86 L.V. Thompson Figure 7 is a representation of the low-field EPR spectra collected during maximal isometric contraction. During an experiment, three different spectra are collected, rigor, relaxation and contraction. Each rigor, relaxation, and contraction spectrum is collected at the same field positions (e.g., 1,024) and over the same gauss range (e.g., 38-gauss). After collection, the 38-gauss low-field EPR spectra are analyzed to determine the fraction of myosin heads in the strong-binding struc- tural state (x) during muscle contraction. For each experiment (e.g., fiber bundle), the spectrum obtained during maximal isometric contraction (V Con ) are analyzed as a linear combination of the spectra obtained during rigor and relaxation using ( ) 1 Con Rig Rel V xV x V= +− (1) where V Rig (rigor) corresponds to all heads in the strong-binding structural state (x = 1) and V Rel (relaxation) corresponds to all heads in the weak-binding structural state (x = 0). Thus, for the contraction spectrum, x is solved at each of the field A B Relaxation Contraction Rigor Fig. 7 Portion of the low-field EPR spectra of spin-labeled muscle fibers. The EPR spectra are collected (3,425 G central peak, 38 G sweep width, 5.0 G peak-to-peak modulation amplitude, and 16 mW microwave power) under conditions of rigor in which all heads are in the strong-binding structural state (black), relaxation in which all heads are in the weak-binding structural state (red), and contraction (green). Spectra obtained during maximal isometric contraction are analyzed as a linear combination of the spectra obtained during rigor and relaxation. For each fiber bundle, the spectrum obtained during maximal isometric contraction (V Con ) is analyzed as a linear combination of the spectra obtained during rigor and relaxation using V Con = xV Rig + (1 − x)V Rel , where V Rig (rigor) corresponds to all heads in the strong-binding structural state (x = 1), and V Rel (relaxation) corresponds to all heads in the weak-binding structural state (x = 0). Thus for the contraction spectrum, x is solved at each of the 1,024 field positions to determine the fraction of myosin heads in the strong-binding structural state, using x = (V Con − V Rel )/(V Rig − V Rel ). A/B is equal to the mole fraction of x of myosin heads in the strong-binding state, as indicated in Eq. 2 87Age-Related Decline in Actomyosin Structure and Function positions (e.g., 1,024) to determine the fraction of myosin heads in the strong-binding structural state as follows x = (V Con − V Rel )/(V Rig − V Rel ) = A/B and as illustrated in Fig. 7. 4.3 Age-Related Structural Changes in Myosin Protein Structure Figure 8 summarizes the major findings of one of the first studies to investigate age-related alterations in the distribution of myosin structural states (Lowe et al. 2001). The study hypothesized that the reduced force-generating ability of skeletal muscle fibers from aged animals was due to a decreased population of myosin heads in the strong-binding (force-generating) structural state during muscle contraction relative to skeletal muscle fibers from younger animals. The results clearly demonstrated that the only detected difference between young and aged fibers is in the mole fraction x of myosin heads in the strongly bound structural state. During a maximal isometric contraction (same length contraction), 30% fewer A B + A B Young adult Aged % of young adult 120 100 80 60 40 20 0 Specific Force x, fraction strong-binding myosin Rigor ab Relaxation Contraction Young Old Fig. 8 Differences in fiber-specific force and myosin structural distribution with aging. Panel A – Representative EPR spectra of spin-labeled muscle fibers from young adult and aged rats. For the aged fibers the contraction spectrum (green) is closer to the relaxation spectrum (red) than is the contraction spectrum (green) to its respective relaxation spectrum (red) of the young adult fibers; i.e., A/B is less for the aged fibers. Panel B – Deficits of specific force (force/cross- sectional area) and fraction of myosin heads in the strong-binding structural state during maximal isometric contraction. * – Significantly different from young adult. The myosin structural changes can provide a molecular explanation for age-related decline in skeletal muscle force generation (Modified from Lowe et. al. 2001) 88 L.V. Thompson myosin heads are in the strong-binding, i.e., force-generating, structural state in fibers from aged animals (x = 0.22) than in fibers from younger animals (x = 0.32). Because it is usually proposed that the force generated