IN a comprehensive review in 1987, Rusy and Komai1discussed the possible mechanisms by which volatile anesthetics inhibit cardiac contraction. Since that time, there have been advancements in the understanding of excitation-contraction coupling, cardiac mechanics, and the actions of volatile anesthetics. There are essentially three major factors that determine the force of contraction of heart muscle cells: the magnitude of cytosolic Ca2+increase after electrical excitation, the responsiveness of the contractile proteins to Ca2+, and the sarcomere length (SL) at which the contractile proteins are activated. Hence, there are two possible ultimate direct negative inotropic actions of volatile anesthetics: a reduction in Ca2+availability or a decrease in the Ca2+-responsiveness (Ca2+-sensitivity or maximal Ca2+-activated force) of the contractile apparatus. The rate of relaxation of the muscle cells, on the other hand, depends on the rate at which Ca2+is cleared from the cytosol, which facilitates its dissociation from the regulatory proteins of the contractile system. The control of calcium cycling and the activity of the contractile proteins consume energy that must be continuously supplied by the mitochondria, another potential site volatile of anesthetic action.

A diagram of cardiac excitation-contraction coupling is shown in figure 1. Action potentials, initiated by sinoatrial nodal cells, are rapidly conducted throughout the heart, facilitated by the His-Purkinje system, and activate the force (pressure) generating myocardial cells within the ventricles. Rapid depolarization in myocytes is mediated by voltage-gated Na+channels, different subtypes of which are located in the transverse tubules (Nav1.1, Nav1.3, and Nav1.6) and intercalated disks (Nav1.5).2The depolarization of the cell leads to the activation of L-type Ca2+channels, which are primarily encoded by the α1Cgene (Cav1.2),3and are the next key element in cardiac excitation-contraction coupling. In addition, other ionic currents, such as that attributable to Na/Ca exchange, as well as chloride and potassium currents, all shape the action potential, whose duration is ∼300 ms. The action potential spreads from cell to cell, a process that is facilitated by Nav1.5 channels (in the intercalated disks) and extensive gap junctions between cells, each of which is connected to ∼15 of its neighbors.4In ventricular cells, the action potential passes into transverse tubules (t-tubules) that serve to minimize delay in excitation throughout the cell.5 

Fig. 1. Excitation-contraction coupling in a ventricular myocyte. Excitation of the sarcolemma and t-tubules by an action potential leads to activation of L-type Ca2+channels and Ca2+-induced Ca2+release. The resulting transient elevation of cytosolic Ca2+serves to activate the contractile apparatus, producing contraction. Removal of Ca2+from the cytosol by various Ca2+transporters facilitates mechanical relaxation. 

Fig. 1. Excitation-contraction coupling in a ventricular myocyte. Excitation of the sarcolemma and t-tubules by an action potential leads to activation of L-type Ca2+channels and Ca2+-induced Ca2+release. The resulting transient elevation of cytosolic Ca2+serves to activate the contractile apparatus, producing contraction. Removal of Ca2+from the cytosol by various Ca2+transporters facilitates mechanical relaxation. 

Surface membrane depolarization promotes the influx of Ca2+via  the voltage-gated L-type (also called dihydropyridine-sensitive) Ca2+channels and possibly the Na/Ca exchanger (NCX). In the case of the L-type Ca2+channel, Ca2+influx is limited by both Ca2+-dependent inactivation as well as voltage-dependent and time-dependent inactivation.6–9Ca2+-dependent inactivation depends on both the Ca2+that enters the cell via  the channel itself (e.g. , Bechem and Pott10) and Ca2+release from intracellular stores.11,12The intracellular mechanism of Ca2+-dependent inactivation appears to involve calmodulin.13,14 

Although Ca2+influx via  L-type calcium channels will increase intracellular Ca2+directly by a small amount, the influx of Ca2+is normally amplified by a larger release of Ca2+from the sarcoplasmic reticulum (SR) in a process known as Ca2+-induced Ca2+release (CICR).15This mechanism resides in the Ca2+-dependent gating of ryanodine-sensitive Ca2+-release channels in the SR (ryanodine receptors; RyRs) and of the three isoforms known, the RyR2 isoform predominates in heart (for review of junctional proteins, see Muller et al.  16). Because activation of RyRs depends only on an elevation in intracellular Ca2+, any source of Ca2+could, in principle, activate CICR.

Alternative Trigger of CICR

The NCX, which serves as the primary route for Ca2+extrusion at rest,17,18can be reversed during depolarization as a result of the action potential to bring Ca2+into the cell. This Na/Ca exchange “reverse mode” Ca2+influx has also been implicated in triggering CICR.19–21Although this possibility was first demonstrated under pathologic conditions of calcium overload, where the cell is highly sensitive to trigger calcium concentrations,19subsequent experiments showed that conditions which would lead to NCX reversal (such as strong depolarization or increased intracellular Na+concentrations) could also trigger CICR.20–22It has even been suggested that the NCX might be a major source of trigger Ca2+during the normal action potential,21although this has been debated.23Blockade of NCX with an inhibitory peptide suggested that only up to 27% of the trigger might be attributable to NCX.22In addition to the membrane potential, internal Na+concentrations will also be critical for determining the NCX contribution.24During the upstroke of the action potential, Na+influx might increase Na+concentrations locally to promote reverse mode exchange20,25as well modify SR Ca2+content.26Some researchers have been unable to demonstrate potent triggering of CICR at negative potentials by reverse mode exchange when the Na+current is activated,24,27and it is likely that a major part of this controversy resides in the difficulty of obtaining good voltage control during the Na+current while also trying to prevent all Ca2+influx via  L-type Ca2+channels by pharmacological agents.

Colocalization of Dihydropyridine Receptors and RyRs

The reliability of excitation-contraction coupling is also dependent on the physical relation between the proteins that provide/regulate the triggers for CICR as well as the location of the RyRs that respond to the trigger signal. The membranes of the t-system and the terminal cisternae of the SR are closely opposed in junctional areas or “dyads,”28and such structures will increase the reliability of CICR by limiting diffusional loss of the trigger Ca2+signal within them.29–31Although the dihydropyridine receptors (L-type Ca2+channels) and the RyRs (Ca2+-release channels) are colocalized,32,33a combination of immunofluorescence and data deconvolution techniques have led to the suggestion34that neither voltage-gated Na+channels nor the NCX are located in the dyad. On the contrary, more recent immunolocalization data from Thomas et al.  35suggest that the NCX is indeed concentrated in t-tubules.

Elementary Ca2+Release Events: Ca2+Sparks

Over the past decade new insights into the basic events underlying CICR have been obtained by application of confocal microscopy and fluorescent calcium indicators. Such experiments have shown that the close opposition of the dihydropyridine receptors and the RyRs enables local control of Ca2+release into the cytosol as proposed by Stern.36In this model, it is not the global cytosolic concentrations of Ca2+(or any other messenger) that are important but rather the micro-environment around the RyR. The importance of local rather than global signals is based on the idea that most biologic processes are inherently nonlinear. Thus, in terms of cardiac excitation-contraction coupling, we must consider the microscopic environment produced by the local activation of trigger mechanisms (such as L-type Ca2+channels) and the behavior of a local cluster of RyRs in their vicinity. The discovery of “Ca2+sparks” or microscopic release events resulting from the local coactivation of a cluster of RyRs5,37,38has reinforced this idea. Ca2+sparks are seen both as spontaneous and evoked microscopic SR release events inside the cell. The latter generally require the use of Ca2+channel antagonists to reduce the probability of spark activation so that individual sparks can be observed.37,39From the site of release (near z-lines) the Ca2+spark diffuses to cover a region of the cell approximately 2 μm in diameter5with a slightly greater spread in the longitudinal direction40(fig. 2). When measured at room temperature, the Ca2+spark reaches a peak in ∼10 ms and lasts ∼40–80 ms and peak Ca2+during the spark is typically 200–400 nm. The decay of the Ca2+spark is a result of both Ca2+diffusion and SR uptake.41When sparks are activated by depolarization, they summate to produce a larger and slower Ca2+transient, and the reduced rate of decline of Ca2+can be ascribed to the loss of the diffusive component of spark decay; during the whole-cell Ca2+transient Ca2+is globally increased so diffusion cannot serve to reduce Ca2+.5 

Fig. 2. Elementary Ca2+release events (Ca2+sparks) in a ventricular myocyte. The top panel shows confocal images (x,y scans) of a myocyte obtained using a confocal laser scanning microscope. The local regions of increased fluo-3 fluorescence are Ca2+sparks. The middle panel shows a Ca2+spark recorded using the line-scan (x,t) mode of imaging. 

Fig. 2. Elementary Ca2+release events (Ca2+sparks) in a ventricular myocyte. The top panel shows confocal images (x,y scans) of a myocyte obtained using a confocal laser scanning microscope. The local regions of increased fluo-3 fluorescence are Ca2+sparks. The middle panel shows a Ca2+spark recorded using the line-scan (x,t) mode of imaging. 

From studying Ca2+sparks we now know that excitation-contraction coupling in the heart is attributable to the spatio-temporal summation of a very large number of “elementary” Ca2+sparks.37,42,43It has been estimated that during excitation-contraction coupling the spontaneous spark rate is increased by a factor of ∼10,000 by a “local” 100-fold increase in Ca2+resulting from the trigger Ca2+influx.44Such a local increase in Ca2+is consistent with some computer simulations of changes in Ca2+attributable to L-type Ca2+channels.31Moreover, the latency for SR Ca2+release is <2 ms,45consistent with the rate of RyR opening expected from such high local trigger Ca2+concentrations.30Although strictly speaking, Ca2+sparks cannot be elemental if attributable to the concerted activation of a number of local RyRs in a cluster,40the fact that evoked Ca2+sparks appear to have a modal amplitude distribution at fixed locations within the cell46suggests that each spark site behaves in an “all or none” fashion and are therefore “elementary” in a functional sense (but see Lipp and Niggli47). In any case, it is generally agreed that coordinated activation of multiple spark sites gives rise to the global increase in cytosolic [Ca2+] that occurs under normal conditions. More recently, Wang et al.  48have shown that the local increase in Ca2+concentration (“sparklet”) produced by an extended (drug modified) opening of a single L-type Ca2+channel can trigger a cluster of about 4–6 peripheral RyRs, which then produce a spark. On the other hand, from both experimental and theoretical approaches, both Bridge et al.  46and Soeller and Cannell49suggest that sparks may in fact arise from somewhat larger clusters of RyRs, consisting of >15 receptors. The number of RyRs underlying a Ca2+spark is important because it sets limits on the degree to which different amounts of Ca2+may be released by modulation of RyR open probability as well as the “safety factor” inherent in RyR activation.

SR Ca2+Content

The amount of Ca2+released into the cytosol depends not only on the magnitude of the Ca2+-influx current (the trigger for Ca2+release) but also on the SR Ca2+content.50,51For a given concentration of trigger Ca2+, the Ca2+release increases as a function of SR Ca2+content. High SR luminal [Ca2+] appears to increase the open probability of the RyRs,52,53which may explain why transient inward currents, afterdepolarizations, and aftercontractions are often seen when the SR contains a high Ca2+load. Under steady state conditions (constant heart rate and constant neurohumoral input), the magnitudes of Ca2+influx and efflux across the sarcolemma are balanced, and thus there is no net change in mean SR Ca2+content. After β1-receptor stimulation, for example, L-type Ca2+current as well as SR Ca2+-pump activity increases, resulting in net Ca2+influx over successive contraction cycles that loads Ca2+into the SR. The increase in Ca2+current (trigger Ca2+) and SR Ca2+store induced by β1-receptor stimulation gives rise to larger Ca2+transients and therefore more contractile protein activation. However, it is still unclear whether this is accompanied by an increase in the number of Ca2+sparks as well as individual spark amplitude.

