Ca²⁺ Signaling in Striated Muscle Cells During Intracellular Acidosis

Abstract

The cytosolic pH (pH_i) of mammalian cells is tightly maintained at values ~7.2. Cytoplasmic acidosis (pH_i < 6.8) occurs when the intracellular proton concentration ([H⁺]_i) exceeds the buffering capacity of the cytosol and transport processes to extrude protons are exhausted. During intracellular acidosis, the contractility of cardiac and skeletal muscle cells is strongly reduced, often at sufficient Ca²⁺ levels. A contraction of striated muscle is achieved when the intracellular calcium (Ca²⁺) concentration rises above resting levels. The amplitude and kinetics of Ca²⁺ signals are controlled by Ca²⁺ handling proteins and force is generated if Ca²⁺ ions interact with contractile filaments of the sarcomere. Some aspects of this phenomenon, such as the biochemical origin of excessive protons in working muscle cells and molecular interactions of protons with Ca²⁺ handling proteins or contractile filaments, are not yet fully understood. This review summarizes our current understanding of how striated muscle cells handle Ca²⁺ and H⁺ and how a rise in [H⁺]_i may interfere with Ca²⁺ signaling in the working skeletal muscle (fatigue) or during ischemic events in cardiac muscle. Finally, we briefly address experimental strategies to measure Ca²⁺ signaling at different pH values with fluorescent probes and highlight their limitations.

1. Introduction

The concentration of free protons ([H⁺]) in aqueous solutions is low and commonly expressed on a logarithmic scale as pH = −log [H⁺] [1,2]. In mammalian cells, extracellular pH (pH_o) is rigorously maintained between 7.3–7.4 [3,4,5], whereas intracellular pH (pH_i) ranges between 7.0–7.2 (skeletal muscle cells and cardiac myocytes) [6,7]. When transient metabolic changes in free [H⁺] occur, pH_i is kept constant by means of buffering. In biological solutions, weak acids or bases, such as phosphate groups and side chains of amino acids of proteins, serve as proton buffers. In addition, cells employ transport systems for acids and bases, such as proton pumps and transporters or bicarbonate shuttles to fine-tune pH_i on the long-term range [4]. There is a modulatory role of pH_i on Ca²⁺ signaling in striated muscle cells [8,9,10,11] and non-excitable cells [12]. Cells use Ca²⁺ ions as secondary messengers to regulate different cellular functions, including contraction of smooth and striated muscle [13,14,15]. Ca²⁺ signaling occurs when the low, resting Ca²⁺ concentration (~100 nM) suddenly increases to values up to 1–2 µM and Ca²⁺ sensor proteins integrate the information encoded by Ca²⁺ signals, such as amplitude, frequency and duration, with cellular functions. Endogenous Ca²⁺ sensing proteins relevant for striated muscle cell function are calmodulin (CaM), which controls enzymes and ion channels [16] and troponin, which controls muscle contraction (see below) [17,18]. The intracellular Ca²⁺ concentration ([Ca²⁺]_CYT) increases due to activity of plasmalemmal ion channels mediating Ca²⁺ influx and/or intracellular Ca²⁺ release channels, which liberate Ca²⁺ from intracellular stores, such as the endoplasmic reticulum (ER) or the sarcoplasmic reticulum (SR) in muscle cells. Ca²⁺ signals that control muscle contraction are brief and reversible (“Ca²⁺ transient”) and resting Ca²⁺ levels are established via removal of Ca²⁺ ions from the cytosol by Ca²⁺ pumps, such as the plasma membrane Ca²⁺ ATPase (PMCA, Ca²⁺ extrusion), the sarcoplasmic/endoplasmic Ca²⁺ ATPase (SERCA, store-refilling) and plasmalemmal Ca²⁺ exchangers [15,19,20,21]. In addition to Ca²⁺ release and removal mechanisms, intracellular Ca²⁺ signals are shaped by reversible binding of Ca²⁺ ions to intracellular proteins serving as Ca²⁺ buffers [22]. Ca²⁺ signaling is affected by pH_i via two mechanisms: First, H⁺ and Ca²⁺ compete for identical binding sites at protein buffers. This has the consequence that protonation of a buffer will increase its K_d for Ca²⁺ [23] and a rise in [H⁺] (a fall in pH) will unavoidably increase the free [Ca²⁺] and vice versa [3]. Second, the protonation of Ca²⁺ handling proteins at secondary sites [24] may allosterically alter their function to transport Ca²⁺ [25].