during contraction is directly proportional to x and because the decrease in x (30%) is in good agreement with the decrease in specific force (27%), the study provides direct evidence that the decre- ment in force-generating capacity of fibers from aged animals is a direct result of a reduced fraction of myosin heads in the strong-binding structural state. This approach has effectively proven the number of force-generating cross-bridges per unit area during a muscle contraction is reduced with age, indicating that age- related changes in myosin structure represent a likely molecular mechanism under- lying muscle weakness. Therefore, evidence suggests that the structure and function of myosin are altered with age. 5 ATPase Activity 5.1 Experimental Approaches ATPase measurements are required for progress in understanding the molecular basis of age-related deterioration of contraction velocity because it is well known that shortening velocity is directly related to, and dependent upon, myosin ATPase activity. Several experimental approaches are used to determine ATPase activity, permeabilized skinned fibers, myofibrils, and purified proteins (actin and myosin). The myofibril preparation is a relatively simplistic experimental system for study- ing ATPase activity. Myofibrils are functional muscle units comprised of sarcom- eres such that the thick-thin filament structure is maintained. Myofibrils in solution containing ATP and Ca 2+ contract freely corresponding to the condition of an unloaded muscle fiber during shortening. The myofibrillar preparation allows quantitative determination of the protein concentration and specific enzymatic activity (ATPase). In muscle, total protein content is mainly actin (18–22%) and myosin (43–50%) (Ingalls et al. 1998; Yates and Greaser 1983). However, since the interaction between actin and myosin in the muscle filament lattice also depends on other structural and regulatory proteins, a more direct assessment of age-related changes in the interaction between muscle myosin and actin requires specific measurements of biochemical and structural properties of actin and myosin. Purified actin and myosin can be used to determine ATPase activity too, permitting the determination of enzymatic changes in each of these proteins, independently of the other and without interference from other myofibrillar proteins. Myosin ATPase activity can be determined at high and low salt concentrations, revealing critical information about the proteins (e.g., Bobkova et al. 1999). Myofibrillar ATPase activity at high salt concentration eliminates effects of actin and other proteins, and is sensitive to post-translational changes in myosin, 89Age-Related Decline in Actomyosin Structure and Function particularly the covalent modification of specific cysteines and lysines, depending on the principal cation present (Ca 2+ or K + ) and on the affected sites and modifying reagents. Although myofibrillar ATPase activity at high salt concentration provides critical information about myosin, it is not physiological. Physiological ATPase depends on the state of myosin as well as actin and is directly associated with con- traction velocity or V o (e.g., Barany 1967; Marston and Taylor 1980). Physiological ATPase can be determined in myofibrils or in purified myosin and actin (i.e., less than 0.3 M ionic strength, in the presence MgCl 2 ). In addition to determining myosin ATPase with purified proteins, purified actin and myosin can be directly studied using the in vitro motility assay, a novel method for analyzing the interaction of actin and myosin at the single-molecule level. In this assay, isolated myosin molecules are immobilized on the glass surface, fluorescent actin filaments are added, and sliding movement of these filaments, initiated by addition of ATP, is directly observed under an optical microscope. 5.2 Age-Related Changes in Myosin ATPase 5.2.1 Myofibrils To more directly assess the possibility of low myosin ATPase activity being a mechanism underlying age-reduced shortening velocity, Ca 2+ – activated myosin ATPase activity is measured in freely contracting myofibrils corresponding to conditions of maximal unloaded shortening velocity. Under these conditions, Ca 2+ - activated myosin ATPase activity is 16% lower with age and this decrease is similar to the reduced shortening velocity in permeabilized fibers from the same experimental group (Lowe et al. 2004). The results provide important evidence that changing actin-myosin interactions contribute to age-related inhibition of contractility. Considering the