Ca2+Transport Systems

The increase in cytosolic [Ca2+] and the accompanying force development after electrical activation is transient because Ca2+is rapidly removed from the cytosol after release. There are four main Ca2+transport systems that remove Ca2+from the cytosol: the SR Ca2+-pump (sarco-endoplasmic reticular Ca2+-adenosine triphosphate [ATP]ase), the sarcolemmal Ca2+-pump (Ca2+-ATPase), the NCX, and the mitochondrial Ca2+-uniporter (fig. 1). Although the relative contributions of these systems is species-dependent, the predominant Ca2+transport systems are the SR Ca2+-ATPase and the NCX.50,54,55The activity of the SR Ca2+-ATPase can be increased by cAMP-dependent phosphorylation of the endogenous inhibitor phospholamban,56whereas the activity of the NCX can be modulated by phosphorylation via  protein kinase C.57The NCX exchanges 3 Na+for 1 Ca2+, and therefore it is electrogenic. Hence, its direction (Ca2+-influx versus  Ca2+-efflux mode) is determined by the prevailing transmembrane gradients for Na+and Ca2+, as well as membrane potential.58,59Indeed, the NCX may transiently reverse after membrane depolarization, especially during the peak of the action potential. It should be reiterated that the magnitude and direction of the NCX is strongly (in a cubic fashion) dependent on intracellular [Na+], as discussed in detail by Cooper et al.  58(see also Evans and Cannell24). Hence, the activity of the Na pump (Na+,K+-ATPase) plays a critical role in determining the Gibbs free energy of the exchanger. It should also be noted that activation of the exchanger after spontaneous Ca2+release after the Ca2+transient (in Ca2+overloaded cells60or in nonuniform cardiac muscle61) will lead to current flow across the membrane which can lead to after depolarizations sufficiently large to produce action potentials and extrasystoles.

Activation of the Contractile System

Activation of the contractile system by Ca2+is mediated by its binding to the regulatory protein troponin C (TnC).62The contractile proteins (myofibrils), which occupy around 60% of the cell volume,63,64consist of thick (myosin) and thin (actin) filaments which are arranged between z-lines to form sarcomeres, the repeating units of the myofibrils (fig. 1). The thin filaments are ∼1 μm in length and protrude from anchoring points on the z-line.65,66Tropomyosin lies in the groove between the two actin chains of the thin filament and its movements are regulated by the Ca2+-sensitive troponin complex.67Interdigitating with the thin filaments are the thick filaments, the globular heads of which form the cross-bridges which interact with the thin filament and contain actomyosin-ATPase.

The troponin complex has three subunits: TnC (the Ca2+-binding subunit), troponin T (the tropomyosin-binding subunit), and troponin I (the inhibitory subunit). Compared with its skeletal counterpart, the cardiac troponin I isoform has a longer amino terminus containing two phosphorylation sites that are substrates for cAMP-dependent protein kinase A.68When troponin I is bisphosphorylated, the Ca2+off-rate of TnC is increased, which enhances the relaxation rate of a contraction.68,69Cardiac TnC has three Ca2+-binding sites, unlike the skeletal isoform, which has four.56,70Two of the sites bind both Ca2+and Mg2+, whereas the other site specifically binds Ca2+and therefore serves as the regulatory site. In resting cardiac muscle, cytosolic [Ca2+] is low (∼70 nm)71and the Ca2+-specific site of TnC (with a dissociation constant of ∼500 nm)55is unoccupied.72In this state, tropomyosin, lying in the groove of the thin filament, prevents interaction of the myosin heads (cross-bridges) with actin (but see Perry73). When cytosolic [Ca2+] increases, after membrane depolarization and CICR, Ca2+binds to TnC, which causes tropomyosin to move out of the actin groove, thereby allowing myosin to interact with actin, producing force or shortening (for review, see Perry73).

The mechanism by which myosin interacts with actin and uses the energy from ATP hydrolysis to produce mechanical work is known as the “cross-bridge theory of sliding filaments” and was first proposed in 1957 by Huxley.74This mechanism has been extensively reviewed.70,75As long as Ca2+is bound to TnC, the myosin head can form cross-bridges with nearby binding sites on the thin filament. The generation of force or the relative sliding of the filaments is thought to be brought about by rotation of the head, the swinging cross-bridge model.76,77The biochemical steps that drive cross-bridge cycling (hydrolysis of ATP via  actomyosin-ATPase) have been largely characterized.78Basically, ATP binds to myosin, causing the globular head domain to detach from actin, and its subsequent hydrolysis sets the myosin head into a high energy state. Ishijima et al.  79have simultaneously measured the mechanical events and actomyosin-ATPase activity of a single-headed myosin molecule interacting with an actin filament. Under these artificial conditions, force generation did not always coincide with the release of adenosine diphosphate from the myosin head (as previously thought) but could occur several hundred milliseconds after release. The authors proposed that the mechanical and biochemical events of cross-bridge cycling are not tightly coupled in that a myosin head can undergo several conformational changes during a single actomyosin-ATPase cycle. However, it should be stressed that the exact nanomechanics and stoichiometry of cross-bridge cycling still remain to be elucidated.80 

Responsiveness of the Contractile Proteins to Ca2+

Although intracellular [Ca2+] is the major determinant of force at any given SL, the development of force is also dependent on the responsiveness of the contractile proteins to Ca2+. Responsiveness refers to the relation between force and intracellular [Ca2+] (fig. 3), often expressed as pCa (−log10[Ca2+]). The pCa-force relation can be modulated by various factors such as ionic strength, temperature, pH, [Pi], SL (fig. 3), and phosphorylation state of the contractile apparatus.56The [Ca2+] at which force is half-maximal provides an index of ‘Ca2+-sensitivity' of the contractile system whereas the force obtained at TnC-saturating [Ca2+] gives “maximal Ca2+-activated force.” Ca2+binding to TnC is cooperative, that is, the binding of one Ca2+ion facilitates the binding of the next one. This gives rise to a steep pCa-force relation.

Fig. 3. Force-[Ca2+] relations of trabeculae at various sarcomere lengths (SLs). Note that there is a shift of the [Ca2+]-activation curve to the left, indicating that Ca2+-sensitivity of the contractile proteins increases as a function of SL. Adapted with permission from Kentish  et al. 84 

Fig. 3. Force-[Ca2+] relations of trabeculae at various sarcomere lengths (SLs). Note that there is a shift of the [Ca2+]-activation curve to the left, indicating that Ca2+-sensitivity of the contractile proteins increases as a function of SL. Adapted with permission from Kentish  et al. 84 

The relation between intracellular [Ca2+] and force has been well characterized using chemically or mechanically “skinned” muscle preparations. Skinning renders the sarcolemma freely permeable to ions and small molecules, and thus this technique allows the intracellular environment to be well controlled. However, Marban et al.  81showed that the relation between intracellular [Ca2+] and force is strikingly steeper in intact cardiac muscle than is reported for skinned muscle preparations. This apparent difference was reexamined by Gao et al. ,82who determined the intracellular [Ca2+]-force relation before and after skinning in the same muscle preparation (rat cardiac trabecula). The [Ca2+] required for half-maximal force was considerably higher after skinning, suggesting that the skinning procedure may damage or remove components involved in the regulation of actin-myosin interaction. Hence, data obtained from experiments employing skinned muscle preparations cannot be readily extrapolated to the intact system.

Length-dependence of Passive and Ca2+-activated Force

One of the major determinants of force is the SL at which the contractile proteins are activated83,84(fig. 3).Typically, cardiac myocytes have a slack (resting) length of ∼1.9 μm and can be extended to SLs of up to ∼2.3–2.4 μm. In intact cardiac muscle, twitch force (or stress, which is force/cross-sectional area) increases steeply as a function of SL (the Frank-Starling mechanism). This steep increase in force as a function of SL is thought to be attributable largely to a length-dependent increase in the sensitivity of the contractile system to Ca2+, mediated by increased affinity of Ca2+binding to TnC.62,72An increase in overlap of thick and thin myofilaments, a reduction in interfilament lattice spacing,62,85and an increase in intracellular Ca2+release86,87are thought to contribute to the increase in force when SL is increased.

The exact mechanism by which an increase in SL causes an increase in myofilament Ca2+responsiveness remains elusive. Fuchs and Smith72argue that the sensitivity of the contractile system to Ca2+is governed by myofilament lattice spacing rather than muscle length. It has been shown by radiographic diffraction that interfilament spacing decreases as SL increases in both intact and skinned cardiac muscle.88Fitzsimons and Moss85previously proposed that decreased interfilament spacing increases force development by increasing the probability that myosin will form strong cross-bridges with actin. Recent work from Konhilas et al. ,89suggests that changes in lattice spacing may not underlie length-dependent activation. These authors found that osmotic compression of the myofilament lattice spacing, which was measured by radiographic diffraction, did not alter Ca2+-sensitivity in skinned cardiac trabeculae. Hence, the sensor for SL changes probably does not reside in the interfilament space. The protein titin (also called connectin), which spans the entire sarcomere and interacts with the thick and thin filaments,90is in a good position to sense SL changes, but whether this protein signals length changes to the regulatory proteins of the thin filament is not known.91,92 

Extension of cardiac muscle beyond a SL that optimizes actin-myosin interaction (a SL of 2.2–2.4 or so, depending on species), is prevented by the development of large parallel elastic forces,84,93,94as shown in figs. 4A and 4B. The straightening of perimysial collagen fibers, which are thought to be the dominant contributors to passive force at SL >2.2 μm,95is presumed to underlie this end-stop effect (figs. 4C and 4D).

Fig. 4. Passive force-SL relations of rat cardiac trabeculae obtained after  (A ) and before  (B ) skinning. Note that passive force at SLs above 2.2 μm is reduced after skinning. (Modified from Kentish  et al. 84,with permission) At near-resting SL, the perimysial collagen fibers (reconstructed in 3-D by confocal laser scanning microscopy) of cardiac trabeculae are wavy  (C ), whereas they become straight at SLs of approximately 2.3 μm  (D ), corresponding to the steep portion of the force-SL relation. Adapted with permission from Hanley  et al. 200 

Fig. 4. Passive force-SL relations of rat cardiac trabeculae obtained after  (A ) and before  (B ) skinning. Note that passive force at SLs above 2.2 μm is reduced after skinning. (Modified from Kentish  et al. 84,with permission) At near-resting SL, the perimysial collagen fibers (reconstructed in 3-D by confocal laser scanning microscopy) of cardiac trabeculae are wavy  (C ), whereas they become straight at SLs of approximately 2.3 μm  (D ), corresponding to the steep portion of the force-SL relation. Adapted with permission from Hanley  et al. 200 

At SLs < 2.1 μm, where passive force is less than 5% of maximal twitch force, titin overshadows collagen as the main contributor (>90%) to passive force.95This large protein, sometimes referred to as the third filament of the contractile apparatus, is closely associated with the thick filaments in the A-band region.96,97The elasticity of titin resides in its I-band portion, which spans from the z-line to the tip of the myosin filament.98This elastic (I-band) segment of titin is shorter in cardiac muscle than in skeletal muscle and may explain why titin-based passive force is greater in cardiac than most skeletal muscle at a given SL. In addition to acting as a molecular spring, titin may also contribute to the viscoelastic properties of the sarcomeres by interacting with the thin filament.71,98,99 

Cardiac Energy Metabolism and Mitochondrial Function

Cardiac myocytes, the functional unit of the heart (accounting for >90% of heart volume), require a continual supply of free energy to perform mechanical work, synthesize various molecules, and do electrochemical work (maintain ion gradients across membranes). This free energy comes from the oxidation of metabolic substrates such as long-chain fatty acids and, to a lesser extent, glucose and lactate.100–102Mitochondria occupy ∼30% of the myocyte volume4,63,64and have a highly ordered distribution in the living myocyte, lying in the interfibrillar space between z-lines (fig. 5). Oxygen is the ultimate acceptor of electrons when metabolic substrate is oxidized. Electrons are transferred to oxygen via  the electron transport chain, a series of proton pumps and electron carriers located in the mitochondrial inner membrane. A proton gradient is established across the mitochondrial inner membrane, giving rise to a potential of ∼200 mV, as electrons are transferred along the various complexes of the electron transport chain. The energy stored in this electrochemical gradient is used to drive the synthesis of the energy carrier molecule ATP. Mitochondrial matrix ATP is then tranported via  the ATP-adenosine diphosphate translocase to the cytosol where, via  the creatine-creatine phosphate shuttle system,103it serves as a readily accessible source of free energy for mechanical and electrochemical work.

