activity of the combined data-set for both plantaris and cardiac muscles for both resting and exercised conditions. Based on individual comparisons, where exercise consisted of a 25 m/min treadmill run on an 8% grade for 60 min 55 Figure 8. Pearson's correlational analysis (SPSS for Windows; version 6.1) illustrating the: Relationship between myeloperoxidase and calpain-like activity of plantaris muscle for both resting and exercise conditions. Based on individual comparisons, where exercise consisted of a 25 m/min treadmill run on an 8% grade for 60 min 56 Figure 9. Pearson's correlational analysis (SPSS for Windows; version 6.1) illustrating the: Relationship between myeloperoxidase and calpain-like activity of cardiac muscle for both resting and exercise conditions. Based on individual comparisons, where exercise consisted of a 25 m/min treadmill run on an 8% grade for 60 min 57 vii Figure 10. Bi-axial bar plot illustrating the effects of E64c pre-treatment on myeloperoxidase and calpain activities following treadmill running. Values are means • S.D.; * denotes significant difference from control at p < 0.05. Exercise consisted of a 25 m/min treadmill run on an 8% grade for 60 min. Runners in the E64c group were pre-injected 1 hour prior to the running protocol 58 viii ACKNOWLEDGMENT I would like to thank: Dr. Angelo N. Belcastro Dr. Blair A.M. Walker & Corinne Rocchine Dr. Darlene W. Reid Dr. Donald C. McKenzie Gavin D. Arthur Leann D. Shewchuk Tracie A. Albisser Timothy S. Booker ChadG. Ball Lorna Christie Micheal Boyd and his crew at the animal resource unit INTRODUCTION It is now well established that exercise above moderate intensity (-70% VC^max) causes changes to skeletal muscle reminiscent of both macro- and micro-injury (i.e. injury to whole fibers vs isolated lesions to specific sections in the fiber). These effects of exercise often initiate reactions that resemble the "acute phase response" to infection [12,19] and inflammation [28,70,79,98]. This includes proteolysis of skeletal muscle, mobilization and activation of neutrophils [63,70,98], and increased hepatic production of certain acute phase proteins [12,79]. The physiological changes accompanying this sequelae of events in the acute inflammatory response to exercise have been previously reported [19,20]. However, the mediators and effectors underlying the coordinated response to infection, and inflammation to the region of muscle damage [30,79] remain to be fully elucidated. Studies employing histological and electron microscopic techniques have shown that striated muscle injury is typically characterized by such events as, mitochondrial and sarcoplasmic reticular vacuolization [2,33], and sarcomeric Z-line disruption [2,20,22,30]. These changes are reminiscent of the actions of the calcium activated neutral protease, calpain [37,73,93,104], which have since been implicated with exercise-induced muscle injury [6-8]. As typically seen with strenuous exercise, these myofibrillar alterations have also been associated with the leakage of such enzymes as creatine kinase (CK) [50,51], lactate dehydrogenase (LDH) [38,46] and the release of myosin heavy chain (MHC) 1 [60,61] and the troponins (Tn) [59] from the muscle. With exercise induced muscle injury, neutrophils are the first inflammatory cells present at focal sites of lesion, rising in numbers (>3-fold) immediately after insult and peaking within 12 hours [70,79,98]. That neutrophils accumulate into striated muscle is evidence by tissue myeloperoxidase activity which has been reported to increase by >30% following exhaustive running exercise in rats [8]. Although there is evidence for increased number of neutrophils within skeletal muscles following exercise, the precise mechanism(s) responsible for their margination and emigration (i.e. chemotactic potential) is unknown. Since muscle protease activity and subsequent protein breakdown occur early on in the events mediating injury, the first objective of this study was to assess the extent to which neutrophils would readily chemotax to muscle immediately following exercise. This was achieved by conducting neutrophil chemotaxis assays on prepared cytosolic and membrane-soluble muscle extracts from exercised and sedentary rats. The second aim of this study was to investigate a possible link between the actions of striated muscle calpain and neutrophil accumulation. Since this accumulation of neutrophils appear as the first mflarnmatory occurrence following initial damage to muscle, assessment of its involvement in promoting/augmenting muscle proteolysis is warranted. The hypothesis proposes (Figure 1) that following the activation of calpain through elevated levels of intracellular calcium within the muscle, selective proteolysis of various contractile and cytoskeletal protein elements occurs. Hence, peptides fragments are produced which act as chemotactic signals to neutrophils, drawing them from the peripheral circulation towards focal sites of tissue damage. The early assessment is crucial if a link is to be established 2 between the actions of calpain and neutrophil recruitment, since calpain is now known to be activated as early as 5-10 minutes following the initiation of exercise (unpublished findings). Partial support for a direct link is evidenced from various sequences of fragmented calpain peptides which have shown potential to stimulate neutrophil chemotaxis [54]. Along with calpain peptide fragments, cytoskeletal peptide fragments from early calpain-mediated proteolysis could also be likely chemotactic candidates. Thus, demonstrating a positive relationship between the activities of calpain and myeloperoxidase is critical in supporting the hypothesized link between calpain activation during muscle protein breakdown and neutrophil accumulation. To test this hypothesis, individual animal comparisons were made between the activities of tissue calpain and myeloperoxidase in striated muscle homogenates from exercised and sedentary rats. Calpain involvement was further implicated with the use of the cysteine protease inhibitor, E64c, which was able to prevent exercise-induced increases in the activities of calpain and myeloperoxidase. The extent of initial tissue damage is believed to pre-determine the muscle's fate, inasmuch as its potential for recovery. In studying this early stage of tissue damage, this investigation hopes to contribute to the current understanding on the mechanism(s) underlying tissue injury. 3 Exercise * Ca2+ Imbalance t Calpain Activation t Neutrophil Accumulation [Ca2+] Calpain Fragments/Peptides? ^ <s> Neutrophils Neutrophils ^ (Circulation) <m Figure 1. Proposed sequence of events following exercise induced muscle damage. OBJECTIVES • To quantitate immediately following exercise, neutrophil chemotaxis in the membrane-soluble and cytosolic fractions from muscle preparations of the soleus, plantaris and the heart. • To correlate at rest and immediately following exercise, the activities of calpain and myeloperoxidase from the plantaris and heart muscle. • To quantitate immediately following exercise, the activities of calpain and myeloperoxidase from die plantaris and heart muscle, when animals are pre-treated (1 hour) with the cysteine protease inhibitor, E64c. 5 HYPOTHESES • That compared to control, muscle extracts of both the cytosolic and membrane soluble fractions from exercised rats will demonstrate a significant increase in neutrophil chemotaxis in muscles of the heart, soleus and plantaris. • That compared to control, muscles of both the heart and plantaris from exercised rats will demonstrate significant increases in both calpain and myeloperoxidase activity, and that these will show a significant positive relationship. • That pre-exercise treatment with E64c, the cysteine protease inhibitor, will significantly suppress exercise-induced increases in calpain and myeloperoxidase activity. 6 LITERATURE REVIEW Eccentric Exercise and Muscle Damage Within the last 10 years, there has been increased interest in how muscles resist externally applied forces and how eccentric muscle action leads to delayed-onset muscle soreness (DOMS) [79,87]. It is now established that the eccentric component of muscular contraction, as opposed to concentric or isometric, is the predominant cause of muscle damage and the closely related sensations of DOMS. Typically, an eccentric contraction involves the generation of force while the muscle is lengthening. Since almost all forms of physical activity consist of some combination of concentric, isometric and eccentric muscle contractions, much research has questioned which of these is chiefly responsible in producing muscle damage and the pain experienced during DOMS. Stauber [87], Evans and Cannon [20] have described the actions of eccentric-type contractions, and how they are capable of generating more force than concentric or isometric contractions. They have highlighted that the physiological metabolic cost of perforaiing "negative work" (i.e. eccentric action) is substantially less than performing "positive work" (i.e. concentric or isometric actions) — requiring a smaller oxygen uptake (less VO2), involving lesser recruitment of motor units (reduced EMG) and therefore also decreased energy utilization and requirement (of substrates and metabolites) for the active muscles to develop tension while being stretched. 7 Clearly eccentric-type contractions have their advantages. Moreover, given that it requires fewer muscles fibers and lesser energy to generate more force with eccentric contractions, it is not difficult to visualize how the body may be inclined to exploit this biomechanical efficiency, especially during fatigue. Since these eccentric muscle actions ~ as opposed to concentric or isometric actions — are capable of generating more force and tension within the muscle, they are undoubtedly the prime candidate implicated in muscle damage. Based on studies assessing blood flow to muscles during exercise, comparatively, fast-twitched (type II) as opposed to slow-twitched (type I) muscle fibers are reportedly more susceptible to exercise-induced damage [56,57]. This is plausible since blood flow to exercising muscles is preferentially directed to the oxidative muscle fibers to fuel their increased oxygen requirements. Hence, these more oxidative muscle fibers would also have an expectedly increased capacity to remove damaged muscle debris, and acquire hepatically produced plasma proteins and amino acids from the blood for muscle tissue regeneration. To suggest that myofibrillar damage is complimented/augmented by neutrophil responses to exercise, this study proposes an indirect assessment of this occurrence in an exercise-adapted state. Since eccentric-type contractions exert a greater tension on muscle fibers and/or adjacent connective tissues, resulting in muscle damage and the associated inflammatory reaction, they will also be expected to elicit a more dramatic neutrophil response over concentric-type contractions [2,18]. Previous studies describing inflammatory cells related to this topic have primarily focused on the assessment of plasma concentration from various leukocyte populations [18,29,34,40,68], and tissue biopsies of 8 certain muscles [20,30,78]. Results from studies assessing leukocyte concentration from blood plasma have been far from conclusive. Both increases [68] and decreases [40] have been reported from seemingly similar protocols. This may be explained by possibly different methods and assays used for the various dependent variables. Still, it is important to realize that concentrations of blood-borne variables represent a constant flux of synthesis and breakdown. Therefore, the variations seen may be inherently due to the nature of blood being primarily a medium for transport. In this regard, the assessment of muscle-specific samples would expectedly facilitate clearer understanding of muscle function. However, tissue biopsies are also problematic [79]. With exercise-induced muscle damage, the muscle's response to an exercise stimulus must be expected to be different depending on: the locale of tension (e.g. musculotendinous junction vs muscle belly), muscle fiber type (e.g. fast- vs slow- twitched), and the level of innervation (i.e. number of muscle fibers recruited per motor unit). Since human muscles are not homogenous throughout, a muscle biopsy can therefore only be expected to represent at best, a very localized area and not the whole muscle. To this end, the assessment of exercise-induced muscle damage in this study will be based on whole tissue analysis. More pain (i.e. DOMS) has been linked to muscle damage with downhill running ~ requiring more eccentric muscle action than level or uphill running in humans [22,87]. Likewise, under similar running conditions, muscle damage and its associated inflammatory response has been observed in rats [2]. It seems peculiar how downhill running requiring a lower metabolic cost, and clearly less effort than either level or uphill 9 running, is still believed to cause more inflammatory and degenerative changes to muscle [2,87]. More recently, this relationship between muscle tissue damage and the inflammatory response has been characterized in both humans [18,20,29,30,34] and rats [2,66,78]. Accordingly, the rat serves as an appropriate animal model in the study of exercise-induced muscle damage. Furthermore, since the rat can be easily trained to run on a treadmill [56], it is thus far the animal model of choice for the purposes of this study. 10 Acute Inflammatory Response Inflarnmation is characterized by the movement of fluid, plasma proteins and leukocytes into tissue in response to injury, microbial invasion, foreign material or antigens [98]. This development of an inflammatory response is an important mechanism by which organisms defend themselves against pathogenic agents — by initiating both structural and functional repair of damaged tissues [28,70,79]. It manifests itself by means of an acute inflammatory response, and depending on the extent of the pathogenic insult, the development of a chronic inflammatory response may follow [98]. Chronic inflarnmation can result in a range of outcomes. Here resolution could vary from complete elimination of the pathogenic agent and injured tissue, resulting in repair and return to normal tissue structure and function; to more pathological states involving irreversible injury of tissue, resulting in scar tissue formation and loss of tissue function [98]. While chronic inflammation serves as an end-point to the inflammatory process, it is the acute inflammatory response that sets the stage for the events that will follow. By its initial reaction to injury, the processes occurring during this acute phase will determine the extent and magnitude of the triggered response required [70]. Thus, in exploring the underlying mechanisms related to acute muscle injury, assessment of the early events which trigger this response seem justified. The acute inflammatory response is a sequence of defense reactions that are typically triggered by tissue damage and infection. Although more recognized for its role in antibacterial and antiviral actions, the acute inflammatory response also promotes the 11 clearance of damaged tissue and sets the platform for repair and growth [28,70]. Since exercise-induced damage to muscle must also undergo similar stages of repair and growth, the initiation of the acute inflammatory response has also been implicated in muscle's adaptation to exercise [70,79]. Furthermore, with the classical signs of inflammation: pain, swelling, loss of function, heat and redness being very similar to the effects of DOMS, the relationship between exercise-induced muscle damage and the acute inflammatory response is now more apparent [20,79]. As reviewed by Smith [79], following exercise that involves unaccustomed eccentric muscle action, connective and/or contractile tissue disruption occurs. Almost immediately after a strenuous bout of exercise, the circulating levels of neutrophils rise significantly. These neutrophils then invade the site of injury, exerting their effects for several hours. Next, about 6-12 hours post-exercise, monocytes infilttate into the injured area. Their levels at the site of injury then rise steadily to a peak after around 48 hours, and are non-existent by 72 hours after exercise. These monocytes then mature into adult macrophages, which can synthesize large amounts of PGE2 upon exposure to the inflammatory environment. From this, PGE2 sensitizes the Type III and IV "pain" afferents, causing them to become hypersensitive to stimulation. In addition, the edema associated with DOMS causes an increased intramuscular pressure within the muscle tissue compartment. These combined effects result in a non-compliant, "tight" muscle compartment that is highly sensitive to mechanical stimuli. As such, the slightest movement or palpation, serves as sufficient mechanical stimulus for the sensations of DOMS to be experienced. 