Acidosis is the fall of pH below physiological values [26] and is recognized as a clinical parameter when plasma pH is <7.35 (acidemia) due to metabolic or respiratory disturbances [27]. Respiratory acidosis can cause a rapid fall in pH_o due to an extracellular buildup of CO₂. Because CO₂ is highly diffusible across lipid bilayers, a rise in extracellular CO₂ causes an increase of the cell’s acid load, which lowers pH_i [4,28]. Intracellular acidosis is defined as a fall of pH_i < 6.8 and it occurs when there are more free cytosolic protons than buffers can bind and transport processes can remove from the cytoplasm [4]. Acidosis is frequently observed under physiological conditions due to metabolic processes in working muscle or pathologically in ischemic tissue under hypoxic conditions or during inflammation. It is worth noting that any fall in pH_o during extracellular acidosis will directly cause an accompanying fall in pH_i since protons can easily cross the plasma membrane via diffusion or transport processes [3]. Therefore, a direct relation between low pH_o values and intracellular acidosis has been demonstrated for cardiac, skeletal and smooth muscle cells [29,30]. This review will focus on the effects of intracellular acidosis on striated muscle Ca²⁺ handling and contraction. Intracellular acidosis causes a remarkable reduction in striated muscle contractility, which is the ability of the muscle to generate force. As part of this special issue, our review aims to introduce the Ca²⁺ signaling events that lead to contraction in striated muscle cells and to provide an overview of the most important cellular mechanisms that contribute to the reduction in contractility during acidosis in skeletal and cardiac cells. We will also briefly address optical techniques to assess Ca²⁺ signaling and pH_i with fluorescent probes in living cells to study those processes.

2. Excitation–Contraction Coupling (ECC) in Striated Muscle Cells

The signaling process that mediates contraction of striated muscle cells is called excitation-contraction coupling (ECC) [31,32,33]. The term summarizes a chain of signaling events in which an electrical signal of the plasma membrane, an action potential (AP), is converted into mechanical contraction via intracellular Ca²⁺ release (Figure 1). APs propagate into the depth of muscle cells via invaginations of the plasma membrane named T-tubules (Figure 1, 1). In striated muscle cells, excitation of the T-tubule is coupled with a mechanism that releases Ca²⁺ from the SR (electrical coupling, Figure 1, 2). In skeletal muscle cells, electrical coupling is achieved by activation of the dihydropyridine receptor (DHPR), which is directly coupled to a Ca²⁺ release channels of the sarcoplasmic reticulum (SR), the ryanodine receptors type 1 (RyR1s, Figure 1, box 1, 2a) [34]. In cardiac cells, electrical coupling involves the L-type Ca²⁺ channel (LTCC), which mediates Ca²⁺ influx. Ca²⁺ ions diffuse to adjacent SR membranes and open the Ca²⁺-gated, cardiac ryanodine receptor type 2 (RyR2, Figure 1, box 1, 2b) [35,36]. Opening of either type of RyR causes rapid release of Ca²⁺ from the SR into the cytosol down its concentration gradient (Figure 1, 3), which is converted into mechanical contraction at the sarcomere (Figure 1, 4). The kinetics, amplitude and duration of this intracellular Ca²⁺ elevation define the speed, strength and duration of muscle contractions, respectively [37].