Fig. 5. Confocal fluorescence image of mitochondria in a living, intact ventricular myocyte of guinea pig (  A ) (P. J. Hanley, unpublished data). Mitochondria (and a nucleus) were labeled with the fluorescent indicator rhod-2. Note the highly ordered distribution of mitochondria inside the cell. (  B ) Diagram showing that the mitochondria are spatially arranged within the intermyofibrillar spaces between the t-tubules. 

Fig. 5. Confocal fluorescence image of mitochondria in a living, intact ventricular myocyte of guinea pig (  A ) (P. J. Hanley, unpublished data). Mitochondria (and a nucleus) were labeled with the fluorescent indicator rhod-2. Note the highly ordered distribution of mitochondria inside the cell. (  B ) Diagram showing that the mitochondria are spatially arranged within the intermyofibrillar spaces between the t-tubules. 

Ultimately, all the energy used by the myocyte to maintain its structure and function is degraded to heat. From studies in which the rate of heat production of cardiac trabeculae of guinea pig was measured under various conditions, Schramm et al.  104deduced that actomyosin-ATPase, Ca2+-ATPase, and Na+,K+-ATPase accounted for 76%, 15%, and 9%, respectively, of the overall rate of ATP turnover. In accord with these results, Ebus and Stienen105measured the rate of ATP hydrolysis in saponin-skinned trabeculae of rat and showed that approximately 15% of maximal Ca2+-activated ATPase was membrane-bound, around two thirds of which was attributable to the sarco-endoplasmic reticular Ca2+-ATPase. The source of the heart's extraordinarily high basal rate of metabolism, which accounts for 25–30% of the energy expenditure of the beating heart, remains unknown (for review, see Gibbs and Loiselle106).

Mitochondrial Ca2+Uptake

Mitochondrial Ca2+influx occurs via  a ruthenium-red sensitive Ca2+-uniporter107(fig. 1), whereas its efflux is mediated by a Na/Ca exchange mechanism (and, indirectly, Na+-H+exchange), the stoichiometry of which has not been established.108Ca2+influx into the mitochondria is thought to regulate metabolism because an increase in mitochondrial [Ca2+] stimulates ATP synthase109and the activity of pyruvate dehydrogenase and α-ketoglutarate dehydrogenase, enzymes linked with the tricarboxylic acid cycle.110 

Recent evidence suggests that cytosolic Ca2+signals are probably communicated directly to the mitochondria. After selectively loading the fluorescent Ca2+indicator rhod-2 into the mitochondria of electrically stimulated ventricular myocytes of rabbit, Trollinger et al.  111were able to demonstrate mitochondrial Ca2+transients using confocal microscopy. In accord, in work with patch-clamped ferret or cat ventricular myocytes, Zhou et al.  112were also able to detect mitochondrial Ca2+transients during a twitch, albeit only when the cytosolic resting [Ca2+] exceeded physiologic concentrations. Compared with the cytosolic Ca2+transient, the kinetics of the mitochondrial Ca2+transient were much slower. Griffiths,113using cardiac myocytes and techniques to load indo-1 (a fluorescent Ca2+indicator) selectively into mitochondria, found that mitochondrial transients accompanied twitches in guinea pig but not rat, suggesting that there may be species-differences in mitochondrial Ca2+cycling. It should be noted that the surface area of the mitochondrial inner membrane is 10-fold higher than the area of the sarcolemma and t-tubule system.56Hence, small changes in mitochondrial Ca2+permeability could greatly influence Ca2+distribution in the cell.

Neurohumoral Regulation of Contraction

Although neurohumoral regulation is outside the scope of this review (see, for example, Morris and Malbon114or Xiao115), a brief overview of the major signaling pathways in the myocyte follows. Each myocyte has three major intracellular signaling cascades that are modulated via  G protein-coupled surface receptors, each of which contains a conserved structure of seven transmembrane α-helices.116,117The G protein-coupled receptors regulate the activity of various membrane-bound enzymes including adenylate cyclase, guanylate cyclase, and phospholipase C, which produce, respectively, the secondary messengers cAMP, cGMP, and DAG.118In each case, the secondary messengers activate protein kinases (protein kinase A, protein kinase G, and protein kinase C, respectively) that phosphorylate specific amino acid residues on contractile proteins, ion channels, and pumps. The adrenergic receptors and the muscarinic cholinergic receptors are the most important G protein-coupled receptors in the heart. At least nine subtypes of adrenergic receptors have been cloned: three α1-receptors, three α2-receptors, and three β-receptors. When agonist is bound to the receptor the G proteins dissociate into effector subunits that modulate the activity of various membrane-bound targets such as adenylate cyclases, phospholipases, and ion channels. For example, when agonists bind to β1-receptors, the Gαs subunit dissociates and stimulates adenylate cyclase, which produces the secondary messenger cAMP, a protein kinase A activator. Protein kinase A phosphorylates the following: L-type Ca2+channels (which increases Ca2+influx), phospholamban (which enhances SR Ca2+-uptake, thereby increasing the rate of relaxation), RyRs (which may facilitate Ca2+release, a controversial issue),119troponin I (which reduces filament sensitivity to Ca2+ions and by itself decreases force but accelerates relaxation), and myosin binding protein C. The net effect is positive inotropy and positive lusitropy (increased rate of relaxation). Stimulation of muscarinic receptors yields Gαi, which inhibits adenylate cyclase, producing the opposite effect to adrenergic receptor stimulation.

Volatile anesthetics have been shown to inhibit or stimulate various cellular components such as ion channels, pumps, exchangers, enzymes, gap junctions, and components of the contractile system, as schematically illustrated in figure 6. Most studies have shown effects at moderately high concentrations, suggesting that volatile anesthetics may perturb lipid bilayers, which would explain why these lipophilic agents affect diverse molecular targets. However, the fundamental molecular mechanisms by which these agents inhibit or stimulate various membrane-bound proteins and the contractile apparatus remains unclear. For a recent discussion of the possible mechanisms by which molecules of volatile anesthetics interact with protein structures, lipid bilayers and various molecular interfaces, see a review by Urban.120 

Fig. 6. Sites of action of volatile anesthetics in a ventricular myocyte. The red spots indicate inhibitory actions whereas the green spots indicate stimulatory actions. 

Fig. 6. Sites of action of volatile anesthetics in a ventricular myocyte. The red spots indicate inhibitory actions whereas the green spots indicate stimulatory actions. 

Anesthetic concentrations are usually expressed as mM, volume%, or their equivalent MAC (minimal alveolar [anesthetic] concentration) value, where 1 MAC is defined as the minimal alveolar (anesthetic) concentration at one atmospheric ambient pressure required to prevent movement in response to a noxious stimulus in 50% of animals.121Anesthetic concentrations corresponding to 1 MAC (in rat) at 37°C, for example, are as follows: halothane (0.27 mm), isoflurane (0.31 mm), and sevoflurane (0.35 mm).122When comparing studies performed at different temperatures, it is important to note that MAC expressed as volume% varies considerably with temperature, whereas the equivalent liquid-phase concentration, typically expressed in mM, changes little.122 

Ultimately, for volatile anesthetics to inhibit cardiac contractility they must reduce Ca2+-availability or decrease the Ca2+-responsiveness of the contractile proteins. Although direct actions at the level of the contractile apparatus or components of excitation-contraction coupling are likely to be more important, volatile anesthetics could also inhibit contractile function indirectly by impairing mitochondrial energy supply.

Anesthetic Effects on Ca2+Availability

In 1986, Bosnjak and Kampine123showed that halothane decreased the Ca2+transient in cat papillary muscles that had been microinjected with aequorin. These authors also reported that halothane did not affect resting Ca2+concentrations; however, the luminescence of aequorin is a quadratic function of [Ca2+] and, therefore, is not a suitable indicator for measuring resting Ca2+concentrations.124Another potential limitation of aequorin is that its light-emitting properties may be altered by direct interaction of the anesthetic with the photoprotein,125although, in favor of aequorin, Housmans and Wanek126recently reported that neither halothane nor isoflurane affected aequorin luminescence in the pCa range 2–8. In 1985, the ratiometric fluorescent Ca2+indicators fura-2 and indo-1 were introduced,127followed by fluo-3;128these are technically less difficult to use than aequorin.

A large number of studies using fluorescent Ca2+indicators have corroborated the observations of Bosnjak and Kampine123that volatile anesthetics dose-dependently decrease the amplitude of the intracellular Ca2+transient in intact cardiac muscle.129–136Hence, it is now well established that volatile anesthetics indeed decrease the amount of Ca2+released into the cytosol after electrical stimulation. Figure 7shows an example of the inhibitory effect of halothane on peak twitch intracellular [Ca2+], indexed as fura-2 fluorescence ratio. Note that the anesthetic had no effect on diastolic [Ca2+]. Moreover, in multicellular cardiac muscle preparations, it has also been shown that, after correcting fluorescent indicator signals for autofluorescence changes, volatile anesthetics have no effect on diastolic (resting) Ca2+concentrations.129–131We now turn our attention to the anesthetic actions that may be responsible for the decrease in the cytosolic Ca2+transient.

Fig. 7. Fura-2 fluorescence (top trace), an index of [Ca2+], and cell length (bottom trace), an index of contraction, were measured simultaneously in an electrically stimulated rat ventricular myocyte. Application of halothane initially induced a transient increase in the Ca2+transient and twitch force before both the Ca2+signals and contraction decreased. Reproduced with permission from Prof. M. R. Boyett and the  Journal of Physiology. 

Fig. 7. Fura-2 fluorescence (top trace), an index of [Ca2+], and cell length (bottom trace), an index of contraction, were measured simultaneously in an electrically stimulated rat ventricular myocyte. Application of halothane initially induced a transient increase in the Ca2+transient and twitch force before both the Ca2+signals and contraction decreased. Reproduced with permission from Prof. M. R. Boyett and the  Journal of Physiology. 

Anesthetic-induced Inhibition of L-type Ca2+Current and Shortening of the Action Potential

Studies using the single microelectrode voltage-clamp technique137–139suggested that volatile anesthetics inhibit L-type Ca2+channels. Indeed, using the patch-clamp technique, Bosnjak et al.  140showed that halothane, enflurane, and isoflurane reversibly decreased whole-cell Ca2+current (which is predominantly L-type Ca2+current) in canine ventricular myocytes. In whole-cell and cell-attached recordings using a similar preparation (rat myocytes), Pancrazio141confirmed and extended this work. Halothane (0.9 mm) and isoflurane (0.8 mm) decreased the peak whole-cell Ca2+current by ∼40% and ∼20%, respectively (fig. 8). In cell-attached recordings, the anesthetics decreased both mean open time and open probability without affecting single-channel conductance. The anesthetics also enhanced the slow component of inactivation without affecting the fast component. Inhibition of L-type Ca2+current would lead to a decrease in SR Ca2+content (an important determinant of contractility) over subsequent beats.