12 The extent to which this acute inflammatory response is elicited by exercise has since been shown to be duration-dependent [28]. However, the relationship between exercise intensity and the acute inflammatory response is still illusive. Results from an earlier study (PNWEG '95) with concentric-type uphill running, suggest that when equated on caloric expenditure, the magnitude of neutrophil activation is both intensity-and dose-dependent. Discrepancies seen in the literature may in part be due to experimental complications which arise owing to the progressive nature of the acute inflammatory response. Typically the effects of this response are transient and sequential, with some elements having a very rapid onset within minutes (i.e. complement factors, neutrophils), while other elements involving time-dependent transport processes (i.e. trace metals, acute phase proteins) may require hours or even days to take maximal effect [28,98]. Therefore, the time of sampling is extremely crucial to the characterization of a particular element of interest within the acute inflammatory response. Hence, the justification for proposing immediate sampling of skeletal muscle tissue post-exercise. From this, the results of this study will hopefully describe the early effects of exercise to fatigue and its propensity to elicit the generation of the acute inflammatory response. 13 Neutrophils Neutrophils constitute ~60% of the circulating leukocytes. Characteristic to the neutrophil is a lobulated chromatin-dense nucleus, which gives rise to its being called the polymorphonuclear leukocyte. Either alone or in conjunction with antigen-specific defenses, they serve as the first-line-of-defense against foreign agents [70]. The neutrophil is the most abundant cellular component found in the human immune system, possessing microbicidal, bacterial and viricidal activities [70]. For its wide range of effector functions, the neutrophil has therefore been considered critical to host defense. Following infection or injury, the number of neutrophils circulating in the blood has been documented to increase several fold [28,98]. With acute inflammation, the peripheral neutrophil content rises from 5000/pl to as high as 30,000/pl [98]. Immediately following 20 minutes of downhill running at 60% V02max, plasma neutrophil counts were increased by 33% and plasma concentrations of myeloperoxidase (an enzyme marker for neutrophil content) increased by 97% [18]. Likewise, following 45 minute bout of downhill (16°) running, histological staining revealed a 67% increase in neutrophil accumulation in skeletal muscle 45 minutes post-exercise [30]. This rapid increase is possible, since at rest only about half of the mature neutrophils produced by the bone marrow are within the blood circulation. The other half are adhered onto endothelial cells along blood vessels, thereby remaining temporarily out of the circulation mainstream. Upon activation, these so called marginated neutrophils then move into the circulation to perform their cytotoxic activities. The actual migration of the neutrophil to specific sites 14 of injury is dynamic process. This process is augmented by alterations in vascular caliber that lead to an increase in blood flow, and structural changes in the microvasculature that allow the neutrophils to leave the circulation [24]. The fate of an inflammatory neutrophil in the lumen of the blood vessel involves its initial re-margination, rolling and adhesion onto endothelial cells at the site of injury — a process regulated by the expression of receptors on the surfaces of the neutrophil (i.e. selectins, integrins) and the endothelial cell (i.e. intercellular adhesion molecules: ICAM, vascular cell adhesion molecules: VCAM). This is followed by the inflammatory cell's transmigration across the endothelium; a process also known as diapedesis. Finally, the neutrophil migrates in the interstitial tissues towards the chemotactic stimulus. Clearly, the increased number of neutrophils within injured skeletal muscles are facilitated by some mechanism which brings about an increase in the marginal neutrophil pool at focal regions of tissue damage [66]. This could be particularly evident with strenuous exercise, where an increase in cardiac output causes an increased blood perfusion and blood pressure. In so doing, this may dislodge the neutrophils from their storage sites into the circulation mainstream [98]. However, the exact mechanism involved in the actual movement of the neutrophil, from its circulatory pool into damaged tissues regions is still unclear. Depending on the extent of tissue damage, further neutrophil recruitment may occur during the acute response. Since the bone marrow pool represents a neutrophil population 20 times larger than that in the circulation, it serves as an extended reserve for neutrophils [98]. In response to such inflammatory mediators as tumor necrosis factor (TNF), interleukin-1 (LL-1) and endotoxin, pools of neutrophils within the bone marrow are mobilized into the circulation [20,30,98]. TNF-a and IL-ip are thought to act synergistically to induce specific myofibrillar proteolysis [12,30], perhaps by releasing chemoattractants which draw neutrophils and other leukocytes (e.g. macrophages, eosinophils) to the inflammatory site. Neutrophils have the propensity to phagocytize pathogens and tissue debris. They are also capable of releasing an array of cytotoxic factors, which include proteolytic enzymes (i.e. elastase, collagenase) [18,28] and oxygen radicals [62,65-67]. Each of these factors will react specifically with targeted tissues, to promote the ensuing cascade of events to follow in the inflammatory response. For example, these factors can break down basement membranes of the microvasculature near the site of tissue injury, thereby further increasing vascular permeability which could promote the migration of leukocytes into damaged areas [28]. However, these cytotoxic agents of the neutrophil are also the basis of several inflammatory diseases, since they have no regard for foreign or host tissue, and will as readily break down healthy tissue if not kept in check. As such, the recruitment of neutrophils must be closely regulated to specific sites of injury by chemoattractants such as chemokines and complement factors. This provides tight regulation for the recruitment, and directing of inflammatory cells to the site of injury. Neutrophils in response to chemoattractants, orientate toward the pathogenic stimulus by forming a knob-like protrusion toward it. It is through this process that neutrophils and other leukocytes alike, are thought to "sense" target stimuli [98]. Chemotactic factors and protein fragments released as a result of tissue degradation, are thought to "chemically attract" inflammatory 16 cells to the site of infection or injury [70,98]. Nevertheless, specific work towards this regard has not been done with exercise-induce muscle tissue damage. 17 Chemotaxis and Chemotactic Agents Neutrophils are mobile cells. This motility plays a central role in the ability of these cells to accumulate at inflammatory sites — initially migrating from the bone marrow into the circulation, then to adhere onto endothelial surfaces of blood vessels, and finally to migrate towards the inflammatory focus. Chemotaxis is defined as the directed movement of cells or organisms towards a concentration gradient of some mediator/chemical stimulus [48,92]. Clearly, chemotaxis is different from normalized undirected movement and random motility where cell movement occurs with equal probability in all directions or chemokinesis. Chemotaxis deals with direction of locomotion, and is distinct from chemokinesis which refers to the alteration in speed of random migration. Chemoattractants can accelerate the random movement of cells (chemokinesis), possibly by activating metabolic and contractile systems within the cell; when this cellular regulation is dictated by concentration gradients, the chemoattractants cause cell locomotion to be directed (Chemotaxis). A variety of substances sometimes referred to as cytotaxins are known to demonstrate chemotactic activity. For example, the complement fragment C5a, fibrinopeptides, kallikrein, prostaglandins, casein, neutrophil-derived factors, bacterial-derived factors and certain oligopeptides all directly attract neutrophils [92]. Substances such as antigen-antibody complexes, endotoxin and zymosan are not directly chemotactic but they can act secondarily as chemotactic mediators by inducing the production of a chemotactic factor (i.e. C5a). Hence, chemotactic factors can initiate both chemokinesis and chemotaxis in the neutrophil. 18 When unstimulated neutrophils are spherical in morphology. Upon treatment with chemotactic agents (i.e. /MLP) neutrophils become polarized even in the absence of a chemoattractant gradient. This polarizing ability may be an intrinsic response of the cell to orientate themselves prior to movement towards a particular direction. Neutrophils are believed to sense stimulation by either having an asymmetrical distribution of receptors on the plasma membrane, or by possessing a capacity to rapidly redistribute receptors to the front end of the cell. Hence, their polarization toward the chemoattractant. In order for chemotaxis to be maintained along a concentration gradient, there must exist a continuous re-expression of receptors via recycling of receptors, or the mobilization of new receptors at the front "sensing" end of the cell. When unstimulated there are approximately 15,000 receptors per neutrophil, and this number can be rapidly up-regulated via Ca2+-dependent processes to 30,000 - 50,000 through the mobilization of subcellular internal stores, at a rate of up to 10,000 receptors/min [27]. Neutrophils possess a complex system of microtubules and microfilaments which significantly contribute to its motility. Accordingly, neutrophil chemotaxis is inhibited when microtubule-microfilament inhibitors such as colchicine and cytochalasin B are used in vitro [92]. Following membrane receptor recognition, occurs electrolyte shifts with calcium and sodium influx leading to membrane depolarization and cell activation. Almost all receptor-mediated neutrophil functions are mediated by GTP-binding proteins. G-proteins serve a the link between membrane receptor occupancy and activation of such intracellular enzymes as the phospholipases and protein kinases [27]. 19 It is now recognized that the many functions of the immune system are carefully regulated by cytokines. Cytokines are small (8.5-40 kDa) polypeptides that serve as intercellular signaling molecules for a wide range of local and systemic responses. Monocytes and macrophages are probably the major source of cytokines, however neutrophils, T-cells, endothelial cells and fibroblasts also have the capacity to synthesize and secrete pro-inflammatory cytokines. Cytokines are typically subdivided into several families based on structural similarities of their receptors or by their biological activities [86]. Grouped for their biological similarities, the pro-inflammatory subset of cytokines which include tumor necrosis factor-a (TNF-a), interleukin-1 (IL-1), interleukin-6 (IL-6), interleukin 8 (IL-8), monocyte chemoattractant protein-la (MCP-la) and macrophage inflammatory protein-1 (MIP-1) belong to several different cytokine families. While numerous cytokines can mediate similar responses, on its own, a cytokine can also have multiple and divergent effects. For example, while IL-1 (3, IL-6, TNF-a and MIP-1 are all pyrogenic (fever inducing), singly, EL-8 is a neutrophil chemotactic and activating factor, and MCP-1 is chemotactic for monocytes. Bacterial lipopolysaccharide (LPS) [94] otherwise known as endotoxin, is believed to initiate the inflammatory process through the induction of the pro-inflammatory cytokines interleukin (J-L)-l and tumor necrosis factor (TNF). IL-1 and TNF in turn contribute to the progression of acute inflammation by inducing the expression other chemoattractant and adhesion molecules, such as CINC and MIP-2. For its numerous influences on the neutrophil even at minute concentrations (ng), endotoxin is always a 20 cause for concern in any neutrophil preparation. As such, meticulous care should be taken to minimize the possibility of its contamination. TNF is a macrophage-derived regulatory cytokine induced in vivo by LPS [90]. In certain instances, EL-1 and TNF act synergistically to produce a concerted response (e.g. fever, B and T cell activation). Some of the effects of TNF on endothelial cells include general activation, promotion of neutrophil adherence, induction of procoagulant activity and direct cytotoxicity. TNF-stimulated neutrophil functions include adherence, aggregation, chemotaxis, hydrogen peroxide and superoxide anion generation, degranulation, phagocytosis, fungicidal activity and antibody-dependent cellular cytotoxicity. While numerous cytokines have been implicated in triggering neutrophil chemotaxis, aggregation and degranulation, interleukin-8 (IL-8) has since received much attention. DL-8 is a small (8 kDa) protein originally purified in from culture supernatants of LPS-stimulated monocytes and/or macrophages [27]. In addition to LPS, inflammatory mediators such as zymosan, IL-1 and TNF-a can also induce IL-8 production by macrophages, and certain other cell types: endothelial cells, fibroblasts, epithelial cells and hepatocytes. IL-8 affects neutrophils in a similar fashion as the chemotactic complement C5a and the formyl peptide /MLP [26]. Like them, LL-8 is an extremely potent chemotactic agent for the neutrophil, inducing changes in shape and increases in the expression of surface adhesion molecules of the cell. As well, EL-8 is capable of triggering the release of proteolytic enzymes, the production of oxygen radical species and elevations in intracellular free Ca2+ in human neutrophils. Currently, EL-8 is a favorable chemotactic 21 mediator in attracting neutrophils during the acute mflammatory response to muscle injury, since it has more cell selectivity than C5a, /MLP, PAF and LTB4, having litde (if any) chemotactic affinity for eosinophils or basophils, and no effect on platelets or monocytes and macrophages [4]. The cytokine-induced neutrophil chemoattractant (CINC) is a member of the C-X-C (alpha) or IL-8 surjerfamily in rats. It is highly chemotactic for rat neutrophils and is known to be induced by IL-1, TNF and endotoxin. While numerous chemotactic peptides have been identified in a variety of species, a strict IL-8 analogue has not been identified for the rat. As a result, EL-8 per se has yet to be directly implicated in neutrophil recruitment in the rat model. However, for its cell selectivity and chemotactic affinity, IL-8 and/or its rat analogue CINC do appear to be very likely neutrophil chemotactic candidates. CINC exhibits its most significant homology to human melanoma growth stimulatory activity MGSA/GRO (69%) and to the mouse KC (91%) gene product [101] -- both of which are related to IL-8, the most topical of the C-X-C family of neutrophil chemoattractants. To date, IL-8 and CINC have not been shown to be produced specifically by skeletal muscle cells, but surrounding endothelial cells, resident macrophages and fibroblasts undoubtedly have this capacity. Both EL-8 and CINC have been implicated in numerous inflammatory disorders of the heart, lung, kidney, intestinal tract and smooth muscle. As such, their possible involvement in the neutrophil's response to exercise-induced muscle injury in this study deserves consideration. 