The molecular steps controlling Ca²⁺-dependent contractions are identical for cardiac and skeletal muscle cells: a rise in intracellular Ca²⁺ induces cross-bride formation between actin and myosin, a process that is regulated by the Ca²⁺-sensitive troponin–tropomyosin complex [38]. At low [Ca²⁺]_CYT, troponin masks the myosin interaction sites on the actin molecule and cross-bridge formation is prevented. At high [Ca²⁺]_CYT, Ca²⁺ binds to the regulatory protein troponin and induces a conformational change of the troponin–tropomyosin complex, which liberates myosin binding sites on the actin molecule (Figure 1, box 2) [39]. Cross-bridge formation will shorten sarcomeres, which are the contractile units of striated muscle cells (Figure 1, 4) [17,40,41]. The force a striated muscle can generate depends on the sum of all cross-bridges, a process with steep Ca²⁺ dependence [42,43]. The second important molecule that regulates the cross-bridge cycle is ATP, which binds to myosin [44]. The affinity of myosin molecules for actin is low when ATP is bound to the myosin head (loose cross bridges) and high if ATP is hydrolyzed and ADP is bound to myosin (Figure 1, box 2). The hydrolysis rate of ATP and the ATP-to-ADP exchange rate at myosin heads determine the speed of new cross-bridge formations (i.e., the speed of contraction) [45,46,47].

The muscle relaxes when [Ca²⁺]_CYT falls back to resting levels. This is controlled by Ca²⁺-dependent inactivation of RyRs at high cytosolic Ca²⁺ concentrations [35,36,48] and by activation of Ca²⁺ transport mechanisms that remove Ca²⁺ from the cytosol. The major transport processes in striated muscle cells involve SERCA at the SR (Figure 1, 5) and the plasmalemmal sodium calcium exchanger (NCX) [33,37,49] (Figure 1, 6). The latter uses the Na⁺ gradient across the plasma membrane for Ca²⁺ extrusion, which is generated by the Na⁺/K⁺-ATPase (Figure 1, 7). For skeletal muscle cells there is experimental evidence that refilling of the SR with Ca²⁺ ions is supported by Ca²⁺-influx via store-operated Ca²⁺ entry (SOCE, Figure 1, 9) [50]. In contrast, cardiac SOCE generates only locally restricted Ca²⁺ signals that do not seem to contribute to contractile Ca²⁺ signaling [51]. Established functional roles for cardiac SOCE are the regulation of pacemaker activity of the sinoatrial node [52] and activation of gene transcription processes associated with cardiac hypertrophy and heart failure [53,54].

Additionally, mitochondria can store Ca²⁺ ions and thus may affect [Ca²⁺]_CYT and ECC. Any elevation in cytoplasmic Ca²⁺ may cause an uptake of Ca²⁺ into mitochondria via Ca²⁺ diffusion through the mitochondrial uniporter (Figure 1, 8) [55]. The exact [Ca²⁺] within the mitochondrial matrix is cell type-specific. In resting striated muscle cells, it is equal or close to [Ca²⁺]_CYT [56]. In rat cardiac myocytes, expression of a mitochondria-specific Ca²⁺ biosensor revealed that resting mitochondrial Ca²⁺ levels were between 70 nM and 100 nM and the mitochondrial Ca²⁺ concentration increased with each cytosolic Ca²⁺ transient when ECC was evoked by electrical stimulation of the cells, an effect that was attributed to diffusion of Ca²⁺ from the cytosol into mitochondria [57]. The effect of mitochondrial Ca²⁺ uptake on [Ca²⁺]_CYT is more pronounced in cardiac cells, in which mitochondria occupy ~30% of the cell volume [58,59]. Whether mitochondrial Ca²⁺ buffering is relevant in skeletal muscle cells is a matter of debate [60]. One limiting factor might be that mitochondria occupy only a small volume (~2% to 6%) of skeletal muscle cells [61]. Functional roles of mitochondrial Ca²⁺ uptake/buffering include shaping of Ca²⁺ transients, fine-tuning of CICR at ER/mitochondrial contact sites [62] and providing a feedback signal that couples mitochondrial ATP production to its cytoplasmic demand during cell contractions [58,59].

Taking this complexity of ECC into consideration, there are many molecular steps that can be modified by acidosis. We will summarize next how a fall in pH_i below normal values can alter ECC, Ca²⁺ signaling and Ca²⁺ buffering. This is relevant for the contractile function of striated muscle, which is severely depressed by intracellular acidosis occurring in physiological and pathological conditions.