contracting myofibrils and the respective ATPase activity data, it is possible to draw conclusions about how age influences the ATPase cycle. It has been shown that ADP release is the rate-limiting step during an isometric contraction in fibers and in myofibrils that are chemically cross-linked such that they do not shorten during contraction (Dantzig et al. 1992; Lionne et al. 2002). In contrast, both ADP release and P i release have been implicated as the rate-limiting step during shortening contractions (Lionne et al. 1995, 2002). Therefore, it is probable that P i or ADP release from myosin during a shortening contraction is reduced with age. Based on the available evidence, it is likely that ADP release is the rate-limiting factor with age because during a maximal isometric contraction there is an increase in the apparent rate constant for myosin detachment from actin with age and ADP release is the critical step controlling detachment (Fig. 4). 90 L.V. Thompson 5.3 High-Salt ATPase Age-related molecular changes in myosin are observed in high-salt ATPase activities of myofibrils and purified myosin (Fig. 9b) (Prochniewicz et al. 2005). Like single muscle fibers described previously, the age-induced changes are muscle-specific, strain-dependent and independent of changes in myosin isoform expression (reviewed in Prochniewicz et al. 2007). The observed changes in the ATPase activities provide strong indication of age-related post-translational modifications of myosin. However, high-salt ATPase activities do not provide sufficient information to determine the sites or nature of modification, nor to predict the functional consequences. 0 0.4 0.8 1.2 Fraction of A y M y Fraction of A y M y 0 0.4 0.8 1.2 * * V max * K m 0 0.4 0.8 1.2 ATPase, fraction of young K-ATPase Ca-ATPase * Actin, µM ATPase rate, sec 1 2006040 0 16 12 8 4 YoungYoung OldOld A y M y A y M o A o M y A o M o A y M y A y M o A o M y A o M o Fig. 9 Age-related molecular changes in myosin are observed in high-salt ATPase activities of purified protein. Panel A – Representative experiment with isolated myosin and actin proteins from young and old rats to determine Vmax (the maximum rate) and Km (actin concentration at half Vmax). The actin-activated myosin ATPase rate was measured at increasing concentrations of actin. The Vmax and Km were determined with O = young myosin, young actin, M y A y , Vmax = 16.43 ± 1.00 s −1 , Km = 7.44 ± 1.63 mM; • = old myosin, old actin, M o A o , Vmax = 12.55 ± 0.62 s −1 , Km = 8.98 ± 1.48 mM. Panel B. Age-related changes in myosin high-salt ATPase experiments. Age-related changes in the ATPase activity of myosin. K-ATPase was determined in the presence of 0.6 M KCl 50 mM Tris, and 10 mM EDTA; Ca-ATPase was determined in the presence of 0.6 M KCl, 50 mM Tris, and 10 mM CaCl 2 . The ATPase rates for old myosin are expressed as fraction of the rates for young myosin. Panels C, D – Age-related changes in the actomyosin function are due primarily to changes in myosin. V max and K m for actin-activated myosin ATPase are deter- mined as in panel A. Data are normalized to the value for young actin and young myosin, A y M y . A o M o = old actin and old myosin; A y M o = young actin and old myosin; A o M y = old actin and young myosin. * – Statistically significant (Modified from Prochniewicz et al. 2005) 91Age-Related Decline in Actomyosin Structure and Function 5.4 Purified Protein Biochemical experiments on purified actin and myosin provide more direct and detailed evidence about age-related changes in actin-myosin interactions with age, thus contributing to age-related inhibition of contractility (Prochniewicz et al. 2005). The studies show a decrease in two parameters of the actomyosin ATPase, V max (activity extrapolated to infinite actin concentration) and K m (the concentration of actin at half V max ) (Fig. 9a), providing direct support for the role of changes in the contractile proteins in the deterioration of muscle function. Subsequent mixing of actin and myosin from young and old muscle in four combinations shows that the age-related decrease in V max is primarily due to changes in myosin. This decrease in V max is consistent with the findings showing age-related alterations in the structural states of myosin (transitions from weak to strong interactions, Fig. 9c). Yet, the age-related decrease in K m is due to changes in both proteins, as actin from young muscle attenuates the age-related decrease in K m for myosin from old muscle (Fig. 9d). The changes in K m are related to changes in the equilibria between actin and myosin-nucleotide complexes at the final stages of the cycle. The data from these experiments indicate that changes in actin, together with changes in myosin, are involved in the molecular mechanism of age-related deterioration of muscle contractility. 