22 Myeloperoxidase Myeloperoxidase (donor: hydrogen peroxide oxyreductase, EC is a highly cationic, iron containing protein located within the azurophilic granules of mature neutrophils [58,67,102]. It is a 120-150 kDa glycoprotein which exhibits three major isometric forms designated: MPO I, MPO II and MPO III [65]. Native MPO is composed of a pair of heavy-light protomers, each containing a heavy chain (a) of 59 kDa and a light chain ({$) of approximately 13.5 kDa [67]. While all the isomeric forms of MPO have the same sized (3-subunit, they differ in the size of the oc-subunit. The three major isoforms contained within the neutrophil exhibit differences in molecular weight, cationicity, hydrophobicity, sensitivity to inhibitors and localization within azurophilic granule subpopulations [65]. Of these, MPO I is the largest and most hydrophobic, while MPO lU is the smallest and most cationic. However, the three species do possess similarities. They are recognized by the same monoclonal antibodies, have identical spectra, the same P-subunit amino acid composition, the same specific activity for H2O2, identical prosthetic groups and same carbohydrate content [67]. Myeloperoxidase plays an important role in the host defense against invading microorganisms [62] by aiding in the production of cytotoxic oxygen radical species. Being the most abundant protein found in the human neutrophil, it is clear how the quantification of MPO could arguably serve as an excellent marker for neutrophilic granulocytes within blood and tissue [78]. MPO oxidizes a variety of compounds in the 23 presence of hydrogen peroxide, and in doing so, it plays a crucial role during neutrophilic phagocytosis [47,62,67]. MPO catalyzes the conversion of H2O2 and CT into the potent oxidant, hypochlorous acid (HOC1) via reactions (1) and (2); where MP3* is ferric myeloperoxidase (i.e. MPO with three positively charged ions), and Compound I is MPO in a transitional state [47]: (1) MP3* + H2O2 <=> Compound I (2) Compound I + CI" <=> MP3* + HOC1 It is this HOC1, produced from the MPO catalyzed reaction above, that is responsible for the microbial activity of the enzyme [102]. The presence of MPO in this system augments the microbicidal activity of H2O2 by nearly 50 fold [67]. In this context then, Nauseef has suggested the MPO-H2O2-O" pathway to be the center of the neutrophil's oxygen-dependent host defense system [67]. Similarly, the MPO-H2O2-I" system is capable of mediating the oxidation of iodide by forming a reactive iodinating species which can iodinate, and/or oxidize several compounds including proteins and lipids of bacterial membranes [62]. Although this iodide potentiates a bactericidal system, bacterial lolling in vivo is thought to involve chloride rather than iodide. This is primarily because the chloride concentrations present in the cell are considerably greater than that of iodide. Thus, the chloride reaction plays the principle role (in the production of HOC1), while the iodide reaction serves a more secondary facilitating role [62]. The quantification of MPO has consistently been employed as a marker for neutrophil accumulation [14,58,66]. While a direct relationship was not clearly shown, Bury and Pimay [14] showed that exercise in humans at 45%, 60% and 75% V02max 24 concomitantly increased leukocyte count (primarily neutrophils) by up to 2.5 times, and plasma [MPO] by 85% immediately following the exercise bouts. Specifically, downhill running (20% grade) at 60% V02max increased plasma neutrophil count (33%) and [MPO] (97%) immediately following the exercise [18]. Both these plasma parameters, indicative of neutrophil presence returned to baseline 20 minutes post-exercise, suggesting the quick removal of neutrophils from the central bloodstream, owing perhaps to their infiltration into the muscles and/or their adhering onto endothelial cells. Similarly in the rat model, Morozov et al. [66] have shown that strenuous swimming exercise results in an immediate increase (50%) in plasma [MPO], and a more gradual increase in skeletal muscle [MPO] peaking (to 7 times of control) 24 hours post-exercise. These studies provide clear evidence to support significant exercise-induced elevations in circulating neutrophils, and both plasma and skeletal muscle MPO. In an isoproterenol-induced model of cardiac hypertrophy [71], we have shown that cardiac tissue myeloperoxidase activity was elevated by 65%, concurrent with a 31% increase in calpain activity. The activities of both these enzymes were suppressed with prior treatment of the cysteine protease inhibitor, E64c. Collectively, these findings are suggestive of the involvement of the calpain system in neutrophil accumulation in the heart. In addition, our kinetic analysis of MPO activity revealed that Vmax was increased in the: liver > heart > skeletal muscle immediately following exercise; however, the enzyme's Km was significantly decreased (28%) only in skeletal muscle [8]. Therefore suggesting that, with exercise, myeloperoxidase and its associated neutrophils are more readily 25 available in skeletal muscle compared to other metabolically active tissues such as the heart and liver. As outlined above, the assessment of MPO within exercise-induced muscle tissue will allow for the quantification of neutrophil accumulation within the muscle. This will hopefully demonstrate the relationship between the early occurrences of the acute inflammatory response and exercise. Since monocytes/macrophages also possess MPO, early tissue sampling is crucial to minimize possible effects of these inflammatory cells, confounding our index of neutrophil accumulation. Eccentrically based exercise is reportedly most responsible in causing muscle damage, and has therefore also been implicated in eliciting the experiences of DOMS. Since there exists apparent relationships between these events, it is proposed that they will be reflected in the MPO findings of this study. 26 Calpain Calcium-dependent protease (CANP, calcium-activated neutral protease, calpain, EC is a nonlysosomal intracellular cysteine protease that requires calcium ions for activation. Two types of CANP differing in Ca2+ requirements exist ubiquitously in all mammalian tissues examined thus far. They are the low and high Ca2+-requiring types designated: p.-CANP (calpain I) and m-CANP (calpain II) respectively. Consequently, micromolar (pM) concentrations of Ca2+ are required to optimally activate p-CANP, while millimolar (mM) concentrations of Ca2+ are needed for m-CANP activation. [73]. Since fluctuations in intracellular [Ca2+] occur predominantly at submicmolar levels, p-CANP appears to be the more likely isomer functioning under physiological conditions: Both types possess a similar but distinct large 80-kDa subunit and an identical small 30-kDa subunit [73,81]. The large (80-kDa) subunit can be divided into four domains (I-IV) (Figure 2). Domains II and IV are the cysteine protease and Ca2+-binding domains, respectively. The functions of domains I and III are less clear, however the NH2-terminal region of domain I is removed during autocatalytic activation of the enzyme [81]. The small (30-kDa) subunit is involved in the regulation of Ca2+ sensitivity. It contains four potential Ca2+-binding regions in the C-terminus. The NH2-terminus is rich in glycine and possibly interacts with membrane phospholipids [81] to localize or stabilize the enzyme complex. 27 MPTVISPTVAPR p94 m- & Li- Calpain Large Subunit Small Subunit MSEEIITPVYC- / \ //MFLVNSFLKGGG Figure 2. Schematic representation of calpain. The primary amino acid sequence(s) of neutrophil chemotactic regions are shown. As illustrated in figure 2, a novel type of calpain, /i-CANP (p94), has more recently been identified and studied [81,82]. Distinct from both U.-CANP and m-CANP, n-CANP is believed to occur specifically in only skeletal muscle cells [80]. While the existence of a possible regulatory small subunit is still unclear, the well characterized large 94 kDa subunit named p94, has become the focus of a series of muscle tissue-specific calpain studies by Sorirnachi et al. [83]. Although there exists an overall significant amino acid sequence homology between p94 and the other conventional calpains (-50%), p94 contains distinct sequences within domains I, II and III — namely the sub-regions NS, IS1 and IS2 respectively. However, domain II, the enzyme's cysteine protease region, is most conserved compared to the other domains [80]. Like (i- and m-type calpain, since the N-terminal portion of p94 is removed during autocatalytic activation, there is strong 28 suggestion that p94 possesses very similar proteinase abilities [80]. Hence, possessing similar myofibrillar substrates, which upon degradation could result in Z-line streaming/smearing and loss of cytoskeletal structural integrity/assembly [37]. p94 mRNA expression in skeletal muscle is ~ 10-fold larger than the conventional calpains [83]. Despite this transcriptional potential, the p94 protein has to date only been detectable in very small amounts. The p94 protein, largely autocatalytically regulated by its IS2 region, is believed to move along the cytoskeleton to the nuclear membrane, where it subjected to autolysis almost immediately after its translation. A small percentage of p94 is however imported into the nucleus, where it is thought to control levels of short lived regulatory proteins such as transcriptional factors. Particularly, the IS2 region in domain III of n-CANP is thought to be responsible for regulating the extremely rapid and extensible autocatalytic degradation of p94 ~ which in part contributes to its difiicult detection. Proteolytic activity of p94 has, however, been detected in the cytosol at [Ca2+] between 10-1000 nM. Thus, indicating that the Ca2+ sensitivity of p94 could be in the nanomolar range, making it much more sensitive than either m- or U.-CANP. In light of this, p94 is seen as the first example of the calpain family which demonstrates proteolytic activity within the cytosol that is active at physiological (100-800 nM) [Ca2+]. Therefore, if this muscle-specific form of the calpain family has a similar propensity for contractile and cytoskeletal protein degradation [81], its immediate actions resulting in muscle-specific cleavage which produces associated peptide fragments, could act as priming reagents for other adaptational regulatory processes (i.e. degradation and re-synthesis). 29 The intracellular enzyme family of calpain is found in association with cytosolic membrane component structures such as phospholipids, structural proteins, ion transport channels and receptors [42]. As well within the sarcomere, calpain is actively found in close association to the I-bands (desmin) and Z-bands (a-actinin). Consequently, this direct contact of calpain with its cellular environment, predisposes its activity to be under direct influence of the intracellular events. To this end, numerous proteins have been identified as substrates for calpain degradation [73]. Among them, cytoskeletal proteins such as: actin-binding proteins (fodrin or spectrin, talin, fUarnin, a-actinin) and microtubule-associated proteins. As well, membrane proteins such as: growth factor receptors (EGF receptors), adhesion molecules (integrin, cadherin, N-CAM) and ion transporters (Ca2+-ATPase) have also been identified as specific substrates for proteolysis by calpain (see appendix: Saido et al., 1994 [73]). It is believed that activation of calpain through increased intracellular [Ca2+] within muscle, results in the selective proteolysis of these various contractile and/or structural elements. The mechanism which contributes to the activation of calpain and the subsequent degradation of selected myofibrillar, Z-line and cytoskeletal proteins is not known but may involve mechanically- and/or metabolically linked changes in Ca2+ homeostasis. There is some evidence to suggest that exercise can cause a loss of Ca2+ homeostasis in muscle [25,85]. Indeed, disturbances in Ca2+ metabolism have been associated with altered morphology and muscle weakness after prolonged low intensity loading [17]. Although specific injury to the fibres was not assessed, Tate et al. [25] observed elevated mitochondrial [Ca2+] in the muscle of rats exhausted by treadmill exercise. Similarly, Duan et al. [85] observed elevated mitochondrial calcium concentrations in injured soleus and 30 vastus intermedius muscle immediately following downhill walking. In there assessment, both histological injury and the accumulation of Ca2+ in the mitochondria were markedly reduced when animals were treated with EDTA or EGTA during and after the exercise. Muscle fibres injured by eccentric activity show swollen regions with a marked loss of membrane integrity and elevated free cytosolic [Ca2+] [49]. The histological appearance of injured fibres immediately after downhill walking suggests that there may be disruption of the sarcolemma [85]. It is thus conceivable that changes in membrane permeability, sufficient to allow the leakage of large intracellular enzymes, lead to ultrastructural damage via an influx of Ca2+ from extracellular space. Removing Ca2+ from the media in an isolated soleus preparation protects against ultrastructural damage and enzyme efflux after an intense contraction protocol, but it has no protective effect on the contractile properties of muscle [91]. Major changes in the structure and function of the sarcoplasmic reticulum (SR) occur following strenuous exercise [59], and the decrement in force production associated with exhaustion may be due to an interruption in the function of the SR to transport Ca2+ [76]. Vollestad and Sejerstad [96] suggested that the inability of the SR to reaccumulate Ca2+ contributed to a decrease in the degree of actin-myosin interaction, thereby promoting muscle weakening and/or peripheral fatigue. It has also been postulated that conditions of prolonged muscle use lead to Ca2+ overload within the cell [61]. Sembrowich and Gollnick [95] reported that the Ca2+ uptake activity of the SR isolated from rat fast-twitch muscle was depressed following exhaustive treadmill running. Alterations in the rate of Ca2+ uptake by the SR parallelled a diminished activity of the 31 ATPase pump found along the longitudinal portions of the SR. As well, Belcastro et al. [11] reported an 18% reduction in Ca2+-dependent ATPase activity of the SR following prolonged running exercise. Changes in the capacity and rate of Ca2+ uptake and release by purified SR vesicles have also been reported [60]. Ultrastructural deformities of the SR are accompanied by depressed rates of Ca2+ uptake, diminished Ca2+ release and an increase in intracellular free [Ca2+] [59]. Therefore, this failure in calcium regulation by the SR may result in a favorable environment for die initiation of myofilament degradation by calcium-sensitive proteases. Calpastatin is an endogenous inhibitor of calpain and is equally effective in inhibiting all isozymes [37]. It prevents both enzyme activation and the expression of catalytic activity. The calpastatin prepared from mammalian sources can be classified into two types, based on molecular weight. The