3. Mechanisms Inducing Intracellular Acidosis in Striated Muscle Cells

Skeletal muscle. Fatigue is the inability of the muscle to generate sustained force or power during repetitive contractions [63,64]. Within this period, the speed of individual muscle contractions slows down and maximal force is reduced. Fatigue is well pronounced in fast muscle fibers and less or not at all pronounced in slower muscle fibers [65]. Initially it was assumed that fatigue is the simple result of excessive consumption of energy (ATP) during heavy workloads of the muscle [66]. In this scenario, fatigue occurs whenever muscle fibers consume ATP faster than it can be supplied by metabolic processes. While this mechanism may contribute to fatigue in fast-twitching muscle fibers, some studies indicated that intracellular ATP levels are not likely to fall to a level that may compromise contraction in most muscle types [67] and muscle fibers displayed fatigue at constant ATP levels [68]. Instead, there is evidence that skeletal muscle fatigue is the result of an accumulation of protons and P_i in the working muscle. Acidosis has been associated with fatigue and occurs in a reversible manner during intense exercise [69], in which pH_i can fall from ~7.0 (at rest) to ~6.4 (fatigued) in human skeletal muscle [7]. During longer periods of exercise, ATP synthesis depends on glycolysis that generates pyruvate, which either fuels ATP generation in mitochondria (via the tricarboxylic acid cycle) or which is converted to lactate by the enzyme lactate dehydrogenase (LDH) in the cytosol when mitochondrial uptake is saturated [70]. Since it was discovered that the lactate concentration rises in working skeletal muscles [71], the term “lactic acidosis” has been associated with fatigue. The underlying concept implies that accumulation of lactic acid or lactate due to increased glycolysis rates will cause intracellular acidification [7,72]. Others have argued that the biochemistry of skeletal muscle cells would not be compatible with this concept. Some counter arguments include the following: (1) The final product of glycolysis in the cytosol is pyruvate. Mitochondria import pyruvate and consume protons to supply ATP in working muscle, which limits pyruvate accumulation in the cytosol [73,74]. Furthermore, when mitochondrial import is exhausted, excessive pyruvate in the cytosol is converted by LDH to lactate, but not to lactic acid (which could lower pH_i). Lactate, in turn, stimulates monocarboxylate transporters (MCTs), which are symporters that transport lactate and protons out of the cell (Figure 2) [75]. Due to this transport mechanism, the plasma concentration of lactate can rise about five times to ~8 mM during intense exercise [76]. Extracellular lactate can enter neighboring muscle cells or cells of other tissues via MCTs, which operate bidirectionally, where it fuels ATP synthesis following its conversion back into pyruvate [77]. The conclusion is that mitochondrial metabolism and MCT-mediated transport will limit the accumulation of lactate and protons, so that the fall in pH_i cannot be caused by glycolysis alone. (2) Based on experiments using skinned skeletal muscle fibers, there is no direct evidence that lactate impairs contractility, even at high concentrations [70]. It has been suggested that the high rates of ATP hydrolysis at myosin molecules during heavy exercise would generate excessive protons in the cytoplasm [70,78,79,80]. Irrespective of the biochemical origin of those protons, significant acidification of skeletal muscle cells occurs during long periods of intense exercise and intramuscular pH_i was reported to fall to values ≤ 6.4 in fatigued skeletal muscle [7]. Considering all this, mechanisms inducing intracellular acidosis are not restricted to ATP deficiency, lactate accumulation or high glycolysis rates, but also caused by direct accumulation of protons stemming from alternative biochemical processes. It is likely that mechanisms other than ATP hydrolysis may contribute to proton accumulation, such as release of protons from buffers or using protons as counter ions for Ca²⁺ transportation into organelles. Those mechanisms are not well investigated yet (discussed in Section 5), and they may contribute to local rather than global changes in pH.