Fig. 8. Effect of halothane (  A ) and isoflurane (  B ) on whole-cell Ca2+current (  I Ca), measured in guinea pig ventricular mycoytes. Both anesthetics (recordings indicated by filled circles) decreased peak  I Caand enhanced the apparent rate of channel inactivation. Reproduced with permission from Pancrazio.  141 

Fig. 8. Effect of halothane (  A ) and isoflurane (  B ) on whole-cell Ca2+current (  I Ca), measured in guinea pig ventricular mycoytes. Both anesthetics (recordings indicated by filled circles) decreased peak  I Caand enhanced the apparent rate of channel inactivation. Reproduced with permission from Pancrazio.  141 

In addition to exerting a negative inotropic effect, inhibition of L-type Ca2+current should also manifest as shortening of the action potential. Indeed, Lynch et al.  137found that halothane (>2%) shortened the action potential, measured in guinea pig papillary muscle. Subsequent studies, using enflurane and isoflurane in addition to halothane, have confirmed the observations of Lynch et al.  137that anesthetics shorten the action potential.138,139,142,143Rithalia et al.  144showed that action potential duration was decreased to a greater extent in myocytes isolated from the endocardium than those isolated from the epicardium of rat heart. These authors speculated that this difference might account for the greater negative inotropic effect of halothane on the subendocardium. At lower halothane concentrations, which did not shorten action potential duration, negative inotropy was still observed by Lynch et al. ,137suggesting that additional actions underlie the negative inotropic actions of volatile anesthetics. In accord with Lynch et al. ,137Harrison et al.  132showed that the negative inotropic effect of halothane on rat ventricular myocytes was similar even when action potential duration was maintained constant by applying voltage clamp.

Inhibitory and Stimulatory Effects of Anesthetics on K+Channels

Halothane and isoflurane,145as well as sevoflurane,146have been shown to inhibit inwardly rectifying K+(Kir) channels in guinea pig ventricular myocytes; that is, at potentials positive to the equilibrium potential for K+, the anesthetics increased outward current whereas at more negative potentials inward current was decreased. Transient outward K+current (I  to), important in the early phase of action potential repolarization, has also been shown to be inhibited by halothane, as well as by isoflurane.147Moreover, these anesthetics have been reported to inhibit delayed rectifier K+current (I  K)148and Ca2+-activated K+(KCa) channels.149Unlike KATPchannels, which are activated primarily under pathologic conditions, Kirchannels, I  to, I  Kand KCachannels play important roles in stabilizing the resting potential or modifying the shape of the action potential under normal conditions.150 

Volatile anesthetics have been reported to exert complex effects on sarcolemmal KATPchannels. Using inside-out patches excised from rabbit ventricular myocytes, Han et al.  151found that isoflurane decreased the duration of KATPchannel burst activity and increased the interburst interval without affecting channel kinetics within a burst. It should be noted that KATPchannels characteristically exhibit rundown (decreasing Poas a function of time) and show intermittent burst activity after patch excision.152These characteristics have been shown to be fully prevented by endogenous molecules such as PIP2(phosphatidylinositol 4,5-bisphosphate) and long-chain acyl-CoA esters,152suggesting that burst activity may in fact be restricted to excised patches and, possibly, pathologic conditions.152Han et al.  151also showed that isoflurane shifted the relation between [ATP] and Poto the left; that is, it decreased the sensitivity of the channel to inhibitory ATP. More recently, Stadnicka and Bosnjak153reported that isoflurane facilitates KATPchannel opening at reduced pH. Hence, under physiologic (and pathologic) conditions isoflurane may facilitate KATPchannel activation, which could explain, at least in part, its ability to shorten the action potential duration.

Volatile anesthetics have also been shown to confer ischemic-like preconditioning via  putative mitochondrial KATPchannels, with downstream signaling mediated by reactive oxygen species.154,155Cope et al. ,156for example, demonstrated that halothane, enflurane, and isoflurane decreased infarction size by about 50% (compared to control conditions) in rabbit hearts subjected to regional ischemia either in vitro  or in situ . In another study, the cardioprotective effect of isoflurane was shown to be blocked by 5-hydroxydecanoate, thereby implicating mitochondrial KATPchannels as mediators of volatile anesthetic-induced preconditioning.155In support of this conclusion, Nakae et al.  157recently reported that isoflurane increases the open probability of mitochondrial KATPchannels reconstituted in lipid bilayers. However, there is controversy as to whether mitochondrial KATPchannels play a role in preconditioning because the evidence for their involvement rests mainly on pharmacological foundations.158–161 

Recently, volatile anesthetics have been shown to activate the two-pore domain K+channels TASK and TREK,162,163which may be important targets contributing to cerebral depression. Meuth et al. ,164for example, recently suggested that TASK1 and TASK3 contribute to the activity of thalamocortical relay neurons involved in sleep-wake cycles. TASK and TREK channels, which are weakly expressed in heart compared with the brain, may play a role in modulating resting membrane potential and repolarization, such that these channels could contribute to anesthetic-induced shortening of the action potential. Although the effects of volatile anesthetics on various K+channels in the heart are unlikely to contribute significantly to the negative inotropy, their effects on the the action potential may contribute to anesthetic-induced arrhythmias.

Anesthetic Effects on SR Ca2+Content and Ca2+Release

In 1990, Herland et al.  165showed that halothane stimulated Ca2+efflux from the sarcoplasmic reticulum of chemically-skinned cardiac trabeculae. Connelly and Coronado166confirmed this observation using an alternative approach. After incorporating SR-rich vesicles into artificial lipid bilayers, these authors found that halothane and enflurane, but not isoflurane, increased open probability of the Ca2+-release channel (RyR). In accord with these findings, Lynch and Frazer167showed that halothane, but not isoflurane, enhanced the binding of [3H]-ryanodine to RYRs (note that ryanodine binds to RYRs with high affinity when channels are in the open state). Consistent with activation of the SR Ca2+-release channel, halothane, but not isoflurane, has been shown to evoke a transient increase in [Ca2+] in resting cardiac trabeculae.129This transient effect has also been seen in electrically stimulated myocytes. Harrison et al.  132showed that application of halothane, but not isoflurane, transiently potentiated the Ca2+transients and accompanying contractions of electrically paced myocytes (fig. 7).

Halothane and, to a lesser extent, isoflurane have been reported to decrease the Ca2+content of the SR of isolated intact myocytes132,133(but see Hannon and Cody136) and intact cardiac trabeculae,130,131whereas sevoflurane appears to have no effect.133,136Moreover, isoflurane and sevoflurane, but not halothane, were found to decrease fractional release, the fraction of the content that is released after stimulation.133,136 

Anesthetic Effects on Sarcolemmal NCX

The sarcolemmal NCX is a potentially important target of volatile anesthetics because stimulation of its Ca2+-efflux mode (forward mode) would diminish the SR Ca2+load and thereby decrease contractility. In a radioisotope study using rat myocyte suspensions, Haworth and Goknur168found that halothane, isoflurane, and enflurane dose-dependently inhibited the reverse mode, Na+-dependent 45Ca2+influx, of the NCX. This action could result in less trigger Ca2+for CICR (see Part I). Anesthetic sensitivity of the predominant forward mode, Na+-induced Ca2+-efflux, was not tested. In a more recent study, halothane (1–2 MAC) and sevoflurane (1–2 MAC) were also found to inhibit both the forward and reverse modes of the NCX in fluo-3 loaded rat cardiac myocytes.169On one hand, inhibition of the dominant forward (Ca2+-efflux) mode would be expected to produce positive inotropy whereas, on the other hand, inhibition of the reverse mode (Ca2+-influx) would produce the opposite effect. On balance, the above studies suggest that inhibition of the NCX would have either no net inotropic effect or, if anything, increase inotropy. It is worth noting that volatile anesthetics could exert a positive inotropic effect indirectly by inhibiting the Na+,K+-ATPase,170which would cause intracellular [Na+] to increase (favoring reverse mode NCX activity).

Anesthetic Inhibition of the Sarcolemmal and SR Ca2+Pumps (Ca2+-ATPases)

After inhibiting the NCX with Na+- and Ca2+-free solution, Hannon and Cody136showed that isoflurane and sevoflurane, but not halothane, decreased the rate of relaxation of caffeine-induced Ca2+transients in ferret ventricular myocytes. The authors concluded that isoflurane and sevoflurane inhibited the sarcolemmal Ca2+-ATPase. However, inhibitory effects of the anesthetic on the sarco-endoplasmic reticular Ca2+-ATPase isoform in cardiac SR (sarco-endoplasmic reticular Ca2+-ATPase 2a)171could have accounted, at least in part, for the observed reduction in the rate of Ca2+extrusion. In line with the interpretation of Hannon and Cody,136halothane and isoflurane have been reported to decrease surface membrane Ca2+-ATPase activity in erythrocytes172and neurons.173In any case, inhibition of the sarcolemmal Ca2+-pump (Ca2+-ATPase) would not be expected to produce a negative inotropic action but rather to produce positive inotropy secondary to facilitated SR Ca2+loading. On the contrary, inhibitory effects of volatile anesthetics on sarco-endoplasmic reticular Ca2+-ATPase171,174could contribute to the depletion of SR Ca2+stores by volatile anesthetics.

Gap Junctions

Halothane and isoflurane have been shown to uncouple electrically pairs of myocytes, indicating that these agents block gap junctions.175In accord, Burt and Spray176showed that halothane (2 mm) decreased gap junction conductance by reducing the number of open channels without affecting unitary conductance. In further work, He and Burt177have shown that halothane gates gap junctions to the closed state, the extent of which is dependent on anesthetic concentration and the connexin composition of the channel. Heteromeric channels were found to be more sensitive to inhibition by halothane than homomeric channels composed of either connexin 40 or connexin 43. The ability of halothane to induce uncoupling would not inhibit the contractility of individual myocytes, but it may perturb the normal spread of excitation, rendering the heart more prone to arrhythmias.

Anesthetic Effects on the Contractile Machinery and the pCa-force Relation

In addition to reducing Ca2+availability, volatile anesthetics may also decrease the responsiveness of the contractile proteins to a given amount of Ca2+. Bosnjak and Kampine123examined the effects of halothane on [Ca2+] (measured with aequorin) and tension, simultaneously measured in intact papillary muscle. Halothane decreased force proportionately more than peak Ca2+, suggesting that the anesthetic was having a direct inhibitory action at the level of the contractile proteins. In agreement with Bosnjak and Kampine,123Housmans178demonstrated that isoflurane depressed force, even when the Ca2+transient (measured using aequorin) was restored to control concentrations by elevating extracellular [Ca2+].

Many recent studies using various muscle preparations (single myocytes and thin ventricular trabeculae) have established that halothane and other anesthetics do indeed inhibit the Ca2+-responsiveness of the contractile proteins. For example, using thin ventricular trabeculae (diameter <250 μm) loaded with fluo-3, Hanley and Loiselle129showed that force remained depressed when the cytosolic Ca2+transient was restored to control concentrations by increasing extracellular [Ca2+] in the presence of halothane or isoflurane (fig. 9). Similar results with halothane and isoflurane were obtained by Jiang and Julian,130,131who also used rat trabeculae. Furthermore, Davies et al.  133likewise found that halothane and isoflurane, but not sevoflurane, decreased Ca2+-responsiveness of the contractile proteins in fura-2-loaded rat ventricular myocytes. In contrast to Davies et al. ,133Bartunek and Housmans179found that sevoflurane, in addition to decreasing Ca2+-availability, decreased myofibrillar Ca2+-responsiveness (see also Graham et al.  180).

Fig. 9. Simultaneous measurement of force and fluo-3 fluorescence, an index of [Ca2+], in a rat cardiac trabecula. Application of isoflurane decreased force and [Ca2+]. Restoration of the Ca2+transient amplitude by elevation of external [Ca2+] did not recover force, indicating that, in addition to decreasing Ca2+availability, the anesthetic decreased Ca2+-responsiveness of the contractile proteins. Adapted with permission from Hanley and Loiselle.  129 

Fig. 9. Simultaneous measurement of force and fluo-3 fluorescence, an index of [Ca2+], in a rat cardiac trabecula. Application of isoflurane decreased force and [Ca2+]. Restoration of the Ca2+transient amplitude by elevation of external [Ca2+] did not recover force, indicating that, in addition to decreasing Ca2+availability, the anesthetic decreased Ca2+-responsiveness of the contractile proteins. Adapted with permission from Hanley and Loiselle.  129 

Volatile anesthetics could depress myofibrillar Ca2+-responsiveness by decreasing Ca2+-sensitivity or maximal Ca2+-activated force of the contractile proteins. At least part of the depressive action of halothane and isoflurane is attributable to a reduction in maximal Ca2+-activated force because Hanley and Loiselle129showed that these agents decreased maximum force when trabeculae were tetanized in the presence of ryanodine and high extracellular [Ca2+] (fig. 10). Whether volatile anesthetics shift the pCa-force relation in “intact” cardiac muscle remains to be elucidated.