erythrocyte inhibitor (48 kDa) is smaller than the inhibitor from other tissues (73 kDa). The reactive site of calpastatin shows no apparent homology to the reactive sites of other known protease inhibitors but does contain a consensus (TIPPXYR) sequence essential for inhibition [45]. Immunolocalization studies provide evidence to suggest that calpain and calpastatin are co-localized within the cell [52]. In the presence of Ca2+, calpastatin binds to the large subunit of calpain with high affinity. This inhibitor is a substrate of the protease and its degradation correlates with the general mechanism of regulation. Calpastatin binds to calpain at a [Ca2+] lower than that required to initiate proteolytic activity [44] and the [Ca2+] required to induce proteolytic activity is much higher than the [Ca2+] that normally exists within the cell. Therefore, cells possess some mechanism to reduce the Ca2+ 32 requirement for proteolytic activity of calpain and to regulate inhibition by calpastatin [37]. In addition to [Ca2+], the enzyme-inhibitor interaction is probably regulated by some other complementary mechanism, possibly the phosphorylation of calpastatin [1]. While the physiological function(s) of the calpain system is still unclear, its ubiquitous distribution has led to numerous associations with a large number of different Ca2+-related proteolytic changes in cells [37] (Appendix: Table 4 Goll et al., 1992). In vitro assessments have shown that the calpains cleave at a limited number of specific sites in native proteins, producing large polypeptide fragments rather than degrading the proteins to small peptides and amino acids. While still retaining their enzymatic activity, these polypeptide fragments, following cleavage, are no longer subject to the controls which regulate the intact enzyme [88]. As well, the calpains are known to cleave a limited number of sites in many cytoskeletal proteins. With this occurrence, the resultant polypeptide fragments of calpain degradation looses its ability to re-bind or cross-link with their previously associated proteins. For this reason, the calpain system is believed to play a key role in the disassembly and re-modeling of the cytoskeletal matrix; particularly at the level of its filamentous cytoskeletal attachments to the plasma membranes and/or subcellular structures. Reports of a transient and specific removal of Z-disks from striated muscle has aroused the suggestion that calpain initiates metabolic turnover of myofibrillar proteins by releasing them from their cytoskeletal filament structures [9,37]. However, despite the existence of numerous substrate proteins within muscle, no single characteristic has been identified which predisposes or targets a protein substrate for proteolysis by calpain. 33 There is no direct evidence in the literature to support a relationship between increased calpain activity and muscle injury. However, there is partial support since purified myofibrils from exercised muscle demonstrate a pattern of myofibrillar proteolysis reminiscent of the actions of calpain (i.e. loss of Z-line integrity and/or its associated proteins) [10,36]. Belcastro et al. [10] described the specific loss and/or disruption of the Z-line structure in 22% of myofibrils isolated from exercised skeletal muscle in the rat. They observed an extensive loss of two proteins (58 kDa and 95 kDa) which were thought to be associated with the Z-line, probably desmin and a-actinin. In vitro, activated calpain has been showed to readily degrade desmin, thereby promoting the release of a-actinin [36]. Furthermore, Belcastro [6] showed that immediately following exhaustive running exercise, p.- and m- calpain activity increased by 18% and 22%, respectively. The increase in calpain activity was accompanied by a heightened Ca2+ sensitivity of the enzyme to half maximal activation (lowered pCaso). Moreover, calpain activation was associated with increased susceptibility of its substrate proteins to degradation, evidenced by the varying degradation rates of proteins from the myofibrillar complex, the Z-line and the cytoskeleton. Although calpain activation in skeletal muscle appears to be dependent on the volume of work performed, the intensity of the work does not significanUy affect the activity of the enzyme (unpublished observations). When equated on the basis of caloric output, Belcastro et al. (manuscript submitted) found that calpain is activated to different extents in the soleus, plantaris and vastus during treadmill exercise. It is conceivable that the extent of muscle recruitment and its contribution towards the required movement play a significant role in the activation of calpain. 34 In a model of ventilatory failure induced by resistive loading, Reid et al. [72] observed severe myofibrillar alterations and inflammatory cell infiltration in the diaphragm. Moreover, proteolysis of purified diaphragm muscle digested with calpain demonstrated faster degradation rates of tropomyosin and a-actinin in tracheal-banded hamsters. Increased calpain activation has also been shown in an isoproterenol-induced model of cardiac hypertrophy in rats [3]. The associated increase in calpain activity in the hypertrophic hearts were attenuated when the animals were pre-treated with the cysteine protease inhibitor, E64c. Furthermore, electron micrographs depicting the isoproterenol-induced injury demonstrated extensive damage of sarcomeres in the region of the Z-lines, concurrent with calpain activation. These myofibrillar alterations were not evident in either the control or the E64c pre-treated groups. By inhibiting the activation of calpain in vivo, E64c appears to have some protective effect in maintaining the registry of the muscle. Collectively, these fmdings suggest that muscle is predisposed to the actions of calpain during periods of increased demand. Kunimatsu et al [53,54] demonstrated (dose-dependent) neutrophil chemotactic potential from peptide fragments of calpain (Figure 2). Because we have provided evidence that calpain is activated during exercise and that it selectively degrades muscle proteins, it is our contention that calpain (or the resultant peptide fragments) may be associated with the neutrophil chemotaxis reported to occur during or immediately following exercise. The potential of a calpain-mediated process(es) in stimulating neutrophil movement stems from the neutrophils' ability to recognize a chemotactic gradient and to transform this signal into directional locomotion. While specific surface 35 receptor sites have been identified for C5a and another glycoprotein urate crystal-induced chemotactic factor [92], a number of peptides of bacterial origin, such as N-formylated oligopeptides are also potent activators of neutrophils [48]. Of these, iV-formyl-methionyl-leucyl-phenylalanine (/MLP) is one of the most significant. Formylated peptides such as /MLP bind to specific receptors on the plasma membrane of neutrophils. Depending on the concentration used, /MLP can trigger chemotaxis, aggregation, reactive oxidant production, cytoskeletal alteration and (in the presence of cytochalasin B) degranulation [27]. The biosynthesis of such bacterial proteins as /MLP typically code for formyl-methionine at the NH2 terminal. Due to proteolytic cleavage of Af-formyl-meuhionine however, few mature bacterial proteins possess methionine at the NH2 terminal. Interestingly, all native forms of the calpain family are encoded with methionine as the first amino acid at the NH2 terminal [80]. Using synthetic peptides of various lengths from the large and small subunits of p.- and m-calpain, Sasaki et al. [74] demonstrated dose-dependent neutrophil migration. Presumably, an oligopeptide released from the NH2-terrninal of domain I during autolysis [81] was responsible for the chemotactic activity. While acetylated iV-methionyl peptides from the small calpain subunit demonstrated chemotactic activity, the formylated peptides displayed additional dose-dependent superoxide generation and myeloperoxidase/elastase releasing activity [54]. It is thus conceivable that the calpain peptide fragments bind to a similar neutrophil receptor as /MLP. These observations provide the rationale this study and other investigations directed at understanding the interaction of calpain-mediated processes (through formation of various //-formylated oligopeptides) to stimulate neutrophil accumulation into muscle 36 following exercise-induced muscle injury. Therefore, in light of its numerous responses to exercise induced muscle damage, it is clear how calpain can be involved in this study. 37 RESEARCH METHODS Animal Care and Training Male rats (-325 g) were housed 2-4 per cage in a room maintained at 23 ± 1°C. Days were divided into 12 hours each of light and darkness. Food (commercial rat chow) and water was provided ad libitum. All animals were familiarized on a motorized h-eadmill 10-15 min/day for 3 days prior to experimental treatments. During these sessions, the running speeds were progressively increased to a maximum of ~25 m/min and maintained for the final 2 min. At the end of this familiarization period, the rats were not considered "exercised trained" in a physiological sense [56,57] but rather, "accustomed" to the exercise setting. Approval for this study was granted by the Animal Care Committee at the University of British Columbia (reference number: A93-1437). All the animal experiments were conducted according to the guidelines of the Canada Council on Animal Care (1980-1984). 38 Exercise Protocol — Chemotaxis Following the initial exercise familiarization period, Sprague-Dawley rats (~325 g) were randomly assigned to either a control (n = 10) or an exercise (n = 10) group. The experimental treatment constituted a -14% grade downhill run, at 25 nVmin on a motorized treadmill for 1 hour or until voluntary termination. Noise and/or air was used to motivate the animals. Prior to sacrifice, the sedentary control group were exposed to the same housing conditions as the running animals. Muscle Preparation Immediately following exercise, the rats were sacrificed with a lethal injection of sodium pentobarbitol i.p. (1 ml Euthanol/kg body weight). The heart was then quickly excised followed by the skeletal hind-limb muscles of the soleus (slow-twitch) and plantaris (fast-twitch) . All tissue samples were excised in <7 min to mimrnize post mortem artifact (i.e. rigor mortis associated metabolic changes, proteolytic changes). Upon excision, the muscle samples were immediately suspended individually in vials containing 5 ml of phosphate-buffered saline (PBS) and shaken (Fisher Vortex Genie 2-model G-560: shake setting of 2) on ice for 2 hours (Figure 3). This process yielded the membrane soluble fraction and was centrifuged at 400 x g (1600 rpm; Hermle-model Z 233-M) for 10 min at 4°C to remove any cellular debris or red blood cells (RBC). The cytosolic fraction was processed by homogenizing the muscle sample in 5 ml PBS with 2 x 39 10 sec bursts on a tissue homogenizer (Ultra-Turrax: setting 6.5). The resulting muscle suspension was then centrifuged at 22,000 x g (13,000 rpm; Beckman ultracentrifuge-model L7-65: rotor SW28) for 20 min at 4°C, and the supernatant collected as the cytosolic fraction. All samples, the membrane soluble and the cytosolic fractions were processed fresh and used on the day of experimentation. These samples were used to test the first objective, which assessed the possibility that a myogenically produced chemotactic factor could attract neutrophils to the muscle. PBS \_ muscle shake centrifuge -t Soluble I'"*-re-suspend muscle w&j rf Cytosolic fraction pellet centrifuge _ PBS ' tissue homogenize • muscle muscle suspension Figure 3. Schematic representation of muscle tissue processing. Chemotaxis Research Design Neutrophils were isolated and purified from blood samples from a separate group of control Sprague-Dawley rats (-325 g). This served as the neutrophil pool from which 40 chemotactic activity was assessed for control and exercised tissues from the two muscle fractions of the heart, soleus and plantaris. Neutrophil chemotaxis was assayed on all muscle fractions undiluted (muscle suspended in 5 ml PBS), and in serial dilutions of 1/10, 1/100 and 1/1000 in PBS (Table 1). Negative control Hank's balanced salt solution (HBSS) and PBS Positive control fMLP at 10"8M and 10"7M Chemoattractants Muscle suspensions [-0.5-1.0 mg/ml] were serially diluted with PBS to concentrations of 1/10, 1/100 and 1/1000 Table 1. Control vs exercise muscle tissue neutrophil chemotaxis -- done in triplicate For the purposes of this study, the cytosolic fraction represents chemotactic potential housed within the boundaries of the cell; not bound by any membranous stmctures (i.e. organelles) in the cell, but rather resident "free" within the cytoplasm. This pool therefore serves as an extended reservoir of proteins which could potentially leak out of the cell. Conversely, the membrane soluble fraction represents the chemotactic portion which would have leaked out of the cell during extraction (i.e. shaking). This fraction then represents the fraction that would have in a physiological sense, leaked out of the muscle and into the circulatory mainstream/interstitium. Neutrophil chemotaxis was found to be negligible or undetectable in muscle fractions diluted 1/10 (<3 cells/high-powered field), 1/100 (<1 cell/high-powered field) and 1/1000 (<1 cell/high-powered field). 41 Therefore, all the results concerning chemotaxis will represent exclusively, the undiluted muscle fractions (i.e. muscle in 5 ml PBS). Isolation of Neutrophils The purification of neutrophils was based on a modified version of a method by Walker, et al [99]. Whole blood (-15 ml), anticoagulated with heparin (Hepalean 1,000 U.S.P. units/ml), was collected from untreated Sprague-Dawley rats by a cardiac puncture while under halothane anesthesia. The anticoagulant-treated blood was then diluted 1:1 with Hepes normal saline (H-NS) to yield 30 ml of diluted blood. This was then carefully layered 2:1 onto 15 ml Ficoll-hypaque 1077 in a 50-ml polypropylene centrifuge tube, and centrifuged at 400 x g (1400 rpm; Beckman rotor TJ-6 / 1600 rpm; Hermle-model Z 233-M) for 30 min at room temperature. Following centrifugation, the plasma, mononuclear cell and platelet interface, and the Ficoll layer were aspirated off ~ leaving the red blood cell (RBC) layer which contained the polymorphonuclear cells (PMN). The centrifuge tubes were then filled with NH4CI lysing solution to resuspend the RBC layer. They were allowed to sit until the suspension turned from a matte red to glossy black; indicating RBC lysis. The tubes were centrifuged at 400 x g for 7 min at 4°C. The supernatant was then aspirated down to the PMN pellet and the lysing process repeated once more. After this second lysing step, the PMN pellet was washed with H-NS and centrifuged at 500 x g (1900 rpm) for 6 min at 4°C. The neutrophils were then resuspended at 1 million cells/ml 42 in H-NS for temporary storage (< 2 hours), and finally in Hanks balanced salt solution (HBSS) just prior to use. The preparation at this point typically contained >75 ± 5% neutrophils, <15 ± 5% monocytes and <5 ± 2% of others cells (e.g. eosinophils, lymphocytes). In addition, cell viability was consistentiy >95 ± 2% with trypan blue exclusion. The neutrophil yield from 20 ml blood is about 5 x 105 cellsAnl; therefore approximating 10 x 106 neutrophils in total. Chemotaxis Protocol Neutrophil chemotaxis was performed on a 48-well modified Boyden chemotaxis chamber (NeuroProbe Inc.: Cabin John, MD) through a 3.0 pm pore sized (polyvinyl pyrrolydone-free) polycarbonate filter membrane (Figure 4). The final loading concentration was comprised of neutrophils suspended at 1 x 106 cells/ml in HBSS. Therefore, a total volume of >2.5 ml of cell suspension was needed to load all 48 wells. Figure 4. Schematic representation of modified 48-well