Cardiac muscle. Intracellular acidosis does not occur during normal activity of the heart. Metabolic changes that occur during physiological heartbeats do not cause acidification [81], probably due to the large buffering capacity of cardiac myocytes for protons (20–90 mM/pH unit) [31]. However, severe intracellular acidosis occurs in cardiac cells during pathologies that alter the acid–base homeostasis and lead to extracellular acidification, due to respiratory or metabolic disturbances, sleep apnea and ischemia. Respiratory acidosis is characterized by a buildup of extracellular CO₂, which induces a rapid fall in pH_i [4]. This is caused via diffusion of CO₂ across the plasma membrane and/or via cellular uptake of protons from the extracellular space. The latter is evenly effective, because the plasma membrane has a large conductance for protons [82] and the electrochemical driving force for protons favors proton influx [4]. Consequently, there is a direct relationship between the fall of pH_o and a corresponding fall of pH_i in muscle cells [30,83]. Cardiac ischemia lowers pH_i significantly: in diabetic cardiac myocytes, pH_i values as low as 6.0 have been observed during ischemic episodes, an effect that was reversible once normal oxygenation was established [84]. When pH_i falls this much, the cardiac contractility is markedly reduced [85]. Thus, in contrast to skeletal muscle, the mechanisms leading to intracellular acidification of cardiac myocytes are rather caused by extracellular pH changes than by intrinsic metabolic changes in pH_i.

4. Effect of Low pH_i on Striated Muscle Contractility

Skeletal muscle. High-intensity exercise affects ECC via mechanisms that are pH-independent and pH-dependent. In intact muscle tissue, there is an accumulation of extracellular K⁺ within T-tubules due to K⁺ efflux during APs, which depolarizes the tubules. ECC and Ca²⁺ release was shown to be efficient up to [K⁺]_o~10 mM but strongly impaired when [K⁺]_o reached concentrations of 14–15 mM [86]. Such large K⁺ concentrations were observed locally, within the small volume of T-Tubules [87]. The resulting membrane depolarization limits the spread of excitation within the T-tubule and delays the recovery of the DHPR voltage sensor from inactivation, both of which render electrical coupling less efficient [88]. Intracellular acidosis has several consequences on ECC. (1) Inhibition of RyR1 by protons reduces Ca²⁺ release (Figure 2, 1). Single channel recordings demonstrated maximal RyR1 channel open probabilities Po at pH = 7.4, but almost complete closure of the RYR1 at pH = 6.5 [36]. This limits the amount of Ca²⁺ that is available for contractions and can reduce the peak force per individual twitch by more than 30% in skinned skeletal muscle fibers [89]. (2) There are multiple effects on proteins of the contractile apparatus (Figure 2, 2): first, protonation reduces the Ca²⁺-sensitivity of purified troponin C [90] and approximately three-times more Ca²⁺ is required to form all cross-bridges required for 50% of force generation (pCa₅₀) in skinned muscle fibers [91]. Furthermore, the interaction of troponin C and troponin I, which defines the speed of cross-bridge formation, is less efficient, as shown in myofibril preparations [92]. Protonation of myosin reduces its affinity for actin, which affects the total number of cross-bridges and reduces the muscle tension by about 20% and 45%, depending on the type of skinned muscle fiber (fast or slow twitch) studied [93,94]. Importantly, those negative inotropic effects persisted even at saturating Ca²⁺ concentrations [95], suggesting that pH-related modifications of contractile proteins dominate the negative inotropic effect. (3) High rates of ATP hydrolysis in the working muscle result in accumulation of ADP, P_i and H⁺ within the sarcomere (Figure 2, 3). Protons and P_i act synergistically and have a large negative effect on contractility: P_i reaches peak concentrations between 15 mM and 30 mM in working skeletal muscle [96]. Raising P_i to 30 mM induced a large shift in the force–pCa relationship to the right for both slow- and fast-type, skinned muscle fibers [97], which means that more Ca²⁺ is required to generate force. This suggests that, like protons, P_i alone can strongly depress the generation of force. Furthermore, an accumulation of P_i stabilizes the ADP + P_i-bound conformation of myosin, which slows down the transition from ADP-to-ATP-bound form. This reduces the velocity of cross-bridge formation and impairs the formation of new cross-bridges. In addition, it reduces the force that previously formed cross-bridges had already developed [97,98]. Moreover, P_i can react with protons to its diprotonated form, which seems to have a larger negative inotropic effect than P_i or protons alone when applied to skinned fibers [99]. In combination, all those molecular events reduce the efficiency of Ca²⁺ ions to activate the cross-bridge cycle and/or reduce the force that existing cross-bridges can produce. (4) A rise in intracellular protons reduces the pump function of SERCA (Figure 2, 4). When SERCA function was measured in vesicles obtained from skeletal muscle SR preparations, a change in pH from 7.0 to 6.0 reduced maximal pump rates (V_max) by ~50%, an effect that was accompanied by a reduction in the pump’s Ca²⁺-affinity [100]. This is consistent with the observed decline in SR Ca²⁺-content and Ca²⁺ transient amplitude during acidosis in skeletal muscle fibers [101,102]. (5) In sarcolemmal vesicle preparations, the transport rate of NCX, which is stimulated by intracellular Ca²⁺, depends on pH_i: transport by NCX was stimulated at pH = 9, but inhibited at pH = 6 [103]. This inhibition occurred allosterically following protonation of two histidine residues (H124, H165) within the transporter molecule, which are not located within the Ca²⁺ binding domains [104]. Any reduction in NCX transport rate may cause an elevation of [Ca²⁺]_CYT (Figure 2, 5). Because the cardiac isoform NCX1 has higher maximal Ca²⁺ transport rates than the skeletal isoform NCX3 [33,105], its inhibition by protons may cause a larger rise in [Ca²⁺]_CYT in cardiac cells than in skeletal muscle cells [106,107]. (6) Ca²⁺ currents underlying SOCE (Figure 2, 7) were dependent on pH_i. Ca²⁺ release-activated Ca²⁺ currents (I_CRAC) recorded at pH_i = 6.3 were reduced by more than 50% as compared to control currents recorded at pH_i = 7.3 [108]. Furthermore, I_CRAC was strongly dependent on pH_o, which falls during ischemia or tissue inflammation. A reduction in pH_o from 7.4 to 6.3 almost completely abolished I_CRAC. The dependencies of SOCE currents on internal and external pH have been attributed to protonation of several key residues distributed throughout the Orai molecule [109]. SOCE activity contributes to SR Ca²⁺ load in intact skeletal muscle fibers [50], suggesting that any reduction in SOCE-mediated Ca²⁺ influx may reduce the filling status of the SR in skeletal muscle cells during acidosis.