Fig. 10. Halothane and isoflurane decrease maximal Ca2+-activated force. Ryanodine-induced tetani were used to obtain maximal Ca2+-activated force in a trabecula (  A ). In the presence of halothane or isoflurane, maximal Ca2+-activated force was decreased (  B ). Reproduced with permission from Hanley and Loiselle.  129 

Fig. 10. Halothane and isoflurane decrease maximal Ca2+-activated force. Ryanodine-induced tetani were used to obtain maximal Ca2+-activated force in a trabecula (  A ). In the presence of halothane or isoflurane, maximal Ca2+-activated force was decreased (  B ). Reproduced with permission from Hanley and Loiselle.  129 

Studies using chemically or mechanically skinned cardiac muscle preparations have shed light on the direct effects of volatile anesthetics on the contractile proteins. Halothane, enflurane, and isoflurane have been shown to decrease maximal Ca2+-activated force in detergent-skinned human181and rat cardiac fibers.182Herland et al.  183found that high doses of halothane, but not enflurane or isoflurane, decreased maximal Ca2+-activated force in mechanically disrupted or saponin-skinned rat cardiac muscle preparations. When membranes were further disrupted with Triton X-100, maximal Ca2+-activated force was decreased by both halothane and isoflurane but not by enflurane. Using Triton X-100-skinned rat cardiac muscle, Prakash et al.  184found that halothane and sevoflurane, at 1 or 2 MAC, decreased maximal Ca2+-activated force. Hence, the majority of data using skinned cardiac muscle, as well as the study of Hanley and Loiselle129using intact trabeculae, indicate that volatile anesthetics decrease maximal Ca2+-activated force.

The next question is whether volatile anesthetics decrease the Ca2+-sensitivity (pCa for half-maximal activation) of the contractile proteins. Studies using skinned fibers have produced conflicting results. Murat et al.  182reported that halothane, enflurane, and isoflurane each increased the [Ca2+] for half-maximal activation in skinned cardiac fibers. In accord, Tavernier et al.  181found that halothane and isoflurane, each at 1 MAC, decreased Ca2+-sensitivity of the contractile proteins. However, Herland et al.  183found that the effect of anesthetics on Ca2+-sensitivity depended on the method of skinning employed. When the muscle was mechanically skinned or treated with saponin, sensitivity was increased by the anesthetics, whereas addition of Triton X-100 abolished the sensitizing effects of halothane and isoflurane and reduced the effect of enflurane. Hence, the disparate observed effects of anesthetics on Ca2+-sensitivity of the contractile proteins may reflect differences in technique, including differences in skinning procedure, species, muscle preparation, and anesthetic concentrations. Further studies using intact muscle preparations and graded tetani may help resolve whether volatile anesthetics decrease, increase, or have no effect on Ca2+-sensitivity of the contractile system. It should be noted that force will be reduced at any given [Ca2+] when sensitivity is unchanged but maximal Ca2+-activated force is reduced.

Anesthetic Effects on Cross-bridge Cycling

Murat et al.  185examined the effects of volatile anesthetics on force and stiffness during rapid length perturbations at controlled levels of contractile activation. Halothane, enflurane and isoflurane each decreased active stiffness, increased the stiffness/force ratio and increased the time constant of force recovery. These findings indicated that the anesthetics decreased the number of force-generating cross-bridges, decreased the force per cross bridge and reduced the rate of actomyosin-ATPase activity. Halothane and isoflurane, as well as sevoflurane, were also found to decrease dynamic stiffness, as well as shortening amplitude, in intact ferret papillary muscles.186In further work, Hannon et al. ,135using intact papillary muscles and ryanodine-induced tetani, found that isoflurane shifted the relation between [Ca2+] and rate of tension redevelopment toward higher [Ca2+], suggesting that the anesthetic reduced the rate of cross-bridge turnover. Thus, direct inhibitory effects of volatile anesthetics at the level of actin-myosin protein interaction may account for the ability of these agents to decrease maximal Ca2+-activated force.

Inhibitory Effects of Anesthetics on Mitochondrial Function

Volatile anesthetics have long been known to impair nicotinamide adenine dinucleotide

(NADH) oxidation by mitochondria,187–190suggesting that these agents may inhibit complex I (NADH:ubiquinone oxidoreductase) of the electron transport chain. Using isolated liver mitochondria, Hall et al.  190showed that halothane inhibited NADH oxidation at concentrations that did not affect succinate oxidation. Consistent with inhibition of NADH oxidation, halothane and isoflurane,129,191,192as well as sevoflurane,193have been shown to increase NADH fluorescence in intact cardiac muscle.

Recently, Hanley et al. ,194using heart submitochondrial particles, have shown that halothane, isoflurane, and sevoflurane indeed inhibit the activity of NADH:ubiquinone oxidoreductase (complex I), which accounts for about 40% of the proton pumping capacity of the respiratory chain (fig. 11). Halothane also inhibited, albeit less potently, succinate dehydrogenase, the catalytic component of complex II (fig. 11). At concentrations equivalent to 2 MAC, halothane and isoflurane decreased activity of complex I by ∼20%, whereas sevoflurane decreased activity by ∼10%. Volatile anesthetics are usually administrated in combination with nitrous oxide (N2O), which has itself been reported to inhibit the respiratory chain, albeit at complex IV (cytochrome c  oxidase).195,196 

Fig. 11. Mitochondrial sites of action of volatile anesthetics. Halothane, isoflurane, and sevoflurane partially inhibit complex I of the electron transport chain. Halothane is also a weak inhibitor at complex II, whereas N2O (which is commonly coadministered with volatile anesthetics), like NO, has been reported to exert an inhibitory action at complex IV. Reproduced with permission from Hanley  et al. 194 

Fig. 11. Mitochondrial sites of action of volatile anesthetics. Halothane, isoflurane, and sevoflurane partially inhibit complex I of the electron transport chain. Halothane is also a weak inhibitor at complex II, whereas N2O (which is commonly coadministered with volatile anesthetics), like NO, has been reported to exert an inhibitory action at complex IV. Reproduced with permission from Hanley  et al. 194 

Could inhibitory effects on electron transport chain activity account for the negative inotropic action of volatile anesthetics? In 31P nuclear magnetic resonance studies, 1.5% halothane (2 MAC) was shown to not decrease creatine phosphate or ATP concentrations.197,198Moreover, the observed ∼50% decrease in rate pressure product (an index of cardiac mechanical work) induced by halothane was not accompanied by an increase in the concentrations of Pi(inorganic phosphate) or H+, inhibitors of excitation-contraction coupling and the contractile apparatus,94indicating that impairment of oxidative metabolism is not the major mechanism by which volatile anesthetics exert negative inotropy. Although effects of volatile anesthetics on mitochondrial function cannot explain the negative inotropy, the inhibition of the respiratory chain (energy supply) will decrease cardiac reserve.194Inhibition at complex I may explain how volatile anesthetics increase the rate of production of reactive oxygen species, which are thought to mediate anesthetic-induced, ischemic-like preconditioning.154,199 

Although volatile anesthetics have been shown to target multiple sites in heart muscle cells, the sites of action responsible for negative inotropy are predominantly the L-type Ca2+channels, the SR, and the contractile apparatus. Volatile anesthetics reduce L-type Ca2+current (the trigger for SR Ca2+release) and, depending on the agent used, deplete SR Ca2+content or decrease fractional release of Ca2+from the SR. These anesthetic actions depress the elevation in cytosolic [Ca2+] after membrane depolarization. This negative inotropic effect of reduced Ca2+-availability is compounded by inhibitory effects of the volatile anesthetics at the level of the contractile apparatus (decreased Ca2+-responsiveness). Volatile anesthetics impair cross-bridge cycling, decrease maximal Ca2+-activated force, and, possibly, shift the [Ca2+]-force relation to higher [Ca2+]. Anesthetic actions on mitochondrial function, K+channels, the sarcolemmal Ca2+-ATPase, and the NCX probably contribute minimally to the negative inotropy.