Boyden chamber set-up. 43 The bottom (agonist-loaded) plate was orientated such that the trademark logo faced the upper left corner. The wells were then carefully loaded with 27 ui of either blanks (i.e. HBSS, PBS), standards (i.e. 10"8M/MLP, 10"7M/MLP) or muscle suspensions in serial dilutions. Next, a small cut was made on the upper left corner of the membrane for orientation purposes. With forceps, the membrane was then aligned over the bottom plate to ensure it covered all the wells just before it was laid down. With the membrane bowed in the middle, it was carefully placed onto the plate. The gasket was then placed on top of the membrane — again ensuring correct orientation with cut corner to the upper left. The top plate was fitted on next; orientated such that the trade mark was at the upper left and smooth side facing up, then screwed on finger-tight with the thumb nuts. This upper plate was loaded with 48 u.1 of the neutrophil cell suspension. With all loading, extreme care was taken to ensure that no air bubbles got lodged in the wells, since this greatly impedes the surface contact area between the lower well, the membrane and the upper well. When loading in the upper plate was complete, the chemotaxis chamber was placed in a 37°C humidified (5% C02) incubator for 30 minutes. After incubation, the apparatus was carefully disassembled and the membrane peeled off with forceps and clamps. At this point, care was taken to ensure the locale of the "cell-side" (migrated side) which was to be fixed, and the "non-cell-side" which was wiped off. The non-cell-side (unmigrated side) of the membrane was then wet in 0.9% saline and wiped off over a wiper blade. This wiping process was repeated twice more. When done, the membrane was immersed in methanol, then left to air dry. Diff-Quik solutions I (5 dips) and II (10 dips) were then used to stain the membrane filter. Once 44 appropriately stained and dried, the membrane was mounted on a slide with Permount and stored for counting. Cell quantification was based on the mean count of 4 high-powered fields from each of the 48 stained wells on the membrane viewed at 10 x 40 magnification. All samples counts were normalized to maximal chemotaxis towards the positive control, 10"7M /MLP, and these normalized values are reported as a percentage of maximal chemotaxis. * For further details and illustrations see appendix Exercise Protocol — Correlation Following an initial exercise familiarization period, male Wistar rats (~325 g) were randomly assigned to either a control (n = 5), exercise (n = 5) or (2S,3S)-trans-epoxysucdnyl-L-leucylamido-3-methylbutane (E64c) pre-treated (n = 5) group. Animals in the E64c pre-treated group were subcutaneously injected with the cysteine protease inhibitor (1 mg/kg) 1 hour before running. The exercise test was accomplished with an 8% grade treadmill run at a speed of 25 rn/min for 60 minutes or until voluntary termination — which was reflected by the animals' inability to right themselves. Once again, noise and air were used to motivate the animals. Immediately after running, the rats were administered a lethal injection, i.p., of sodium pentobarbitol (1 ml Euthanol/kg body weight). The cardiac ventricles and plantaris muscles were dissected out and quickly trimmed of visible fat and connective tissue, then frozen in liquid nitrogen with pre-cooled tongs. All samples were then stored at -75°C for subsequent analysis. 45 Calpain Assay Extraction and determination of calcium-dependent, calpastatin-inhibited proteolytic (calpain-like) activity was accomplished on all tissue samples [3,103]. Briefly, tissue samples (approximately 100 mg) were suspended with an Ultra-Turrax homogenizer (IKA Laboratories-model TR-10) for 20 sec at a setting of 6.5, in 10-15 vols of buffer containing 80 mM KC1, 20 mM Tris (pH 7.5), 5 mM EGTA and 2 mM DTT. The suspension was centrifuged (4°C) at 22,000 x g for 15 minutes (Hermle-model Z 233-M) and the supernatant (soluble fraction) decanted and stored in polypropylene tubes on ice (20 min) for subsequent assay of calpain-like activity. Following centrifugation, this particulate material was homogenized with a 2-ml Wheaton glass-homogenizer (10 strokes) in 10-15 vols of a similar buffer (as above) with the addition of 0.35% Triton-X 100 and re-centrifuged under the same previous settings. This procedure was judged to be adequate in removing the 80 kDa (possibly the large subunit of calpain) band from the first pellet, because the second pellet (following glass homogenization with Triton-X 100) contained negligible (if any) amounts of this protein as assessed by PAGE and immunoblotting [6]. Moreover, Tan et. al. [89] have reported that calpain activity is not altered as a result of Triton-X 100 treatment. The supernatant from the second centrifugation step (particulate fraction) was stored on ice and assayed for calpain-like activity. The total calcium-dependent proteolytic activity of the samples were determined by a microplate assay using casein as the substrate [100]. Briefly, 200 |il of extract were 46 added to a reaction mixture containing: 2 mg/ml casein, 50 mM Tris (pH 7.5) and 20 mM DTT (in duplicate). Following a 5 minute pre-incubation at 30°C, 5 mM total calcium (800 pmol/L free calcium as determined by IONS software program) was added to one of the duplicates while the other contained 5 mM EGTA. After 30 minutes at 30°C, an aliquot (100 pi) of each sample was assayed for calcium proteolysis in a total volume of 325 pi using a Bio-Rad protein dye reagent concentrate (Bio-Rad Laboratories). The protein dye reagent is composed of 0.05% (w/v) Coomassie brilliant blue G-250, 23.5% (w/v) ethanol and 42.5% (w/v) phosphoric acid. The enzyme activities (carried out in the absence and presence of calcium) are expressed as caseinolytic activity and calculated based on a 0.1 absorbance change at an optical density of 595 nm being equivalent to 1 unit of enzyme activity. The calcium-dependent, caseinolytic activities of the soluble and particulate fractions are expressed as calpain-like activities, because minimal activity (<5%) was observed when calpastatin (a calpain-inhibitor) was added to the assay. Typical caseinolytic activities for cardiac preparations (i.e. total calpain-like activity) were calculated as the sum of the soluble and particulate fractions. Similarly, the values for total calpain-like activity of rat skeletal muscle determined in this study are comparable to those reported for partially purified calpain preparations [6]. 47 Myeloperoxidase Assay Samples were prepared for the assessment of myeloperoxidase (MPO) activity by homogenization in 10 vol (v/w) of a 50 mM potassium phosphate (pH 7.4) and 0.5% hexadeytrimylammonium bromide (HTAB) solution with an Ultra-Turrax homogenizer (IKA Laboratories-model TR-10) for 2 x 15 sec bursts. Following centrifugation (Hermle HM-22059V05 Rotor, Germany) at 850 x g for 12 rninutes (4°C), samples were frozen and thawed in liquid nitrogen (x2) and re-centrifuged. This procedure was found to yield supernatant fractions with minimal levels of hemoglobin [8]. Quantification of myeloperoxidase activity was performed as previously described [78]. Briefly, 450 u.1 of supernatant was added to a reaction mixture containing: 0.8 mM hydrogen peroxide (H2O2), 10 mM potassium phosphate (KH2PO4) (pH 6.0) and 0.4 mM o-dianisidine dihydrochloride (o-DMB). The reaction was followed continuously (150 sec) in a spectrophotometer (Shimadzu UV-160, Japan) set at 480 nm and 37°C. The absorbances at 480 nm over reaction time (5 sec intervals) were then curve-fitted onto a first order rate constant (GraFit version 3.0, Erithacus Software) to calculate rnaximal activity (see appendix). FL^-dependent MPO activity (with o-dianisidine dihydrochloride as the substrate) was expressed in U/g wet weight of tissue, where 1 unit of MPO activity was defined as a 0.1 absorbance change at an optical density of 480 nm. 48 STATISTICAL ANALYSES Sample size was estimated from results of the concentric study mentioned above, where an n = 5 per cell achieved statistical significance on a 1-way factorial design ANOVA for randomized groups and a Tukey's post-hoc test. Using the previous results as a guideline and an approximation of the means and standard deviation for this study at large, Cohen's d was estimated to be >1.40 for a 1-tailed test at p < 0.05. With that, the same sample size of n = 5 per cell for a 2-way ANOVA was found to still accommodate for an approximated power of >0.80 to be achieved [23]. Statistical comparisons (SPSS for Windows; version 6.1) for chemotaxis study was based on two (one for each muscle fraction) 2-way factorial design for randomized groups analysis of variance (2-way ANOVA). It assessed two 2 (Condition) x 3 (Muscle-type) ANOVA designs, with the level of significance set at p < 0.05. Where analysis of variance proved significant, differences across conditions were evaluated with by a Tukey's post hoc test for honestly significant differences. With a total of 60 muscle samples (2 x 30) from 10 animals, there was an n = 5 per cell for the analyses. Table 2 illustrates the ANOVA table: Soleus Plantaris Cardiac Control n = 5 n = 5 n = 5 Exercise n = 5 n = 5 n = 5 Table 2. Summary of 2 (Condition) x 3 (Muscle-type) 2-way ANOVA. 49 Correlation Analysis All results for calpain-like and myeloperoxidase activities from control and exercise samples were subjected to Pearson's correlational analysis. Correlation coefficients were calculated from the collective data-set by assessing the results for all muscle types and conditions. In addition, statistical comparisons (alpha levels) of r values were determined for the muscles when grouped individually (i.e. cardiac and plantaris) during exercise and control. Differences between control and exercise levels of plasma creatine kinase was assessed with a Student's t-test. With calpain and myeloperoxidase activities, differences between control, exercise and E64c pre-treatment were assessed with a one-way ANOVA and Tukey's post hoc test for honestly significant differences. All analyses (SPSS for Windows version 6.1) were conducted with the level of significance set at p < 0.05. 50 RESULTS Neutrophil Chemotaxis Baseline values for neutrophil chemotaxis were not different between the membrane soluble fractions from muscles of the heart, soleus and plantaris (Figure 6). This finding suggests that, at rest, there appears to be a small but consistent neutrophil response from chemotactic factors released by muscles across the fiber types. In the cytosolic fraction however, resting chemotactic potential was more diverse. The heart demonstrated the greatest chemotaxis response; a response larger than the soleus, and significantly greater than the plantaris. Although varying between muscle types, these results support the contention that, at rest, muscles possess storage capacity for neutrophil chemotactic agents within its cytoplasm. All rats in the exercise group ran for 56 ± 3 minutes (mean ± S.D.). In general, regardless of fiber type, exercise results in a reduced chemotactic capacity of the cytosolic fraction in muscle (Figure 5). On the other hand, proportionally similar increases in the neutrophil chemotactic potential are reflected in the soluble fractions of exercised muscle extracts from the soleus and the plantaris muscles. However, this increased chemotactic finding post-exercise was not observed from the soluble fraction of the heart. Neutrophil chemotaxis for control soleus muscle (31.79 ± 14.41 % max chemotaxis) decreased by 71% immediately after exercise (9.26 ± 4.63 % max chemotaxis) (p < 0.05). Likewise, control plantaris muscle (16.16 ± 4.68 % max chemotaxis) reflected similarly a 69% 51 decrease in neutrophil chemotaxis following exercise (4.97 ± 2.49 % max chemotaxis) (p < 0.05). The greatest decline in neutrophil chemotaxis was observed in cardiac muscle, where chemotaxis values from control cardiac muscle (47.14 ± 13.69 % max chemotaxis) declined significantly by 77% with exercise (10.92 ± 4.74 % max chemotaxis) (p < 0.05). These results clearly indicate that exercise has a suppressive effect on the chemotactic potential in all muscle types. Figure 5. Bar-plot representing neutrophil chemotactic potential in the cytosolic fraction of muscle extracts. Neutrophil chemotaxis as normalized and plotted to a percentage of maximum (100%) of 10"7M /MLP. Results are presented as means ± S.D.; * denotes significant difference from control at p < 0.05. Neutrophil chemotaxis from the membrane soluble fraction demonstrated a markedly different response to exercise (Figure 6). In the soleus, values for neutrophil 52 chemotaxis increased by 81% from control (9.03 ± 3.03 % max chemotaxis) to exercise (46.70 ± 1.91 % max chemotaxis) (p < 0.05). Albeit to a lesser extent, control plantaris (9.31 + 1.25 % max chemotaxis) reflected a similar increase of 58% with exercise (22.31 ± 4.06 % max chemotaxis) (p < 0.05). Neutrophil chemotaxis in cardiac muscle however, was not significandy different between control (7.74 ± 2.73 % max chemotaxis) and exercise (4.28 ± 1.83 % max chemotaxis) (p > 0.05). I | Control H Exercise Soleus Plantaris Heart Figure 6. Bar-plot representing neutrophil chemotactic potential in the soluble fraction of muscle extracts. Neutrophil chemotaxis as normalized and plotted to a percentage of maximum (100%) of 10*7M fMLP. Results are presented as means ± S.D.; * denotes significant difference from control at p < 0.05. 53 Relationship: Calpain and Myeloperoxidase At rest, baseline calpain-like activity and myeloperoxidase activity were significantly higher in the heart compared to the plantaris (p < 0.05) (Figure 10). Rats in the exercise group for this study ran for 58 ± 6 minutes (mean ± S.D.) and had increased levels of plasma creatine kinase of 211 ±41 U/L compared to 89 ± 20 U/L (control samples) (p < 0.05). Increases in total calpain-like activity were observed for both skeletal and cardiac muscle immediately after exercise. The activity for control plantaris muscle (10.78 ± 7.36 U/g wet wt.) was increased by 68% after exercise (33.82 ± 15.82 U/g wet wt.) (p < 0.05). Exercise resulted in a 31% increment in the response of cardiac muscle (39.29 ± 11.63 U/g wet wt.) when compared to control (26.93 ± 1.80 U/g wet wt.) (p < 0.05). Increases in myeloperoxidase activity were also seen for both the plantaris and cardiac muscles. Plantaris muscle from exercised animals (1.89 + 1.15 U/g wet wt.) demonstrated a 93% increase (p < 0.05) in MPO activity over control muscles (0.14 ± 0.10 U/g wet wt.). While absolute levels were greater for cardiac muscle compared to plantaris, a smaller albeit significant increase of 51% was obtained for cardiac muscles with exercise (control = 1.56 ± 0.30 vs exercise = 3.18 ± 1.22 U/g wet wt.) (p < 0.05). 54 3 CARDIAC AND PLANTARIS MUSCLE 60 40 h-20 r = 0.8279 O Control Heart # Exercise Heart • Control Plantaris • Exercise Plantaris! J I L J I L 0 2 4 6 Myeloperoxidase Activity (U/g wet wt. tissue) Figure 7. Pearson's correlational analysis (SPSS for Windows; version 6.1) illustrating the: Relationship between myeloperoxidase and calpain-like activity of the combined data-set for both plantaris and cardiac muscles for both resting and exercised conditions. Based on individual comparisons, where exercise consisted of a 25 m/min treadmill run on an 8% grade for 60 min. The relationship between individual comparisons for calpain-like and myeloperoxidase activities for all muscles (Figure 7) at rest and immediately following exercise was found to be significant; with an r = 0.8279 (p < 0.001). From this, the data-set was analyzed separately for the different muscle types. Plantaris muscles (Figure 8) following exercise and at control were correlated for MPO and CANP at an r = 0.6822 (p < 0.05), while similar conditions for cardiac muscles (Figure 9) correlated much higher at an r = 0.9708 (p < 0.001). 