5.5 In Vitro Motility Assay Studies using the in vitro motility assay show direct evidence supporting the role of molecular changes in myosin in age-related deterioration of contractility (D’Antona et al. 2003; Hook and Larsson 2000; Hook et al. 2001). In one study using purified myosin from the human vastus lateralis muscle demonstrates that the observed decrease of sliding speed of actin on myosin from aged muscle is compa- rable to the age-related decrease in the maximal unloaded shortening velocity V o reported in single skeletal muscle fibers (D’Antona et al. 2003). Consistent with the human study, a decrease in actin sliding speed on myosin was confirmed in aging studies using purified proteins from muscles from rodents (Hook and Larsson 2000; Hook et al. 2001). Interestingly, there are differences in the extent of deterioration between experi- mental preparations. For instance, the 12–25% decrease of actin sliding speed on myosin with age is much less pronounced than the decrease in V o (47%) in permea- bilized fibers (Hook and Larsson 2000; Hook et al. 2001; Li and Larsson 1996). The greater age-effects on fiber contractility than on the sliding velocity of purified actin filament in the in vitro motility reflect age-related changes in myosin as well as in the structural and thin filament proteins within the fiber lattice. Furthermore, the quantitative differences between age-related changes in contractile and enzymatic functions could also result from the complex mechanism of mechanochemical coupling in the actomyosin interaction. 92 L.V. Thompson 5.6 Single Permeabilized Skeletal Muscle Fibers/Isometric Contractions In contrast to contractions that allow shortening to occur, investigations focused on isometric contractions (muscle contracts without a change in length), in which the myosin ATPase rate is much slower and its kinetics are strongly affected by cross-bridge strain reveal changes in energetic efficiency and myosin cross-bridge kinetics with age (Lowe et al. 2002). Studies using the permeabilized fiber preparation, in which simultaneous measurements of force and ATPase activity were determined, show that fibers generate ~20 lower maximum force without changes in the ATPase activity (Lowe et al. 2002). This result indicates a decrease in the energetic efficiency, a partial uncoupling between ATPase activity and force generation, during isometric contraction in aged muscle. Moreover, the apparent rate constant for the dissociation of strong-binding myosin from actin was ~30% greater in fibers from aged animals, indicating that the lower force produced by fibers from aged animals is due to a greater flux of myosin heads from the strong-binding state to the weak-binding state during contraction (Fig. 10). The changes in cross-bridge kinetics are consistent with the observed structural changes in myosin during contraction with age. Overall, the observed changes in contractility with age using permeabilized fibers, myofibrils, and purified proteins provide strong indication of age-related post-translational chemical modifications. X w X s (no force) (force-generating) f app g app weak-binding myosin Strong-binding myosin Y = 0.68 A = 0.73 Y = 0.32 A = 0.27 Y = 9.52 A = 12.21 Y = 4.48 A = 4.51 Fig. 10 Myosin structure and kinetics during a maximal isometric contraction are altered with age. Myosin structural data, from electron paramagnetic resonance spectroscopy experiments show that the age-related fractional reduction of myosin in the strong-binding structural state (x s ) during an isometric contraction is proportional to the decline in force in those fibers (i.e., a 16% decline in force generation with age relates to a 16% reduction in x s ). The age-related reduction in strong-binding (force-generating) myosin is due to an increase in the apparent rate constant for myosin detachment from actin (g app ) from 9.52 to 12.21. Y, young adult; A, aged; f app , apparent rate constant for myosin attachment to actin; x w , fraction of myosin heads in weak-binding structural state (Modified from Lowe et al. 2002) 93Age-Related Decline in Actomyosin Structure and Function 6 Oxidative Stress Age-related deterioration of muscle function may involve ‘damage’ of muscle proteins by reactive oxygen and nitrogen (ROS and NOS) species. Post-translational chemical modification of proteins affects the protein’s structural and functional integrity and in vitro studies where there is an elevation in ROS and NOS show contractile inhibition (e.g., Callahan et al. 2001; Lamb and Posterino 2003). In this chapter, the term ROS includes not only the oxygen radicals but also non-radical derivatives of O 2 , such as H 2 O 2 . Hence, all oxygen radicals are ROS, but not all ROS are oxygen radicals. ‘Reactive’ is a relative term because superoxide anions (O 2 • − ) and H 2 O 2 react fast and are very selective in their reactions, whereas, hydroxyl radical (HO • ) reacts fast and is very promiscuous. 