Rusy BF, Komai H: Anesthetic depression of myocardial contractility: A review of possible mechanisms. Anesthesiology 1987; 67:745–66
Maier SK, Westenbroek RE, Schenkman KA, Feigl EO, Scheuer T, Catterall WA: An unexpected role for brain-type sodium channels in coupling of cell surface depolarization to contraction in the heart. Proc Natl Acad Sci U S A 2002; 99:4073–8
Kamp TJ, Hell JW: Regulation of cardiac L-type calcium channels by protein kinase A and protein kinase C. Circ Res 2000; 87:1095–102
Sommer JR, Johnson EA: Ultrastructure of cardiac muscle, Handbook of Physiology; The Heart: The Cardiovascular System. Edited by Berne R. Bethesda, American Physiological Society, 1979, pp 113–86Berne R
American Physiological Society
Cheng H, Lederer WJ, Cannell MB: Calcium sparks: Elementary events underlying excitation-contraction coupling in heart muscle. Science 1993; 262:740–4
Beeler GW, Reuter H: Membrane calcium current in ventricular myocardial fibres. J Physiol 1970; 207:191–209
Cavalie A, Ochi R, Pelzer D, Trautwein W: Elementary currents through Ca2+channels in guinea pig myocytes. Pflügers Arch 1983; 398:284–97
Hadley RW, Hume JR: An intrinsic potential-dependent inactivation mechanism associated with calcium channels in guinea-pig myocytes. J Physiol 1987; 389:205–22
Kass RS, Sanguinetti MC: Inactivation of calcium channel current in the calf cardiac Purkinje fiber: Evidence for voltage- and calcium-mediated mechanisms. J Gen Physiol 1984; 84:705–26
Bechem M, Pott L: Removal of Ca current inactivation in dialysed guinea-pig atrial cardioballs by Ca chelators. Pflugers Arch 1985; 404:10–20
Sipido KR, Callewaert G, Carmeliet E: Inhibition and rapid recovery of Ca2+current during Ca2+release from sarcoplasmic reticulum in guinea pig ventricular myocytes. Circ Res 1995; 76:102–9
Grantham CJ, Cannell MB: Calcium influx during the cardiac action potential in guinea-pig ventricular myocytes. Circ. Res 1996; 79:194–200
Li L, Satoh H, Ginsburg KS, Bers DM: The effect of Ca2+-calmodulin-dependent protein kinase II on cardiac excitation-contraction coupling in ferret ventricular myocytes. J Physiol 1997; 501:17–31
Peterson BZ, DeMaria CD, Adelman JP, Yue DT: Calmodulin is the Ca2+sensor for Ca2+-dependent inactivation of L-type calcium channels. Neuron 1999; 22:549–58
Fabiato A: Time and calcium dependence of activation and inactivation of calcium-induced release of calcium from the sarcoplasmic reticulum of a skinned canine cardiac Purkinje cell. J Gen Physiol 1985; 85:247–89
Muller FU, Kirchhefer U, Begrow F, Reinke U, Neumann J, Schmitz W: Junctional sarcoplasmic reticulum transmembrane proteins in the heart. Basic Res Cardiol 2002; 97 Suppl 1:I52–5
Crespo LM, Grantham CJ, Cannell MB: Kinetics, stoichiometry and role of the Na-Ca exchange mechanism in isolated cardiac myocytes. Nature 1990; 345:618–21
Choi HS, Trafford AW, Eisner DA: Measurement of calcium entry and exit in quiescent rat ventricular myocytes. Pflugers Arch 2000; 440:600–8
Berlin JR, Cannell MB, Lederer WJ: Regulation of twitch tension in sheep cardiac Purkinje fibers during calcium overload. Am J Physiol 1987; 253:H1540–7
Leblanc N, Hume JR: Sodium current-induced release of calcium from cardiac sarcoplasmic reticulum. Science 1990; 248:372–6
Levi AJ, Brooksby P, Hancox JC: A role for depolarisation induced calcium entry on the Na-Ca exchange in triggering intracellular calcium release and contraction in rat ventricular myocytes. Cardiovasc Res 1993; 27:1677–90
Kohmoto O, Levi AJ, Bridge JHB: Relation between reverse sodium-calcium exchange and sarcoplasmic reticulum calcium release in guinea pig ventricular cells. Circ Res 1994; 74:550–4
Adachi-Akahane S, Lu L, Li Z, Frank JS, Philipson KD, Morad M: Calcium signaling in transgenic mice overexpressing cardiac Na+-Ca2+exchanger. J Gen Physiol 1997; 109:717–29
Evans AM, Cannell MB: The role of L-type Ca2+current and Na+current-stimulated Na/Ca exchange in triggering SR calcium release in guinea-pig cardiac ventricular myocytes. Cardiovasc Res 1997; 35:294–302
Levesque PC, Leblanc N, Hume JR: Release of calcium from guinea pig cardiac sarcoplasmic reticulum induced by sodium-calcium exchange. Cardiovasc Res 1994; 28:370–8
Sipido KR, Carmeliet E, Pappano A: Na+current and Ca2+release from the sarcoplasmic reticulum during action potentials in guinea-pig ventricular myocytes. J Physiol 1995; 489:1–17
Sipido KR, Maes M, Van de Werf F: Low efficiency of Ca2+entry through the Na+-Ca2+exchanger as trigger for Ca2+release from the sarcoplasmic reticulum: A comparison between L-type Ca2+current and reverse-mode Na+-Ca2+exchange. Circ Res 1997; 81:1034–44
Franzini-Armstrong C, Protasi F, Ramesh V: Shape, size, and distribution of Ca2+release units and couplons in skeletal and cardiac muscles. Biophysical J 1999; 77:1528–39
Langer GA, Peskoff A: Calcium concentration and movement in the diadic cleft space of the cardiac ventricular cell. Biophys J 1996; 70:1169–82
Cannell MB, Soeller C: Numerical analysis of ryanodine receptor activation by L-type channel activity in the cardiac muscle diad. Biophys J 1997; 73:112–22
Soeller C, Cannell MB: Numerical simulation of local calcium movements during L-type calcium channel gating in the cardiac diad. Biophys J 1997; 73:97–111
Carl SL, Felix K, Caswell AH, Brandt NR, Ball WJJ, Vaghy PL, Meissner G, Ferguson DG: Immunolocalization of sarcolemmal dihydropyridine receptor and sarcoplasmic reticular triadin and ryanodine receptor in rabbit venricle and atrium. J Cell Biol 1995; 129:672–82
Sun X-H, Protasi F, Takahashi M, Takeshima H, Ferguson DG, Franzini-Armstrong C: Molecular architecture of membranes involved in excitation-contraction coupling of cardiac muscle. J Cell Biol 1995; 129:659–71
Scriven DRL, Dan P, Moore EDW: Distribution of proteins implicated in excitation-contraction coupling in rat ventricular myocytes. Biophysical J 2000; 79:2682–91
Thomas MJ, Sjaastad I, Andersen K, Helm PJ, Wasserstrom JA, Sejersted OM, Ottersen OP: Localization and function of the Na+/Ca2+-exchanger in normal and detubulated rat cardiomyocytes. J Mol Cell Cardiol 2003; 35:1325–37
Stern MD: Theory of excitation-contraction coupling in cardiac muscle. Biophys J 1992; 63:497–517
Cannell MB, Cheng H, Lederer WJ: Spatial non-uniformities in [Ca2+]iduring excitation-contraction coupling in cardiac myocytes. Biophys J 1994; 67:1942–56
Cannell MB, Cheng H, Lederer WJ: The control of calcium release in heart muscle. Science 1995; 268:1045–9
Cheng H, Cannell MB, Lederer WJ: Partial inhibition of Ca2+current by methoxyverapamil (D600) reveals spatial nonuniformities in [Ca2+]iduring excitation-contraction coupling in cardiac myocytes. Circ Res 1995; 76:236–41
Parker I, Zang WJ, Wier WG: Ca2+sparks involving multiple Ca2+release sites along Z-lines in rat heart cells. J Physiol 1996; 497:31–8
Gomez AM, Cheng H, Lederer WJ, Bers DM: Ca2+diffusion and sarcoplasmic reticulum transport both contribute to [Ca2+]idecline during Ca2+sparks in rat ventricular myocytes. J Physiol 1996; 496:575–81
Cheng H, Lederer MR, Xiao RP, Gomez AM, Zhou YY, Ziman B, Spurgeon H, Lakatta EG, Lederer WJ: Excitation-contraction coupling in heart: New insights from Ca2+sparks. Cell Calcium 1996; 20:129–40
Niggli E: Localized intracellular calcium signaling in muscle: Calcium sparks and calcium quarks. Annu Rev Physiol 1999; 61:311–35
Santana LF, Cheng H, Gomez AM, Cannell MB, Lederer WJ: Relation between the sarcolemmal Ca2+current and Ca2+sparks and local control theories for cardiac excitation-contraction coupling. Circ Res 1996; 78:166–71
Cheng H, Cannell MB, Lederer WJ: Propagation of excitation-contraction coupling into ventricular myocytes. Pflugers Arch 1994; 428:415–7
Bridge JH, Ershler PR, Cannell MB: Properties of Ca2+sparks evoked by action potentials in mouse ventricular myocytes. J Physiol 1999; 518:469–78
Lipp P, Niggli E: Submicroscopic calcium signals as fundamental events of excitation-contraction coupling in guinea-pig cardiac myocytes. J Physiol 1996; 492:31–8
Wang SQ, Song LS, Lakatta EG, Cheng H: Ca2+signalling between single L-type Ca2+channels and ryanodine receptors in heart cells. Nature 2001; 410:592–7
Soeller C, Cannell MB: Estimation of the sarcoplasmic reticulum Ca2+release flux underlying Ca2+sparks. Biophys J 2002; 82:2396–414
Bers DM: Calcium fluxes involved in control of cardiac myocyte contraction. Circ Res 2000; 87:275–81
Eisner DA, Choi HS, Díaz ME, O'Neill SC, Trafford AW: Integrative analysis of calcium cycling in cardiac muscle. Circ Res 2000; 87:1087–94
Sitsapesan R, Williams AJ: Regulation of the gating of the sheep cardiac sarcoplasmic reticulum Ca2+-release channel by luminal Ca2+. J Membr Biol 1994; 137:215–26
Lukyanenko V, Gyorke I, Gyorke S: Regulation of calcium release by calcium inside the sarcoplasmic reticulum in ventricular myocytes. Pflugers Arch 1996; 432:1047–54
Bassani JWM, Bassani RA, Bers DM: Relaxation in rabbit and rat cardiac cells: Species-dependent differences in cellular mechanisms. J Physiol 1994; 476:279–93
Bers DM: Cardiac excitation-contraction coupling. Nature 2002; 415:198–205
Bers DM: Excitation-contraction coupling and cardiac contractile force, 2nd edition. Dordrecht, Netherlands, Kluwer Academic Publishers, 2001, pp 40–60
Dordrecht, Netherlands
Kluwer Academic Publishers
Blaustein MP, Lederer WJ: Sodium/calcium exchange: Its physiological implications. Physiol Rev 1999; 79:763–854
Cooper PJ, Ward ML, Hanley PJ, Denyer GR, Loiselle DS: Metabolic consequences of a species difference in Gibbs free energy of Na+/Ca2+exchange: Rat versus guinea pig. Am J Physiol 2001; 280:R1221–9
Ward ML, Cooper PJ, Hanley PJ, Loiselle DS: Species-independent metabolic response to an increase of [Ca2+]iin quiescent cardiac muscle. Clin Exp Pharmacol Physiol 2003; 30:586–9
Berlin JR, Cannell MB, Lederer WJ: Cellular origins of the transient inward current in cardiac myocytes: Role of fluctuations and waves of elevated intracellular calcium. Circ Res 1989; 65:115–26
ter Keurs HE, Zhang YM, Miura M: Damage-induced arrhythmias: Reversal of excitation-contraction coupling. Cardiovasc Res 1998; 40:444–55
Solaro RJ, Rarick HM: Troponin and tropomyosin: Proteins that switch on and tune in the activity of cardiac myofilaments. Circ Res 1998; 83:471–80
Barth E, Stämmler G, Speiser B, Schaper J: Ultrastructural quantitation of mitochondria and myofilaments in cardiac muscle from 10 different animal species including man. J Mol Cell Cardiol 1992; 24:669–81
Kim H-D, Kim CH, Rah B-J, Chung H-I, Shim T-S: Quantitative study on the relation between structural and functional properties of the hearts from three different mammals. Anat Rec 1994; 238:199–206
Sugiura S: Actin-myosin interaction. Cardiovasc Res 1999; 44:266–73
Li Y, Mui S, Brown JH, Strand J, Reshetnikova L, Tobacman LS, Cohen C: The crystal structure of the C-terminal fragment of striated-muscle α-tropomyosin reveals a key troponin T recognition site. Proc Natl Acad Sci U S A 2002; 99:7378–83