55 ~ 60 :s f 40 20 PLANTARIS MUSCLE r = 0.6822 o CONTROL • EXERCISE J I J I I I I I I I—I— 0 2 4 6 Myeloperoxidase Activity (U/g wet wt. tissue) Figure 8. Pearson's correlational analysis (SPSS for Windows; version 6.1) illustrating the: Relationship between myeloperoxidase and calpain-like activity of plantaris muscle for both resting and exercise conditions. Based on individual comparisons, where exercise consisted of a 25 m/min treadmill run on an 8% grade for 60 min. 56 CO 3 Vi .53 I U CARDIAC MUSCLE 3 60 h 40 h-20 h-r = 0.9708 o CONTROL • EXERCISE J I I I L J I I I L 0 2 4 6 Myeloperoxidase Activity (U/g wet wt. tissue) Figure 9. Pearson's correlational analysis (SPSS for Windows; version 6.1) illustrating the: Relationship between myeloperoxidase and calpain-like activity of cardiac muscle for both resting and exercise conditions. Based on individual comparisons, where exercise consisted of a 25 m/min treadmill run on an 8% grade for 60 min. To implicate the actions of calpain, a corresponding inhibition in the enzyme's activity should be evident when it is blocked. The pre-administration of the cell permeable cysteine protease inhibitor, E64c, resulted in a suppression in calpain-like activity when compared to the normal running group (Figure 10) (p < 0.05). E64c decreased (p < 0.05) calpain-like activity by 56% in cardiac muscle (17.73 + 2.46 U/g wet wt.), and by 76% in the plantaris (8.02 ± 2.05 U/g wet wt.). Interestingly, E64c also suppressed (p < 0.05) MPO activity by 64% in cardiac muscle (1.16 ± 0.36 U/g wet wt.), and by 48% in the plantaris (0.98 ± 0.22 U/g wet wt.). In both muscles, calpain-like and MPO activities of the E64c pre-treated group were not different compared to control levels (p > 0.05). 57 These findings clearly support the involvement of calpain with exercise. In addition, calpain's role in supporting neutrophil accumulation, as evidenced through tissue myeloperoxidase activity, is further supported by the corresponding decreased response of MPO with calpain inhibition. H Control f~J Exercise H E64c Plantaris Heart Plantaris Figure 10. Bi-axial bar plot illustrating the effects of E64c pre-treatment on myeloperoxidase and calpain activities following treadmill running. Values are means • S.D.; * denotes significant difference from control at p < 0.05. Exercise consisted of a 25 m/min treadmill run on an 8% grade for 60 min. Runners in the E64c group were pre-injected 1 hour prior to the running protocol. 58 DISCUSSION The primary observation in this study was the finding of a muscle-generated (myogenic) neutrophil chemoattactant(s). Results from neutrophil chemotaxis assays on striated muscle extracts support the hypothesis that, exercise-induced injury to muscle stimulates the release of a physiologically viable neutrophil chemotactic agent(s). Exercise brought about a significant decline in the free cytosolic chemotactic potential, and a concomitant increase in the releasable membrane soluble fraction of skeletal muscles. Attempts to identify the agent(s) responsible for this chemotactic link was supported by a significant relationship between the activities of the calcium-stimulated neutral protease calpain, and the neutrophil marker enzyme, myeloperoxidase. These results suggest that the underlying processes regulating calpain's action and neutrophil accumulation are closely linked. Across all tissues a positive relationship (r = 0.83) was estimated between striated muscle protease activation and neutrophil accumulation, as evidenced by myeloperoxidase activity, with a notably greater relationship for cardiac muscles compared to fast type skeletal muscles. While pre-exercise administration of E64c resulted in a predictable reduction in calpain-like activity, it also attenuated exercise-induced myeloperoxidase activity. Because myeloperoxidase activity is not directly affected by E64c (unpublished observations), we conclude that neutrophil accumulation into rat striated muscle is at least partially dependent upon the activation of cellular calcium-stimulated cysteine proteases, such as calpain. 59 In trying^ to understand the mechanics governing the release of the soluble neutrophil chemoattractant(s), focus is directed to chemotactic components found free within the cytoplasm or released from it. Across all muscles, resting chemotaxis values from the membrane soluble fraction were not different, even though resting values from the cytosolic fraction were suggestive of ranging chemotactic potential for different muscles. As such, the release of neutrophil chemoattractant(s) does not appear to be driven by a concentration gradient, since resting membrane soluble values for the heart were similar to those of skeletal muscles. Likewise, a similar suggestion can be made with regard to the potential for peptide "leak" and pore size of the plasma membrane, both of which appear to be consistent at rest across muscle types. Support for a regulated process is evidenced by muscle's capacity to release peptide fragments like myosin heavy chain (MHC) [60,61] and the troponins [59], and metabolic by-product enzymes such as creatine kinase (CK) [50,51] and lactate dehydrogenase (LDH) [38] when metabolically challenged. Such regulated processes are believed to coincide with accompanying changes in pH, core temperature and electrolyte shifts that effect metabolic turnover [17,97]. While its now clear that injured striated muscle has the potential to trigger neutrophil chemotaxis, the exact identity however, of this neutrophil chemoattract(s) remains to be determined. Since a variety of agents are potentially released from the muscle following injury, numerous possibilities exist with such injury models as exercise. Possible chemotactic candidates include MHC [60,61] and the troponins [59], which are known releasable peptides fragments from the disruption of the myofibrilar apparatus. Likewise, other releasable clinical indicators such as CK [50,51], and LDH [38,46] could also be 60 implicated in neutrophil chemotaxis, along with such metabolic by-products as the adenalytes [38,77] which are produced during the cycling of ATP, since these too can be liberated by the muscle. Cytokines are also likely candidates. The pro-inflammatory cytokines IL-1, IL-6 and TNF have previously be implicated in the progression of the acute inflammatory response to exercise [20]. However, it has more recendy been shown that except for EL-6 which acts mainly in B-cells, both IL-1 and TNF are noticeably suppressed immediately after strenuous exercise [25,85,95]. Since members of the monocyte chemoattractant protein (MCP) [39,84] and macrophage inflammatory protein (MLP) [35,49] family of chemokines primarily target monocyte/macrophage chemotaxis, their involvement in muscle-generated neutrophil chemotaxis can only be minimal at best. In the rat model, the human EL-8 analogue CINC, is still to date a good candidate [41,76,91] since (i) it can be generated from resident macrophages within the muscle, (ii) it is a small (8 kDa) and readily releasable peptide, and (iii) it is extremely chemotactic to the neutrophil. While it is beyond the scope of this study to exclude the numerous possible chemotactic candidates, this study has demonstrated that striated muscle releases a chemotactic agent(s). Further studies are however warranted to identify the precise agent(s). The findings from the soluble fraction offer the first direct evidence for a myogenic neutrophil chemoattract released following exercise. While more pronounced in the soleus than the plantaris, both striated muscles representing the predominandy slow and fast muscle fiber types respectively, demonstrated exercise-induced increases in neutrophil chemotaxis. Clearly, these findings are suggestive of a free intracellular pool of 61 chemoattracts, which are at rest contained within the boundaries of the cell, but can be easily released with exercise. In this regard, the proposed mechanism (Figure 1) out-lining the sequence of events following exercise induced muscle damage still holds; at least for skeletal muscle. Why these increases were not reflected in the soluble fraction of the heart is open to conjecture. Although there exists chemotactic potential in the heart as evidenced by the data from the control cytosolic fraction, these resident chemoattracts within the cytoplasm are non-existent following a bout of eccentric exercise. Perhaps the changes in the heart occurred too quickly and/or the techniques used were too crude to detect the change. Although it is tempting to speculate that the heart has a protective mechanism which guards itself against excessive neutrophil and/or inflammatory cell invasion, this is probably not the case. Blood flow dynamics may offer a plausible explanation. Since exercise does bring about increases in cardiac output and its associated increases in blood flow [21] and perfusion pressures [55], it is not inconceivable that the heart, served by its coronary circulation could have released its chemoattracts before the time of sampling. Hemodynamics and muscle recruitment patterns complementing the metabolic dynamics in skeletal muscle [43,75], could also play a contributing role in the release of chemoattractants. Compared to the fast-twitched (glycolytic) plantaris, being a slow-twitched (oxidative) muscle, the soleus would be expected to be more readily innervated and therefore more consistently perfused with blood. Adequate blood flow then, to the used muscles could easily facilitate the release of myogenic chemoattractants into the bloodstream. Alternatively, hypoperfusion (comparatively) in the plantaris which only gets sufficiently perfused upon innervation and muscle recruitment, would be 62 expected to have a delayed neutrophil chemotactic response. The exact identity of the chemoattractant(s) remains unclear, and it still could possibly be any one of the earlier mentioned cytokines (i.e. CINC, MIP-2 etc.). However, what is clear is that this chemoattractant(s) is produced in muscle during exercise and it is released from the muscle immediately thereafter. A depressed chemotactic response from the cytoplasm could suggest loss of the chemotactic agents either to the blood stream, or to being "bound" by membraneous structures within the cell. Neutrophil chemotaxis could also be masked by an anti-chemotactic agent, perhaps produced as a by-product of exercise. With decreases in the chemotactic potential, as evidenced with the muscle cytosolic fractions, care should be taken in interpreting the nature of the apparent chemotactic factor suppression. The decrease could be a function of one, or any combination of events: (i) a decrease in the amount/concentration of the chemotactic factor present, (ii) an increase in the amount/concentration of an anti-chemotactic agent which could feasibly inhibit chemotaxis, (iii) a suppressive effect of a possible cytotoxic environment (enzyme release) on the chemotactic factors, or (iv) an inhibitory effect of the phagocytic environment on the neutrophils per se [13]. With exercise however, numerous other factors need to be taken into consideration. Exercise-induced metabolic by-product release, and resultant electrolyte and fluid shifts could confound and/or block the chemotactic response by themselves being anti-chemotactic, or perhaps have an indirect influence on neutrophil chemotaxis. For example, metabolite increases with exercise associated lactate production consequently reduces muscle pH [17]. Further, imbalances in high-energy adenosine 63 related adenalytes (i.e. ATP/ADP + P; ratios) could affect ATP phosphorylation potential [38], ATPase regulated channels and other ATP regulated processes [17], thereby compromising muscle function and control mechanisms. Therefore, when attempting to identify possible modulators of neutrophil chemotaxis to muscle, consideration should given to these likely confounding influences. Changes in the [Ca2+]i is now known to play an integral role in the regulation of muscle. Alterations in the amount of Ca2+ release and the rate of Ca2+ sequestration by the sarcoplasmic reticulum (SR), have been implicated in the decreased levels of actin-myosin interaction, thereby promoting overall muscle fatigue through compromised functioning of the myofibrillar apparatus [15,97]. While depressed rates in Ca2+ uptake have been found to parallel altered activity of the ATPase pump within the SR, these changes are transient, recovering back to normal in an hour [17]. Impaired Ca2+ uptake with repeated cycling of the cross-bridges, as seen with exercise, dictate the amount of available Ca2+ in the SR for release, and consequently results in a generalized intracellular increase in Ca2+ exposure to the myofibrils. The consequences of increased [Ca2+]j are not completely understood, however some of the proposed outcomes include: (i) activation of proteases and phospholipases, (ii) decreased mitochondrial energy production capacity, and (iii) apoptosis [93]. Depressed rates of Ca2+ sequestration by the SR during fatigue differentially affect muscle recruitment during exercise; where slow muscle is most drastically affected while fast muscle types tend to retain normal rates of Ca2+ uptake [16]. This phenomenon may in part contribute to the greater neutrophil chemotactic response, 64 higher calpain and myeloperoxidase activities found comparatively in slow versus fast twitched muscles. The Ca2+ activated neutral protease, calpain is thought to be largely responsible for the proteolysis of the myofibrillar complex. Increases in Ca2+ stimulated muscle proteolysis were direcdy inhibited by cysteine proteases leupeptin and E-64 but not lysosomotropic agents, thereby barring the involvement of non-lysosomal proteases in Ca2+ dependent proteolysis in muscle [5,105]. Calpain activation is associated with specific peptide cleavage which can alters the regulation of various enzymes (e.g. protein kinase C, myosin light-chain kinase, phosphorylase kinase), and also remodel and/or disassemble the cytoskeleton of the cell (e.g. desmin, nebulin, titin) [37]. This limited proteolytic capacity of calpain will likely cause destabilization and the loss smictural integrity with the cell; predisposing substrate proteins to further degradation [73]. To this end, calpain has be regarded more as a regulatory enzyme rather than an imctiscriminating degradative protease. The activation of calpain has also been implicated in the degradation of the muscle's plasmalemma [64,104]. This occurrence could feasibly allow a passage for some of the smaller degraded cytoskeletal elements to "leak" into the circulatory bloodstream. Intracellular free Ca2+ also plays an important role in stimulus-induced neutrophil activity. Since neutrophil activation negates prior chemotactic movement of the neutrophil to the required locus, the involvement of Ca2+ in this regard seems warranted. Elevations in the [Ca2+]i in vitro with small amounts of Ca2+ ionophores, such as ionomycin, increases the release of O2in neutrophil preparations [31]. This requirement 65 for calcium is also consistent with neutrophil preparations stimulated with /MLP, where peak [Ca2+]j levels and the rate of increase in [Ca2+]; correspond to increases in O2 production ~ increases furthermore, which can be inhibited with prior Ca2+ chelator incubation [32]. Indeed, it appears that increases in [Ca2+]; and the resultant activation of calcium dependent proteases (i.e. protein breakdown) accompany/compliment the increases in neutrophil chemotactic potential. This suggestion was further evaluated in this study by assessing the in vivo relationship between activities of the proteolytic enzyme calpain and neutrophil myeloperoxidase. The finding that general increases in striated muscle protease activation and neutrophil accumulation accompany periods of increased contractile activity, such as prolonged running exercise, agrees with previous observations [6,8,70,79]. Because this