6.1 ROS Generation ROS are generated in multiple compartments and by multiple enzymes within the cell. ROS are continually generated as byproducts of normal aerobic metabolism, yet can be produced to a greater extent under stress and pathological conditions, as well as taken up from the external environment. Examples of ROS include unstable oxygen radicals such as superoxide anion (O 2 • − ) and hydroxyl radical (HO • ), non- radical molecules like hydrogen peroxide (H 2 O 2 ) and peroxynitrite (ONOO − ). While nitric oxide (NO • ) itself is not highly reactive or toxic, the reaction of NO • with other molecules in the cell can produce more toxic species (O 2 -derived species leading to the formation of reactive nitrogen oxide species). For instance, the reaction of NO • with O 2 •− produces peroxynitrite (ONOO − ) which may be the primary mechanism by which NO • causes cellular injury and alterations in function because it is a highly reactive species that can oxidize cellular lipids, proteins, and nucleic acids. 6.2 Role of Mitochondria Intracellular ROS are primarily generated by the mitochondria. Mitochondria consume ~90% of a cell’s oxygen to support oxidative phosphorylation (OXPHOS) which is the major metabolic system for ATP. Specifically, the process uses the oxidation of NADH or FADH 2 to generate a potential energy for protons across the mitochondrial inner membrane. Subsequently, this potential energy for protons is used to phosphorylate ADP. At several sites along the electron transport chain, electrons can directly react with oxygen or other electron acceptors and generate free radicals. 94 L.V. Thompson Figure 11 summarizes the mitochondrial electron transport chain (ETC). Carbohydrates (TCA cycle) and fats (b-oxidation), provide the reducing equivalents necessary to initiate electron transport through the mitochondrial ETC, a series of protein complexes that reside in the mitochondrial inner membrane (MIM; Balaban et al. 2005; Wolkow and Iser 2006). Two electrons donated from NADH + H + to complex I (NADH dehydrogenase) or from succinate to complex II (succinate dehydrogenase, SDH) are passed sequentially to the membrane-bound electron carrier, ubiquinone (coenzyme Q or CoQ) to give ubisemiquinone (CoQH•) and then ubiquinol (CoQH2). Ubiquinol transfers its electrons to complex III (ubiquinol: cytochrome c oxidoreductase), which transfers them to cytochrome c. From cytochrome c, the electrons flow to complex IV (cytochrome c oxidase, COX), which reduces molecular oxygen to water in the final step. Each of these electron transport chain (ETC) complexes incorporates multiple electron carriers. Complexes I, II, and III encompass several iron-sulfur (Fe-S) centers, whereas complexes III and IV encompass the b + c1 and a + a3 cyto- chromes, respectively. Electron transfer by complexes I, III and IV is coupled to proton transport across the MIM to the intermembrane space. Thus, electron transport through the ETC is coupled to the export of 2 (via Complex II) or 3 (via Complex I) protons into the mitochondrial intermembrane space. Superoxide production occurs at two major sites along the electron transport chain, complex I and complex III, because large changes in the potential energy of Fig. 11 Diagram showing the relationships of mitochondrial oxidative phosphorylation to energy production (ATP) and reactive oxygen species (ROS) production. Dashed lines indicate the flow of electrons donated from either NADH or FADH2 to complexes I–IV. As a result of electron transport, protons (H+) are translocated into the intermembrane space of the mitochondria creat- ing a proton gradient across the inner mitochondrial membrane. The proton gradient is necessary to drive ATP production via ATP synthase, but superoxide anions are produced (O 2 •− ) at sites I and III. Uncoupling proteins (UCP) reduce the overall mitochondrial proton gradient. The different complexes of oxidative phosporylation, designated I to IV and