Stewart M: Structural basis for bending tropomyosin around actin in muscle thin filaments. Proc Natl Acad Sci U S A 2001; 98:8165–6
Ward DG, Cornes MP, Trayer IP: Structural consequences of cardiac troponin I phosphorylation. J Biol Chem 2002; 277:41795–801
Turnbull L, Hoh JFY, Ludowyke RI, Rossmanith GH: Troponin I phosphorylation enhances crossbridge kinetics during β-adrenergic stimulation in rat cardiac tissue. J Physiol 2002; 542:911–20
Gordon AM, Homsher E, Regnier M: Regulation of contraction in striated muscle. Physiol Rev 2000; 80:853–924
Stuyvers BDM, Miura M, ter Keurs HEDJ: Dynamics of viscoelastic properties of rat cardiac sarcomeres during the diastolic interval: Involvement of Ca2+. J Physiol 1997; 502:661–77
Fuchs F, Smith SH: Calcium, cross-bridges, and the Frank-Starling relationship. News Physiol Sci 2001; 16:5–10
Perry SV: What is the role of tropomyosin in the regulation of muscle contraction? J Muscle Res Cell Motil 2003; 24:593–6
Huxley AF: Muscle structure and theories of contraction. Prog Biophysics Biophys Chem 1957; 7:255–318
Cooke R: Actomyosin interaction in striated muscle. Physiol Rev 1997; 77:671–97
Huxley AF, Simmons RM: Proposed mechanism of force generation in striated muscle. Nature 1971; 233:533–8
Finer JT, Simmons RM, Spudich JA: Single myosin molecule mechanics: piconewton forces and nanometre steps. Nature 1994; 368:113–9
Lymn RW, Taylor EW: Mechanism of adenosine triphosphate hydrolysis by actomyosin. Biochemistry 1971; 10:4617–24
Ishijima A, Kojima H, Funatsu T, Tokunaga M, Higuchi H, Tanaka H, Yanagida T: Simultaneous observation of individual ATPase and mechanical events by a single myosin molecule during interaction with actin. Cell 1998; 92:161–71
Huxley AF: Cross-bridge action: Present views, prospects, and unknowns. J Biomech 2000; 33:1189–95
Marban E, Kusuoka H, Yue DT, Weisfeldt ML, Wier WG: Maximal Ca2+-activated force elicited by tetanization of ferret papillary muscle and whole heart: Mechanism and characteristics of steady contractile activation in intact myocardium. Circ Res 1986; 59:262–9
Gao WD, Backx PH, Azan-Backx M, Marban E: Myofilament Ca2+sensitivity in intact versus skinned rat ventricular muscle. Circ Res 1994; 74:408–15
ter Keurs HEDJ, Rijnsburger WH, van Heuningen R, Nagelsmit MJ: Tension development and sarcomere length in rat cardiac trabeculae: Evidence of length-dependent activation. Circ Res 1980; 46:703–14
Kentish JC, ter Keurs HEDJ, Ricciardi L, Bucx JJJ, Noble MIM: Comparison between the sarcomere length-force relations of intact and skinned trabeculae from rat right ventricle: Influence of calcium concentrations on these relations. Circ Res 1986; 58:755–68
Fitzsimons DP, Moss RL: Strong binding of myosin modulates length-dependent Ca2+activation of rat ventricular myocytes. Circ Res 1998; 83:602–7
Allen DG, Kentish JC: The cellular basis of the length-tension relationship in cardiac muscle. J Mol Cell Cardiol 1985; 17:821–40
Petroff MGV, Kim SH, Pepe S, Dessy C, Marbán E, Balligand J-L, Sollott SJ: Endogenous nitric oxide mechanisms mediate the stretch dependence of Ca2+release in cardiomyocytes. Nat Cell Biol 3;3:867–73
Irving TC, Konhilas J, Perry D, Fischetti R, de Tombe PP: Myofilament lattice spacing as a function of sarcomere length in isolated rat myocardium. Am J Physiol 2000; 279:H2568–73
Konhilas JP, Irving TC, de Tombe PP: Myofilament calcium sensitivity in skinned rat cardiac trabeculae: Role of interfilament spacing. Circ Res 2002; 90:59–65
Labeit S, Kolmerer B, Linke WA: The giant protein titin. Emerging roles in physiology and pathophysiology. Circ Res 1997; 80:290–4
Cazorla O, Vassort G, Garnier D, Le Guennec J-Y: Length modulation of active force in rat cardiac myocytes: Is titin the sensor? J Mol Cell Cardiol 1999; 31:1215–27
Fukuda N, Sasaki D, Ishiwata S, Kurihara S: Length dependence of tension generation in rat skinned cardiac muscle: Role of titin in the Frank-Starling mechanism of the heart. Circulation 2001; 104:1639–45
Wu Y, Cazorla O, Labeit D, Labeit S, Granzier H: Changes in titin and collagen underlie diastolic stiffness diversity of cardiac muscle. J Mol Cell Cardiol 2000; 32:2151–61
Stuyvers BD, McCulloch AD, Guo J, Duff HJ, ter Keurs HEDJ: Effect of stimulation rate, sarcomere length and Ca2+on force generation by mouse cardiac muscle. J Physiol 2002; 544:817–30
Granzier HL, Irving TC: Passive tension in cardiac muscle: Contribution of collagen, titin, microtubules, and intermediate filaments. Biophys J 1995; 68:1027–44
Kellermayer MSZ, Smith SB, Granzier HL, Bustamante C: Folding-unfolding transitions in single titin molecules charaterized with laser tweezers. Science 1997; 276:1112–6
Tskhovrebova L, Trinick J, Sleep JA, Simmons RM: Elasticity and unfolding of single molecules of the giant muscle protein titin. Nature 1997; 387:308–12
Linke WA, Ivemeyer M, Labeit S, Hinssen H, Rüegg JC, Gautel M: Actin-titin interaction in cardiac myofibrils: Probing a physiological role. Biophysical J 1997; 73:905–19
Stuyvers BD, Miura M, Jin JP, ter Keurs HE: Ca2+-dependence of diastolic properties of cardiac sarcomeres: involvement of titin. Prog Biophys Mol Biol 1998; 69:425–43
Hütter JF, Piper HM, Spieckermann PG: Effect of fatty acid oxidation on efficiency of energy production in rat heart. Am J Physiol 1985; 249:H723–8
van der Vusse GJ, Glatz JFC, Stam HCG, Reneman RS: Fatty acid homeostasis in the normoxic and ischemic heart. Physiol Rev 1992; 72:881–940
Chatham JC: Lactate: The forgotten fuel! J Physiol 2002; 542:333
Bessman SP, Geiger PJ: Transport of energy in muscle: The phosphorylcreatine shuttle. Science 1981; 211:448–52
Schramm M, Klieber H-G, Daut J: The energy expenditure of actomyosin-ATPase, Ca2+-ATPase and Na+,K+-ATPase in guinea-pig cardiac ventricular muscle. J Physiol 1994; 481:647–62
Ebus JP, Stienen GJM: Origin of concurrent ATPase activities in skinned cardiac trabeculae of rat. J Physiol 1996; 492:675–87
Gibbs CL, Loiselle DS: Cardiac basal metabolism. Jpn J Physiol 2001; 51:399–426
Kirichok Y, Krapivinsky G, Clapham DE: The mitochondrial calcium uniporter is a highly selective ion channel. Nature 2004; 427:360–4
Baysal K, Jung DW, Gunter KK, Gunter TE, Brierley GP: Na+-dependent Ca2+efflux mechanism of heart mitochondria is not a passive Ca2+/2Na+exchanger. Am J Physiol 1994; 266:C800–8
Balaban RS: Cardiac energy metabolism homeostasis: Role of cytosolic calcium. J Mol Cell Cardiol 2002; 34:1259–71
McCormack JG, Halestrap AP, Denton RM: Role of calcium ions in regulation of mammalian intramitochondrial metabolism. Physiol Rev 1990; 70:391–425
Trollinger DR, Cascio WE, Lemasters JJ: Selective loading of rhod 2 into mitochondria shows mitochondrial Ca2+transients during the contractile cycle in adult rabbit myocytes. Biochem Biophys Res Commun 1997; 236:738–42
Zhou Z, Matlib MA, Bers DM: Cytosolic and mitochondrial Ca2+signals in patch clamped mammalian ventricular myocytes. J Physiol 1998; 507:379–403
Griffiths EJ: Species dependence of mitochondrial calcium transients during excitation-contraction coupling in isolated cardiomyocytes. Biochem Biophys Res Commun 1999; 263:554–9
Morris AJ, Malbon CC: Physiological regulation of G protein-linked signaling. Physiol Rev 1999; 79:1373–430
Xiao RP: β-Adrenergic signaling in the heart: dual coupling of the β2-adrenergic receptor to Gsand Giproteins. Sci STKE 2001; 104:RE15
Pierce KL, Premont RT, Lefkowitz RJ: Seven-transmembrane receptors. Nat Rev Mol Cell Biol 2002; 3:639–50
Rockman HA, Koch WJ, Lefkowitz RJ: Seven-transmembrane-spanning receptors and heart function. Nature 2002; 415:206–12
Hunter PJ, McCulloch AD, ter Keurs HEDJ: Modelling the mechanical properties of cardiac muscle. Prog Biophys Mol Biol 1998; 69:289–331
Li Y, Kranias EG, Mignery GA, Bers DM: Protein kinase A phosphorylation of the ryanodine receptor does not affect calcium sparks in mouse ventricular myocytes. Circ Res 2002; 90:309–16
Urban BW: Current assessment of targets and theories of anaesthesia. Br J Anaesth 2002; 89:167–83
Eger EI II, Saidman LJ, Brandstater B: Minimum alveolar anesthetic concentration: A standard of anesthetic potency. Anesthesiology 1965; 26:756–63
Franks NP, Lieb WR: Temperature dependence of the potency of volatile general anesthetics: implications for in vitro experiments. Anesthesiology 1996; 84:716–20
Bosnjak ZJ, Kampine JP: Effects of halothane on transmembrane potentials, Ca2+transients, and papillary muscle tension in the cat. Am J Physiol 1986; 251:H374–81
Lee JA, Westerblad H, Allen DG: Changes in tetanic and resting [Ca2+]iduring fatigue and recovery of single muscle fibres from Xenopus laevis.  J Physiol 1991; 433:307–26
Baker PF, Schapira AHV: Anaesthetics increase light emission from aequorin at constant ionised calcium. Nature 1980; 284:168–9
Housmans PR, Wanek LA: Effects of halothane, enflurane, and isoflurane on measurements of Ca2+by calcium electrode and aequorin luminescence. Anal Biochem 2000; 284:60–4
Grynkiewicz G, Poenie M, Tsien RY: A new generation of Ca2+indicators with greatly improved fluorescence properties. J Biol Chem 1985; 260:3440–50
Minta A, Kao JPY, Tsien RY: Fluorescent indicators for cytosolic calcium based on rhodamine and fluorescein chromophores. J Biol Chem 1989; 264:8171–8
Hanley PJ, Loiselle DS: Mechanisms of force inhibition by halothane and isoflurane in intact rat cardiac muscle. J Physiol 1998; 506:231–44
Jiang Y, Julian FJ: Effects of halothane on [Ca2+]itransient, SR Ca2+content, and force in intact rat heart trabeculae. Am J Physiol 1998; 274:H106–14
Jiang Y, Julian FJ: Effects of isoflurane on [Ca2+]i, SR Ca2+content, and twitch force in intact trabeculae. Am J Physiol 1998; 275:H1360–9
Harrison SM, Robinson M, Davies LA, Hopkins PM; Boyett MR: Mechanisms underlying the inotropic action of halothane on intact rat ventricular myocytes. Br J Anaesth 1999; 82:609–21
Davies LA, Gibson CN, Boyett MR, Hopkins PM, Harrison SM: Effects of isoflurane, sevoflurane, and halothane on myofilament Ca2+sensitivity and sarcoplasmic reticulum Ca2+release in rat ventricular myocytes. Anesthesiology 2000; 93:1034–44
Housmans PR, Wanek LA, Carton EG, Bartunek AE: Effects of halothane and isoflurane on the intracellular Ca2+transient in ferret cardiac muscle. Anesthesiology 2000; 93:189–201
Hannon JD, Cody MJ, Housmans PR: Effects of isoflurane on intracellular calcium and myocardial crossbridge kinetics in tetanized papillary muscle. Anesthesiology 2001; 94:856–61
Hannon JD, Cody MJ: Effects of volatile anesthetics on sarcolemmal calcium transport and sarcoplasmic reticulum calcium content in isolated myocytes. Anesthesiology 2002; 96:1457–64
Lynch C, Vogel S, Sperelakis N: Halothane depression of myocardial slow action potentials. Anesthesiology 1981; 55:360–8
Terrar DA, Victory JGG: Effects of halothane on membrane currents associated with contraction in single myocytes isolated from guinea-pig ventricle. Br J Parmacol 1988; 94:500–8