study employed individual comparisons (rather than group means) across both cardiac and fast type muscles of calpain-like protease activity and myeloperoxidase activity, a novel finding of this study was that the changes in these systems were positively related within a single sample. Although this positive relationship was more pronounced for cardiac samples compared to fast type samples, skeletal muscles appear to respond in a similar fashion to an exercise load. The factor(s) contributing to the higher correlation co efficient for cardiac samples is unclear, but it may be related to the greater oxidative capacity and supporting blood flow required for cardiac muscle fibers. It has been suggested that the relatively lower blood flow rates associated with fast type muscles may impede the motility and transport of soluble chemotactic agents, originating from either intra- or extracellular muscle sources, thereby slowing down the process of neutrophil 66 chemotaxis to local inflammatory sites [70]. The correlation data in this study partially support this 'looser' control theory between protease activation and neutrophil accumulation, within fast type skeletal muscle compared to ventricular muscle. Alternatively, the fact that the heart, by virtue of its continuous function and requirement for adequate blood supply, may respond more quickly to and/or receive (and incorporate) more chemotactic signals from the systemic-pulmonary 'venous' return. Although the factor(s) influencing tissue specific differences are uncertain, this does not detract from a major finding of the present study, that the processes of striated muscle calpain-like protease activity and neutrophil accumulation, as determined by myeloperoxidase activity, are positively related within individual samples immediately following exercise. Administration of cell permeable cysteine protease inhibitors, either E64c or E64d, have been reported to reduce the activity of the calcium activated neutral protease in rat striated muscles [100]. The use of (2S,3S)-trans-epoxysuccinyl-L-leucylamido-3-methyl-butane, E64c (at 1 mg/kg), was favored in the present investigation over oral administration of E64d for three reasons: a) E64d requires a much longer time course, 4-6 weeks, before any effect on calpain activity is observed [69]; b) E64d is a pro-drug of E64c, thus to exert any biological activity on tissue calpain levels it must be converted into E64c and circulate in the plasma [100] and c) i.p. administration of E64c to rats has been shown to inhibit tissue enzyme levels [3,69]. Because E64c does not affect the activity of myeloperoxidase in vitro, any changes in the tissue determination of this enzyme can be considered to be through an alternative mechanism(s). The observation that calpain-like activity was reduced following exercise and that myeloperoxidase activity was similarly 67 lowered after E64c pre-treatment, implicates a cause-effect link for the positive relationship reported in this study. Although the mechanism(s) underlying the cause-effect nature of these processes cannot be determined from the present results, the involvement of a myogenic factor linked to protease activation and/or protein breakdown (as reported to occur with exercise), which is chemoattractive to neutrophils is novel and forms the basis of our current working hypothesis. A schematic representation depicting the steps that could be involved in this hypothesis is presented (Figure 1). The suggestion that neutrophils possess a cytotoxic potential incapable of discriminating between host and/or foreign or abnormal proteins and tissues and thus must be strictly controlled by extraneous mediators/regulators [28,70,98] provides partial support for our results and subsequent hypothesis. Clearly in the rat, chemokines (i.e. CINC, MIP-2) and other cytokines (i.e. IL-8, TNF-a, IL-1 (3) produced by endothelial cells and/or resident macrophages within muscle, play a major role in mobilizing and attracting inflammatory cells to sites of injury, however other myogenic factors cannot be ruled out and may also play a role in targeting the tissue for neutrophil infiltration. As observed in these experiments, calpain activation occurs with exercise, and it may indeed play a direct role in the chemoattractive process to neutrophils [53,54,74]. Kunimatsu et al. [54] have shown significant neutrophil chemotactic activity with a peptide fragment of |i- and m-calpain, and have since sequenced an active N-acetyl peptide, which likely acts on the well characterized fMLP receptor of the neutrophil [53]. Because this region of the protease is released upon activation [73,81], this may be the biological mechanism underlying the results of this study. Further work is required to precisely identify the steps involved in the activation of 68 this calcium-dependent cysteine protease, calpain, and the accumulation of neutrophils into striated muscle following exercise. In summary, while much work has gone into the quantitative description of exercise related indicators of muscle injury/damage and accompanying protein degradation, the literature lacks specific mechanism(s) to explain the initiation of these exercise induced changes. This study has confirmed with exercise-induced injury, the release of a muscle-generated neutrophil chemotactic factor from striated muscle. It also reports a significant positive relationship between activation of a calpain-like protease and the biochemical index of neutrophil accumulation (i.e. myeloperoxidase activity) for cardiac and fast type muscles following exercise. In addition, a cause-effect link between these processes is provided with the use of a cell permeable cysteine protease inhibitor, E64c. A working model is proposed which describes a plausible mechanism underlying the data set which demonstrates the involvement of calpain in the generation of a potential myogenic factor with affinity for a neutrophil receptor. Whether these events underlie the initiation processes for exercise-induced muscle injury requires further investigation. 69 CONCLUSION The calcium activated neutral protease, calpain, is known to cause proteolysis in specific cytoskeletal proteins in muscle almost immediately following exercise-induced damage. Likewise, the first cellular occurrence of acute inflammation is the infiltration of neutrophils to the site of tissue damage. Thus, this study has proposed that the initial events in Ca2+-stimulated proteolysis could serve as a precursory link to neutrophil accumulation into injured muscle tissue. This is thought to occur through a mechanism whereby peptide fragments from calpain degradation serve as chemotactic factors which "chemically attract" neutrophils from the circulation mainstream, towards specific sites of tissue damage. Therefore, the hypothesis that there exists a direct/indirect relationship between the processes of calpain degradation and the migratory actions of neutrophils; both synergistically contributing to the active occurrence of the acute inflammatory response in damaged muscle tissue. 70 FUTURE RECOMMENDATIONS • To further explore the involvement of CINC and/or other IL-8 analogues in the progression of muscle-generated injury and neutrophil recruitment. • To explore the involvement of calpain sub-fragments and/or peptide fragments from calpain degradation in promoting neutrophil chemotaxis. • To isolate the specific chemotactic agent(s) involved in the recruitment of neutrophils to focal sites of muscle injury. 71 BIBLIOGRAPHY [I] Adachi, Y., A. 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Evidence for enhanced nonlysosomal proteolysis associated with elevated cytosolic calcium. /. Biol. Chem. 260: 13619-13624, 1985. 80 APPENDIX Table 3. Calpain substrates (adapted from Saido et al., 1994). A. Cytoskeletal proteins Actin-binding proteins (fodrin or spectrin, talin, filamin. a-actinin),* microtubute-associated proteins, etc. B. Membrane proteins Crowth factor receptors (EOF receptors), adhesion molecules (integrin, cadherin, N-CAM), ion transporters (Ca**-ATPase), C. Enzymes'" Kinases (protein kinase C, myosin light chain kinase, calmodulin-dcpendcnt kinase, ppjSO sre), phosphatases (calcineui phospholipases (phospholipase C), etc. D. Others Cytokines (interleukin la), transcription factors (Fos, Jun), lens proteins (crystalline), etc. 'Representative examples are jiven in parentheses. 'Some are cytosolic and othen are membrane associated. Table 4. Some of the proposed physiological functions of the calpain system (adapted from GoII et al., 1992). A. Activate or alter regulation of enzymes Enzyme , Effect of calpain cleavage Protein kinase C Caldoeurin Myosin fight-chain kinase Pbospbocylase kinase Factors V and XIII (transglutaminase) c-Jun and c-Fos Fully active but no longer requires Ca1* and phospholipid Fully active but no longer requires calmodulin Fully active but no longer requires Ca1? and calmodulin Activates independently of phosphorylation Activates both factors identically to the activation that occurs during cloning Inactivates pp**"0", the protein kinase product of the proto-oncogene, c-mas, that mrrcsls cells in (hem-phase ofmeiosis Inactivates both these DNA-binding proteins rapidly B. Remodeling/disassembly of the cytoskeleton System Effect of calpain cleavage Striated muscle Smooth muscfcAcytoskelcton in nonmusde cells Platelets Mitosis Alzheimer's plaques C Cleave hormone receptors Receptor Initiates metabolic turnover of the myofibrillar proteins Rapid cleavage: desmin. nebulin. troponin T Slower cleavage: C-protein, lititv. tropomyosin, troponin I No cleavage: actio, a-actinin, myosin, troponin C Removes dense bodies and attachment plaques: cleaves microfilament/plasma membrane attachments Rapid cleavage: ankyrin, band 3. filamin. MAP 2, protein 4.1, talin. vinculin Slower cleavage: spectrin, tubulin Rapidly cleaves all intermediate filament proteins including the neurofilament proteins (the !60-kDa>68-kD3t>200-kDa) Platelet activation involves massive membranefcytoskeletal rearrangement and is accompanied by calpain-like cleavages in talin, filamin. and spectrin: associated with shedding of vesicles? Intracellular [Ca14] increases at the initiation of mitosis and calpain-induced, degradation of tubulin. MAPs. and the cell cytoskeleton could be critical for spindle disassembly and cytokinesis H-Calpain cleaves amyloid precursor proteins to produce /S-amyloid peptide that is a core component of neurittc plaques*"1 Effect of calpain cleavage EOT receptor Estrogen receptor Progesterone receptor Otucocorlicoid receptor POOF receptor Hype Cal+-chanoel proteins Cleaves 160-kDa receptor to 145-kDa; no significant effect on EOF binding Cleaves &6S receptor to ^L6S forou no significant effect on binding ability . .r. . rt~.™ Cleaves A-subunU from 79-kDa to 43-kDa: B-subunit from 112-kDa to 23-kDa: no significant effect on binding ability -x Cleaves 90-kDa receptor to 50-kDa: no significant effect on binding m -Cleaves 185-kDa wl30-kDa polypeptide that retains Tyr-kinase activity and autophosphorylation sues Destroys Ca1+-channd properties • 81 Polymorphonuclear Cell Isolation Sprague-Dawley rats (~325g) anaesthetized with halothane 1. Draw via cardiac puncture -15 ml of blood into a heparin-coated 20-ml plastic syringe. Dilute the anticoagulated whole blood 1:1 in a 50-ml polypropylene tube with Hepes-normal saline (H-NS). Add 15 ml of Ficoll-hypaque 1077 into a clean 50-ml polypropylene tube and carefully layer on the 30 ml of H-NS diluted blood. Centrifuge at 400 x g (1600 rpm; Hermle-model Z 233-M) for 30 min at room temperature. 2. Aspirate off plasma leukocytes and platelets at the interface along with the Ficoll layer, down to the red blood cell layer (pellet). 3. Fill tube with NFUCl lysing solution. Gently mix and allow tube to sit a color change from matte red to glossy black occurs. Centrifuge at 400 x g for 7 min at 4°C. 4. Pour/aspirate off the supernatant and resuspend pellet in NH4CI lysing solution once more. Centrifuge at 400 x g for 7 min at 4°C. Aspirate off supernatant and resuspend pellet in H-NS. Centrifuge at 600 x g for 6 min at 4°C (1900 rpm; Hermle-model Z 233-M). 5. Resuspend the neutrophils at 1 million/ml in H-NS for temporary storage (i.e. <2 hours in ice), or in Hanks balanced salt solution (HBSS) just prior to use - as assessed by crystal violet staining on a grid hemocytometer. % Yield: Neutrophils >80 ± 4, Monocytes <15 ± 4, Others <5 ± 2. 82 Reagents Phosphate buffered saline: 8.00 g NaCI, 0.20 g KC1,1.15 g Na2P04,0.20 g KH2P04 in 990mlddH2O**, pH 7.40 0.01 M Hepes normal saline: 5 ml IM Hepes in 500 ml 0.9% saline**, pH 7.35 NH4Cl lysing solution: 8.02 g NH4CI, 0.84 g NaHC03, 0.37 g EDTA in 990 ml ddH20** fMLP: 10"2 M in DMSO (stock) diluted to desired concentration with HBSS** Crystal violet solution: 0.1% crystal violet, 0.1% acetic acid, 10% methanol Permount solution: Permount diluted with Histoclear® (xylene substitute) Diff-Quik® stain set: Fixative (methanol), Solution I (xanthene dye), Solution n (thiazine dye mixture) (Baxter™: McGaw Park, IL) * Unless otherwise stated, all chemical reagents were acquired from Sigma™: St. Louis, Missouri. ** Endotoxin-free grade 83 Protocol for Neuro Probem48 well chemotaxis chamber NEURO PROBE, INC PO Box 400 Cabin John, MD 20818 Tel: (301) 229-8598 Fax: (301) 229-8938 PROTOCOL FOR THE NEURO PROBE 48 WELL CHEMOTAXIS CHAMBER Stock Number AP48 PREPARING THE BOTTOM CHAMBER 1. Adjust a variable volume mlcropipette with a 1mm tip so that the ejected liquid fills a bottom well (approximately 25|d). The volume should be such that a slight positive meniscus forms; this prevents air bubbles from being napped when the filter is applied 2. Orient the bottom plate so that the NP trademark is at the upper right Fill the bottom wells with chemoattractants or control reagents which have been warmed to about 37°C and vortexed to expel dissolved gasses; the filling should take no longer than 5 minutes to. prevent excessive evaporative loss. 3. Cut off 1mm of the comer of a filter membrane. lift the filter by the ends with two forceps, and orient it to the chamber so that the cut corner corresponds to the NP trademark. Suspend the filter over the filled wells and lower it onto them, allowing die middle portion of the filter to make contact first (see Fig. 1). The filter position can be adjusted at this point, if necessary, but movement must be minimized to avoid contamination between wells. 4. Apply the silicone gasket with die cut corner at the upper right, then the top plate with the trademark also oriented at the upper right. Push the top plate down against the bottom plate and hold it down firmly with one hand; with the other hand apply the thumb nuts. It is important to maintain even, firm pressure on the assembly while you tighten die hardware; this prevents air bubbles from being drawn into the bottom wells. The thumb nuts should be finger-tight; do not use pliers or other tools to tighten themv PREPARING AND ADDING RESPONDING CELLS 1. Cells are prepared and suspended in medium. The concentration of cells in the suspension should be adjusted so that 45-50jd contains the desired number of cells for one well. Note that the exposed filter area for each well is 8mm2, so, for example, a suspension of 8,000 cells in 50jd will yield 1,000 cells/mm2, and 50,000 cells in 50ui will yield approximately 6,000 cells/mm2. 