coded by color, are complex I (NADH: ubiquinone oxidoreductase) encompassing a flavin mononucleotide and six Fe-S centers; complex II (succinate: ubiquinone oxidoreductase) involving a flavin adenine dinucleotide, three Fe-S centers, and a cytochrome b; complex III (ubiquinol: cytochrome c oxidoreductase) encom- passing cytochromes b, c1 and the Rieske Fe-S center; complex IV (cytochrome c oxidase) encompassing cytochromes a + a3 and CuA and CuB; and the H+-translocating ATP synthase (F 1 and F 0 ) (Figure adapted from Balaban et al. 2005; Wolkow and Iser, 2006) I II II IV F 1 F 0 UCPCoQCyt C H + H + H + H + H + ADPATP O 2 H 2 O FADH2 FAD NADH NAD + Intermembrane space Mitochondrial inner membrane Matrix Cytoplasm O 2 - O 2 - O 2 - 95Age-Related Decline in Actomyosin Structure and Function the electrons, relative to the reduction of oxygen, occur. The relative contributions of complexes I and III to ROS production appear to be dependent on types of tissues, species and experimental conditions. The eight iron sulphur centers and/or the active site flavin mononucleotide are proposed to be the sites of ROS production at complex I (NADH coenzyme Q reductase) (Liu et al. 2002). The production of superoxide at complex I occurs at the matrix side of the inner membrane because the centers are proposed to be at this side of the membrane. The site of superoxide production at complex III is thought to be unstable ubisemiquinone molecules (St-Pierre et al. 2002). Superoxide produced by complex III is released to both the matrix and cytosolic sides of the mitochondria with about 80% going to the matrix and 20% going to the intermembrane space (Cadenas 2004). ROS production is substrate-, tissue-, cell-, and organelle-specific. Under physiological conditions, about 0.2% of the total oxygen consumption is directed to ROS generation (St Pierre et al. 2002). More importantly, the mitochondrial metabolic state can influence the rate of ROS production (Frisard and Ravussin 2006). When oxygen consumption is low and the potential energy for protons is high (state 4), complexes of the ETC are in reduced states and superoxide production is highest. ROS production is increased when the electron carriers in the initial steps of the ETC harbor excess electrons, i.e., remain reduced, which can result from either inhibition of OXPHOS. Electrons residing in the electron carriers; for example, the unpaired electron of ubisemiquinone bound to the CoQ binding sites of complexes I, II, and III; can be donated directly to O 2 to generate superoxide anion. 6.3 Oxidative Stress/Damage to Macromolecules by ROS In general, there is a balance between free radical production and the many antioxidant defense mechanisms within the skeletal muscle fibers. Thus, ‘oxidative stress’ can be viewed as a disturbance in the prooxidant-antioxidant balance in favor of prooxidant, leading to oxidative damage (Fig. 12). Increased levels of ROS can directly or indirectly damage macromolecules such as phospholipids, nucleic acids, and proteins. Since ROS are generated in the mitochondria, they can damage mitochondrial macromolecules either at or near the site of their formation. Mitochondria have two membranes, an inner highly proteinaceous membrane (80% protein) and an outer, porous membrane. Some proteins are attached loosely to the surface of the membranes whereas others are integral parts of the membrane (embedded). ROS damage to proteins as a direct result of oxidative stress or as a consequence of lipid peroxidation can result in protein cross-linking, degradation of proteins and loss of function because of the close physical association of phospholipids and proteins in mitochondrial membranes. ROS can damage other macromolecules, outside the mitochondria, yet within the muscle fiber. Figure 12 is schematic of the sarcomere showing how increased damage to actin and myosin may potentially interrupt actomyosin interactions which could result in skeletal muscle contractility deterioration. . assessment of age-related changes in the interaction between muscle myosin and actin requires specific measurements of biochemical and structural properties of actin and myosin. Purified actin and myosin. 2002) 9 3Age-Related Decline in Actomyosin Structure and Function 6 Oxidative Stress Age-related deterioration of muscle function may involve ‘damage’ of muscle proteins by reactive oxygen and nitrogen. and myosin (43–50%) (Ingalls et al. 1998; Yates and Greaser 1983). However, since the interaction between actin and myosin in the muscle filament lattice also depends on other structural and