Terrar DA, Victory JGG: Isoflurane depresses membrane currents associated with contraction in myocytes isolated from guinea-pig ventricle. Anesthesiology 1988; 69:742–9
Bosnjak ZJ, Supan FD, Rusch NJ: The effects of halothane, enflurane, and isoflurane on calcium current in isolated canine ventricular cells. Anesthesiology 1991; 74:340–5
Pancrazio JJ: Halothane and isoflurane preferentially depress a slowly inactivating component of Ca2+channel current in guinea-pig myocytes. J Physiol 1996; 494:91–103
Hirota K, Ito Y, Masuda A, Momose Y: Effects of halothane on membrane ionic currents in guinea pig atrial and ventricular myocytes. Acta Anaesthesiol Scand 1989; 33:239–44
Puttick RM, Terrar DA: Effects of propofol and enflurane on action potentials, membrane currents and contraction of guinea-pig isolated ventricular myocytes. Br J Pharmacol 1992; 107:559–65
Rithalia A, Gibson CN, Hopkins PM, Harrison SM: Halothane inhibits contraction and action potential duration to a greater extent in subendocardial than subepicardial myocytes from the rat left ventricle. Anesthesiology 2001; 95:1213–9
Stadnicka A, Bosnjak ZJ, Kampine JP, Kwok WM: Modulation of cardiac inward rectifier K+current by halothane and isoflurane. Anesth Analg 2000; 90:824–33
Stadnicka A, Bosnjak ZJ, Kampine JP, Kwok WM: Effects of sevoflurane on inward rectifier K+current in guinea pig ventricular cardiomyocytes. Am J Physiol 1997; 273:H324–32
Hüneke R, Jüngling E, Skasa M, Rossaint R, Lückhoff A: Effects of the anesthetic gases xenon, halothane, and isoflurane on calcium and potassium currents in human atrial cardiomyocytes. Anesthesiology 2001; 95:999–1006
Suzuki A, Bosnjak ZJ, Kwok WM: The effects of isoflurane on the cardiac slowly activating delayed-rectifier potassium channel in guinea pig ventricular myocytes. Anesth Analg 2003; 96:1308–15
Hashiguchi-Ikeda M, Namba T, Ishii TM, Hisano T, Fukuda K: Halothane inhibits an intermediate conductance Ca2+-activated K+channel by acting at the extracellular side of the ionic pore. Anesthesiology 2003; 99:1340–5
Xu Y, Tuteja D, Zhang Z, Xu D, Zhang Y, Rodriguez J, Nie L, Tuxson HR, Young JN, Glatter KA, Vázquez AE, Yamoah EN, Chiamvimonvat N: Molecular identification and functional roles of a Ca2+-activated K+channel in human and mouse hearts. J Biol Chem 2003; 278:49085–94
Han J, Kim E, Ho WK, Earm YE: Effects of volatile anesthetic isoflurane on ATP-sensitive K+channels in rabbit ventricular myocytes. Biochem Biophys Res Commun 1996; 229:852–6
Liu GX, Hanley PJ, Ray J, Daut J: Long-chain acyl-coenzyme A esters and fattya cids directly link metabolism to KATPchannels in the heart. Circ Res 2001; 88:918–24
Stadnicka A, Bosnjak ZJ: Isoflurane decreases ATP sensitivity of guinea pig cardiac sarcolemmal KATPchannel at reduced intracellular pH. Anesthesiology 2003; 98:396–403
Müllenheim J, Ebel D, Fräβdorf J, Preckel B, Thämer V, Schlack W: Isoflurane preconditions myocardium against infarction via release of free radicals. Anesthesiology 2002; 96:934–40
Tanaka K, Weihrauch D, Ludwig LM, Kersten JR, Pagel PS, Warltier DC: Mitochondrial adenosine triphosphate-regulated potassium channel opening acts as a trigger for isoflurane-induced preconditioning by generating reactive oxygen species. Anesthesiology 2003; 98:935–43
Cope DK, Impastato WK, Cohen MV, Downey JM: Volatile anesthetics protect the ischemic rabbit myocardium from infarction. Anesthesiology 1997; 86:699–709
Nakae Y, Kwok WM, Bosnjak ZJ, Jiang MT: Isoflurane activates rat mitochondrial ATP-sensitive K+channels reconstituted in lipid bilayers. Am J Physiol 2003; 284:H1865–71
Hu H, Sato T, Seharaseyon J, Liu Y, Johns DC, O'Rourke B, Marban E: Pharmacological and histochemical distinctions between molecularly defined sarcolemmal KATPchannels and native cardiac mitochondrial KATPchannels. Mol Pharmacol 1999; 55:1000–5
Hanley PJ, Mickel M, Löffler M, Brandt U, Daut J: KATPchannel-independent targets of diazoxide and 5-hydroxydecanoate in the heart. J Physiol 2002; 542:735–41
Lim KHH, Javadov SA, Das M, Clarke SJ, Suleiman M-S, Halestrap AP: The effects of ischaemic preconditioning, diazoxide and 5-hydroxydecanoate on rat heart mitochondrial volume and respiration. J Physiol 2002; 545:961–74
Hanley PJ, Gopalan KV, Lareau RA, Srivastava DK, von Meltzer M, Daut J: β-Oxidation of 5-hydroxydecanoate, a putative blocker of mitochondrial ATP-sensitive potassium channels. J Physiol 2003; 547:387–93
Patel AJ, Honoré E, Lesage F, Fink M, Romey G, Lazdunski M: Inhalational anesthetics activate two-pore-domain background K+channels. Nat Neurosci 1999; 2:422–6
Patel AJ, Honoré E: Anesthetic-sensitive 2P domain K+channels. Anesthesiology 2001; 95:1013–21
Meuth SG, Budde T, Kanyshkova T, Broicher T, Munsch T, Pape HC: Contribution of TWIK-related acid-sensitive K+channel 1 (TASK1) and TASK3 channels to the control of activity modes in thalamocortical neurons. J Neurosci 2003; 23:6460–9
Herland JS, Julian FJ, Stephenson DG: Halothane increases Ca2+efflux via Ca2+channels of sarcoplasmic reticulum in chemically skinned rat myocardium. J Physiol 1990; 426:1–18
Connelly TJ, Coronado R: Activation of the Ca2+release channel of cardiac sarcoplasmic reticulum by volatile anesthetics. Anesthesiology 1994; 81:459–69
Lynch C, Frazer MJ: Anesthetic alteration of ryanodine binding by cardiac calcium release channels. Biochim Biophys Acta 1994; 1194:109–17
Haworth RA, Goknur AB: Inhibition of sodium/calcium exchange and calcium channels of heart cells by volatile anesthetics. Anesthesiology 1995; 82:1255–65
Seckin I, Sieck GC, Prakash YS: Volatile anaesthetic effects on Na+-Ca2+exchange in rat cardiac myocytes. J Physiol 2002; 532:91–104
Molliex S, Dureuil B, Aubier M, Friedlander G, Desmonts JM, Clerici C: Halothane decreases Na,K-ATPase, and Na channel activity in alveolar type II cells. Anesthesiology 1998; 88:1606–13
Karon BS, Autry JM, Shi Y, Garnett CE, Inesi G, Jones LR, Kutchai H, Thomas DD: Different anesthetic sensitivities of skeletal and cardiac isoforms of the Ca-ATPase. Biochemistry 1999; 38:9301–7
Lopez MM, Kosk-Kosicka D: How do volatile anesthetics inhibit Ca2+-ATPases? J Biol Chem 1995; 270:28239–45
Franks JJ, Horn JL, Janicki PK, Singh G: Halothane, isoflurane, xenon, and nitrous oxide inhibit calcium ATPase pump activity in rat brain synaptic plasma membranes. Anesthesiology 1995; 82:108–17
Miao N, Frazer MJ, Lynch C: Anesthetic actions on calcium uptake and calcium-dependent adenosine triphosphatase activity of cardiac sarcoplasmic reticulum. Adv Pharmacol 1994; 31:145–65
Terrar DA, Victory JG: Influence of halothane on electrical coupling in cell pairs isolated from guinea-pig ventricle. Br J Pharmacol 1988; 94:509–14
Burt JM, Spray DC: Volatile anesthetics block intercellular communication between neonatal rat myocardial cells. Circ Res 1989; 65:829–37
He DS, Burt JM: Mechanism and selectivity of the effects of halothane on gap junction channel function. Circ Res 2000; 86:e104–9
Housmans PR: Mechanisms of negative inotropy of halothane, enflurane and isoflurane in isolated mammalian ventricular muscle. Adv Exp Med Biol 1991; 301:199–204
Bartunek AE, Housmans PR: Effects of sevoflurane on the intracellular Ca2+transient in ferret cardiac muscle. Anesthesiology 2000; 93:1500–8
Graham MD, Lambert EL, Hopkins PM, Harrison SM: Mechanisms contributing to the inotropic effects of sevoflurane in rat ventricular myocytes. J Physiol 2003; 551P:C5
Tavernier BM, Adnet PJ, Imbenotte M, Etchrivi TS, Reyford H, Haudecoeur G, Scherpereel P, Krivosic-Horber RM: Halothane and isoflurane decrease calcium sensitivity and maximal force in human skinned cardiac fibers. Anesthesiology 1994; 80:625–33
Murat I, Ventura-Clapier R, Vassort G: Halothane, enflurane, and isoflurane decrease calcium sensitivity and maximal force in detergent-treated rat cardiac fibers. Anesthesiology 1988; 69:892–9
Herland JS, Julian FJ, Stephenson DG: Effects of halothane, enflurane, and isoflurane on skinned rat myocardium activated by Ca2+. Am J Physiol 1993; 264:H224–32
Prakash YS, Cody MJ, Hannon JD, Housmans PR, Sieck GC: Comparison of volatile anesthetic effects on actin-myosin cross-bridge cycling in neonatal versus adult cardiac muscle. Anesthesiology 2000; 92:1114–25
Murat I, Lechene P, Ventura-Clapier R: Effects of volatile anesthetics on mechanical properties of rat cardiac skinned fibers. Anesthesiology 1990; 73:73–81
Bartunek AE, Claes VA, Housmans PR: Effects of volatile anesthetics on stiffness of mammalian ventricular muscle. J Appl Physiol 2001; 91:1563–73
Biebuyck JF: Effects of anaesthetic agents on metabolic pathways: Fuel utilization and supply during anaesthesia. Br J Anaesth 1973; 45:263–8
Cohen PJ: Effect of anesthetics on mitochondrial function. Anesthesiology 1973; 39:153–64
Grist EM, Baum H: A possible mechanism for the halothane-induced inhibition of mitochondrial respiration: Binding of endogenous calcium to NADH dehydrogenase. Febs Lett 1974; 48:41–4
Hall GM, Kirtland SJ, Baum H: The inhibition of mitochondrial respiration by inhalational anaesthetic agents. Br J Anaesth 1973; 45:1005–9
Kissin I, Aultman DF, Smith LR: Effects of volatile anesthetics on myocardial oxidation-reduction status assessed by NADH fluorometry. Anesthesiology 1983; 59:447–52
Jiang Y, Julian FJ: Pacing rate, halothane, and BDM affect fura 2 reporting of [Ca2+]iin intact rat trabeculae. Am J Physiol 1997; 273:C2046–56
Riess ML, Camara AKS, Chen Q, Novalija E, Rhodes SS, Stowe DF: Altered NADH and improved function by anesthetic and ischemic preconditioning in guinea pig intact hearts. Am J Physiol 2002; 283:H53–60
Hanley PJ, Ray J, Brandt U, Daut J: Halothane, isoflurane and sevoflurane inhibit NADH: ubiquinone oxidoreductase (complex I) of cardiac mitochondria. J Physiol 2002; 544:687–93
Einarsdóttir Ó, Caughey WS: Interactions of the anesthetic nitrous oxide with bovine heart cytochrome c  oxidase. J Biol Chem 1988; 263:9199–205
Chervin C, Thibaud MC: Inhibition of plant and animal cytochrome oxidases by nitrous oxide as a function of cytochrome c  concentration. Biochemie 1992; 74:1125–7
McAuliffe JJ, Hickey PR: The effect of halothane on the steady-state levels of high-energy phosphates in the neonatal heart. Anesthesiology 1987; 67:231–5
Murray PA, Blanck TJJ, Rogers MC, Jacobus WE: Effects of halothane on myocardial high-energy phosphate metabolism and intracellular pH utilizing 31P NMR spectroscopy. Anesthesiology 1987; 67:649–53
Novalija E, Varadarajan SG, Camara AKS, An J, Chen Q, Riess M, Hogg N, Stowe DF: Anesthetic preconditioning: Triggering role of reactive oxygen and nitrogen species in isolated hearts. Am J Physiol 2002; 283:H44–52
Hanley PJ, Young AA, LeGrice IJ, Edgar SG, Loiselle DS: 3-Dimensional configuration of perimysial collagen fibres in rat cardiac muscle at resting and extended sarcomere lengths. J Physiol 1999; 517:831–7