2. Pipette 50ul of cell suspension into each upper well. Although different volumes can be used, this volume fills the wells and creates a slight positive meniscus, which makes checking for bubbles easy (note that it is essential to avoid trapping bubbles In this step). Hold the pipette at -a steep angle so that the end of the pipette tip rests against the-wall of the chamber Just above the filter, and the side of the tip rests against the top rim of the well. Eject the fluid with a rapid motion to dislodee air in the bottom of the well. 84 3. Check for trapped bubbles in the upper wells. One easy way to do this is to look at the reflections of overhead lights in the menisci: a well with an abnormally large positive meniscus usually has a trapped air bubble. Remove any bubbles by sucking the well completely dry with a suction line and disposable pipette tip, then refill it. 4. For most chemotaxis assays the filled chamber is incubated at 37°C in humidified air with 5% CO2. Incubation times vary considerably depending on cell types and chemotactic factor(s). One good way to determine the optimum incubation time is to use 6-12 blind well chambers (e.g. catalogue #BW100) set up as negative controls and placed simultaneously in the incubator. Remove one blind well after a set period (e.g. 30 minutes), and remove the rest sequentially, one every 5 minutes. Stain the filters and examine them to see how long cells have taken to migrate to a specified optimum depth in cellulosic filters, or unstimulated cells to simply migrate through polycarbonate filters. REMOVING, WIPING AND STAINING THE FILTER 1. Remove the thumb nuts while holding down the top plate, and invert the entire chamber onto a paper towel. Grasp the four corners of the top plate and push down evenly so the top plate stays parallel to the bottom plate as it drops to the table (see Fig. 3). If the gasket should hang up on the post hardware, carefully push it down evenly onto the top plate. Take care not to touch the filter, which should be stuck to the gasket. Immerse the bottom plate in cool distilled water. 2. The migrated cells are now facing up on the filter — this side of the filter is henceforth referred to as the "cell side." Lift up one end of the filter with forceps and catch 1mm of the edge in the wide plastic filter clamp (see Fig. 4). Lift the filter and quickly attach the small clamp to the edge of the free end. « 3. Keeping the cell side up, wet the underside (non-migrated cell side) of the filter in a dish containing PBS (see Fig. 5). Do not let the PBS wash over the cell side of the filter. 4. Holding the filter by the large clamp, with the small clamp attached to the other end and hanging, free, wipe the cells off the non-migrated side of the filter by drawing the filter up over the wiper blade (Fig. 6). The blade should first contact the filter just below the Jaws of the wide damp. Pressure against the blade should be gende; maintain an angle of about 30° from the vertical for the portion of the filter above the wiper. The time between wetting the filter and completion of wiping must be minimized to avoid drying of the cells on the filter and consequent incomplete cell removal (drying occurs within 10-20 seconds). 5. Clean the wiper with a Q-tip, wet the underside of the filter again in PBS and repeat Step 4. Clean the wiper again, then wet the filter a third time in PBS and repeat Step 4. 6. For granulocytes and monocytes, carefully immerse the filter in methanol, then place the filter cell side up on a disposable lint-free towel for air drying. Rinse all chamber components in cool distilled water. For other kinds of cells, consult the literature for staining techniques. 85 7. When the filter is dry, clamp the edge of one end with a wide blade clamp, weight the other end with the small clamp and stain in Diff-Qulk® (Baxter Scientific Products), or equivalent, according to the manufacturer's instructions. To avoid contaminating the chamber components with stain, it is convenient to have two sets of filter clamps, one for removing the filter from the gasket, and one for staining. 8. Place the wet filter cell side up on a 50x75 microscope slide to dry. When the filter Is dry, center it on the slide and place a drop of immersion oil on It. Rub the oil over the filter with a smooth, blunt instrument to remove all bubbles and wrinkles. The filter is now ready for counting. WARNINGS CONCERNING THE USE OF ACRYLIC CHAMBERS • NEVER autoclave acrylic chambers; they will warp, deform or melt. • NEVER immerse acrylic components of chamber in water hotter than 60° C. • NEVER allow acrylic chamber components or silicone gaskets to come in contact with solvents (e.g. acetone, carbon tetrachloride, alcohol). • Ultrasonic cleaning of chambers is NOT recommended. • NEVER place acrylic components in drying oven or under UV light. • NEVER allow solutions, especially ones containing proteins, to dry on chamber components. • NEVER blow dry with compressed air containing oil or other contaminants. CLEANING AFTER EACH USE This is a precision instrument; It should be handled and washed Individually. The hardware in the bottom of the chamber, or any other hard object, should not be allowed to bang Into the sealing surfaces of acrylic components. 1. Immerse chamber components In distilled or demlnerallzed water immediately after disassembly. Make sure that trapped air bubbles in the wells are eliminated. 2. Rinse all parts in running distilled water and drain on paper towels; blow dry with CLEAN compressed gas, e.g. nitrogen or filtered air. Store chamber components at room temperature, covered for protection. 86 PERIODIC CLEANING Proteins and other contaminants tend to accumulate in the chamber's wells and on the gasket. Accumulation can be fast or slow depending on the nature of the solutions used, but is immediate and severe if protein solutions are ever allowed to dry in the wells. Periodic soaking of all the chamber components in Terg-a-zyme®, an enzyme active detergent (Baxter or Fisher), for 1-3 hours at 60°C usually removes the residues. Follow with thorough rinsing under running distilled/deionlzed water, and soak in the same overnight. You can use this cleaning method weekly, if necessary, depending on the conditions under which you are using the chamber. After cleaning as above, if continued contamination is indicated by the presence of erratic results or by negative controls migrating faster than normal, the gasket is often the source of the problem. The gasket should be soaked in hot Terg-a-zyme® (100°C), and ultrasonically cleaned if possible. (Note that the gasket is significantly heat and scratch resistant, but is very sensitive to solvents.) If problems persist after this treatment, we recommend two courses of action. Firsc, run some controls in blind well chambers parallel to the multiwell chamber. Blind well chambers don't use gaskets, so this will help determine if the multiwell gasket is contaminated. Second, keep a spare gasket on hand and replace the original. CHAMBER STERILIZATION NEVER AUTOCLAVE ACRYLIC COMPONENTS. If sterilization is required for studies with long incubation times, or due to the presence of pathogens, acrylic and metal components may be sterilized in one of three ways: • gas sterilizing with ethylene oxide • soaking for 30-60 minutes in 1 molar sodium hydroxide at 60°C, followed by thorough rinsing in sterile deionlzed water in a sterile environment • soaking in a weak bleach solution (1 T Clorox®/gallon distilled, deionlzed water) for 30-60 minutes, followed by soaking In sterile deionized water in a sterile environment. This procedure will usually kill all bacteria; if other pathogens are present, the strength of the solution and/or duration of the soak may need to be modified. The gasket (ONLY) may be autoclaved or boiled in deionized water. Do not immerse the gasket in bleach or sodium hydroxide solutions; it is porous and will absorb and later bleed these solutions. 87 88 Microplate Assay for Calpastatin-inhibited Calcium-dependent Proteolysis Calpain like Activity * KEEP ALL SAMPLES, TEST TUBES AND EPPENDORFS ON ICE (OR REFRIGERATED) DURING THE PROCEDURE Before Starting: Thaw a vile of DTT (2 ml-200 mM) and a test tube of Casein (5 ml-10 mg/ml). Preheat dry bath to 30°C. PROCEDURES: A. Preparing a Muscle Suspension: -add 1 ml of Buffer I to labeled test tubes (1 tube/sample). -add 0.1 ml DTT to each tube, vortex and place on ice. -weigh muscle sample (~100 mg is best size-cut if required) -if muscle sample is frozen, after weighing, transfer immediately into Buffer I (DON'T LET SAMPLE THAW !) -homogenize sample using a Polytron for ten seconds (twice) at a setting of 6. Be sure that all tissue is in suspension. (IMPORTANT: Clean Polytron After Every Use) B. Separating Soluble and Particulate Fractions: -transfer the suspension (from step A) into eppendorf tubes (~2.0 ml) and place in pre-cooled rotor (0-4°C). -centrifuge samples (HERMLE Z233M) at 13,000 rpm for 15 minutes, -remove the supernatant and transfer it to a labeled eppendorf tube (on ice). This soluble fraction should be removed by the use of a Pasteur pipette and care taken not to dislodge the pelleted material. -to the pellet, add 1 ml of the Buffer II and re-suspend it by vortexing. (Buffer U is prepared by a 2:1 mixture of Buffer I and Myofibril Wash Buffer (with 1.0% Triton X-100). This results in 0.33% Triton X-100 which is critical for removing the bound fraction while maintaining calcium-dependent proteolytic activity, -transfer the re-suspended pellet to a 2 ml glass tissue homogenizer. Homogenize using approx. 8-12 hearty strokes. -transfer contents back into original eppendorf and centrifuge at the same settings as before. -remove the supernatant and place on ice, this is the particulate (bound) fraction. C. Preparation of Samples for Assay: -4 Eppendorf tubes are needed per sample; soluble unactivated, soluble activated, bound unactivated, and bound activated, plus one more for the blank. The tubes should be set up as shown in the following table, using the amount of solution as shown per tube: 89 SOLUTIONS (In ul) BUFFER 1 SAMPLE CAESIN CaCl2 ddFLO EPPENDORF TUBES BLANK 200 0 100 0 75 1) SOLUBLE 0 200 100 0 75 UNACTIVATED 2) SOLUBLE 0 200 100 75 0 ACTIVATED 3) BOUND 0 200 100 0 75 UNACTIVATED 4) BOUND 0 200 100 75 0 ACTIVATED -the order of addition is not important, however the incubation time is started with the addition of sample (either soluble or bound fractions) -as soon as the sample/fraction is added in each eppendorf tube they are vortexed and incubated at 30 C (dry-bath) for 30 min., remembering to vortex each tube again after 15 minutes. D. Prepare Reader. Microplate For Protein Assay: Plates: 1) while the samples are in centrifuge prepare a 1:1 dilution of BioRad protein assay solution and double distilled H2O (ddH20). 2) power up microplate reader (it takes 15 min to warm up and go through self-check diagnostics). 3) prepare microplate by pipetting 100 ul of the diluted BioRad solution into each well (remember each sample, both soluble and particulate fractions, are assayed in triplicate). 4) add 50 ul of ddFLO to each well. Reader: 1) in microplate reader software, click on "File...", then "Open...", then click on FORMAT FILENAME "XXXXXXXXX" (if you do not have a filename set up, you will need to edit an existing file and/or create your own format file. Remember this is important because the program will analyze the samples as you indicate in this format file. 90 2) next click on "Analysis" and then "Read Plate..." to Set Parameters, as follows: a) Single wavelength b) 595 nm measured wavelength c) 340 nm reference wavelength d) 10 sec. Mix Duration e) 9:59 min. Time Delay Protein Assay: -immediately after 30 minutes of incubation, retrieve sample tubes from dry-bath, starting with the blank and continuing in an orderly fashion, vortex each one, then add 100 ul of the sample solution to the appropriate wells on the microplate, remembering to do triplicates (3 WELLS PER SAMPLE) for each sample/fraction. -when finished, place microplate in the reader and start the reader. -print raw data report and click on "Show Calculation Table", print these results. 91 REAGENTS FOR CALPAIN MICROPLATE ASSAY BUFFER 1: (pH7.1) -sodium borate (39 mM) -KC1 (25 mM) -EGTA (5 mM) BioRad PROTEIN ASSAY SOLUTION MYOFIBRIL WASH BUFFER: (as Buffer I with 1.0% Triton X-100 and no EGTA) DTT:(200 mM) CaCl2:(50 mM) CAESIN:(10 mg/ml) 92 Myeloperoxidase Assay Working Solutions: 1. 0.3 % solution of ddH202: 1 ml of 30% stock solution + 100 ml ddH20 2. Phosphate buffer: 500 ml ddH20 + 0.6805 g KH2P04; KOH until pH = 6.0 3. HTAB: 200 ml ddH20 + 1 g HTAB, stir and heat 4. DMB (made fresh daily): 0.0032 g DMB + 25 ml ddH20 PROCEDURE: (to prepare one sample) 1. Stir and heat HTAB while preparing DMB 2. Put TT in ice, with 70 ul of HTAB 3. Weigh muscle sample and put in TT, then add more HTAB to reach 1 ul per mg of muscle 4. Homogenize two times for 15 sec 5. Centrifuge for 12 minutes, at 2900 rpm, at 0 degrees 6. Freeze and thaw 2 times with liquid N2 7. Centrifuge again, same settings READING REACTION with SPEC: (prepare 2 blanks and 1 sample) Program "16-MPO" on the spec will read sample for 150 sec, at intervals of 5 sec, with an initial lag of 2 sec, at 480 nm. Use the circulating heater to maintain 37 degrees. 1. Each tube gets: 500 ul of 0.3% H202,1000 ul of buffer, 50 ul of DMB, then mix 2. 450 ul of HTAB to each blank and zero spec 3. Put solution from #1 into cuvet, put in spec, and allow 2 minutes to heat to 36 degrees 4. Add 450 ul of supernatant to sample cuvet and read against blank for 150 sec while reaction takes place. When adding supernatant, use micro pipette, add 450 ul, suck up solution, and squirt it in again to mix. 93 Table 5. Results: individual sample means for neutrophil chemotaxis 1 1 — V^UIILl UI VJ IUWUV 1 1U Condition 1 2 3 4 5 Mean S.D. Normalized Hanks with Ca2+ 2 3 2 3 2 2.4 0.5 PBS 1 0 1 3 1 1.2 1.1 fMLP 10-8 M 96 107 98 58 115 94.8 21.9 fMLP 10-7 M 124 124 127 101 166 128.4 23.5 Soleus - Undiluted 28 50 58 36 24.8 39.3 14.4 31.8 Soleus -1/10 3 2 3 7 2 3.4 2.1 Soleus -1/100 0 0 1 4 2 1.4 1.7 Soleus -1/1000 0 0 0 2 1 0.6 0.9 Plantaris - Undiluted 13 23 21 20 26 20.6 4.7 16.2 Plantaris -1/10 2 6 2 3 2 3.0 1.7 Plantaris -1/100 0 0 1 2 1 0.8 0.8 Plantaris -1/1000 0 0 0 1 1 0.4 0.5 Heart - Undiluted 68 50 67 42 75 60.6 13.7 47.1 Heart -1/10 3 19 6 24 6 11.6 9.3 Heart -1/100 0 2 1 6 3 2.4 2.3 Heart -1/1000 0 0 0 1 1 0.4 0.5 Condition 1 2 3 4 5 Mean S.D. Normalized Hanks with Ca2+ 3 2 2 1 2 2.0 0.7 PBS 3 3 3 1 1 2.2 1.1 fMLP 10-8 M 147 126 169 223 175 168.0 36.3 fMLP 10-7 M 188 166 221 311 227 222.6 55.3 Soleus - Undiluted 24 14.1 16.5 24 22.3 20.2 4.6 9.3 Soleus -1/10 3 3 2 7 2 3.4 2.1 Soleus -1/100 2 2 2 4 1 2.2 1.1 Soleus -1/1000 0 1 1 1 0 0.6 0.5 Plantaris - Undiluted 12 10 9 14.9 9 11.0 2.5 5.0 Plantaris -1/10 6 2 4 4 1 3.4 1.9 Plantaris -1/100 1 2 3 2 0 1.6 1.1 Plantaris -1/1000 0 1 1 0 1 0.6 0.5 Heart - Undiluted 26.5 22 21 30 18 23.5 4.7 10.9 Heart -1/10 5 6 3 9 2 5.0 2.7 Heart -1/100 4 1 1 3 1 2.0 1.4 Heart -1/1000 2 1 1 1 1 1.2 0.4 94 1 ' . f!onfml Snliihto Fmr>finn j .. Condition 1 2 3 4 5 Mean S.D. Normalized Hanks with Ca2+ 2 3 1 1 1 1.6 0.9 PBS 2 1 1 1 1 1.2 0.4 fMLP 10-8 M 169 165 168 176 163 168.2 5.0 fMLP 10-7 M 245 211 230 240 213 227.8 15.4 Soleus - Undiluted 18 21 17.5 20 25 20.3 3.0 9.0 Soleus -1/10 2 3 2 2 3 2.4 0.5 Soleus -1/100 2 1 2 1 1 1.4 0.5 Soleus -1/1000 1 0 1 0 1 0.6 0.5 Plantaris - Undiluted 20 20 23 21 22 21.2 1.3 9.3 Plantaris -1/10 3 2 2 2 2 2.2 0.4 Plantaris -1/100 2 1 2 1 1 1.4 0.5 Plantaris -1/1000 1 1 1 1 1 1.0 0.0 Heart - Undiluted 15 16 22 16.8 18 17.6 2.7 7.7 Heart -1/10 2 2 2 2 2 2.0 0.0 Heart -1/100 1 0 1 1 1 0.8 0.4 Heart -1/1000 0 1 1 1 1 0.8 0.4 i i i F,w>iv»icA finliihlo PVa/»f!r»n Condition 1 2 3 4 5 Mean S.D. Normalized Hanks with Ca2+ 1 1 0 1 1 0.8 0.4 PBS 1 0 1 3 1 1.2 1.1 fMLP 10-8 M 163 77 132 144 144 132.0 32.7 fMLP 10-7 M 188 161 165 179 182 175.0 11.5 Soleus - Undiluted 82 81 78.8 84 82 81.6 1.9 46.7 Soleus -1/10 15 5 5 2 6 6.6 4.9 Soleus -1/100 4 2 2 1 2 2.2 1.1 Soleus -1/1000 2 0 1 2 2 1.4 0.9 Plantaris - Undiluted 34 38 40.7 37 45 38.9 4.1 22.3 Plantaris -1/10 5 3 3 3 3 3.4 0.9 Plantaris -1/100 2 2 0 2 2 1.6 0.9 Plantaris -1/1000 1 0 0 2 1 0.8 0.8 Heart - Undiluted 9 10 7 5 7 7.5 1.8 4.3 Heart -1/10 1 4 1 2 1 1.8 1.3 Heart -1/100 0 1 0 1 0 0.4 0.5 Heart -1/1000 0 1 0 2 1 0.8 0.8 95 


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