The mechanisms described above collectively contribute to exercise-induced or physiological fatigue at low pH_i values in skeletal muscle. It is important to note that contractility experiments are often conducted using single muscle fibers (intact or skinned) to precisely control experimental parameters. Can one extrapolate those results to the situation of intact muscles in vivo? Performance measurements in muscles from healthy subjects showed that exercise-induced, intracellular acidosis (pH_i~6.5) reduced the total force, speed of contraction and peak power [110] to similar degrees than reported for single fibers. However, a comparison between different muscle types proves difficult because the degree of fatigue appears to be fiber type-specific: whereas intracellular acidosis exerted a pronounced negative inotropic effect in fast twitching fibers, it had little effect on slow twitching muscle fibers [110]. Therefore, not every type of muscle may display the same sensitivity to intracellular pH changes. Moreover, in intact muscle multiple metabolic changes occur at the same time: for example, mitochondria produce large amounts of reactive oxygen species (ROS) [111] and the increasing temperature in working muscle also seems to impair muscle performance [112]. ROS are known to modify the function of Ca²⁺ handling proteins, which impairs cellular Ca²⁺ handling [113,114]. The negative effect of ROS on skeletal muscle function is even more evident when the native ability of the cells to buffer ROS is reduced, as in metabolic disorders or during inflammation [115]. Thus, it may be difficult to estimate the relative contribution of each metabolite to fatigue in vivo (see Discussion below). The development of novel animal models for exercise-induced fatigue may help to better understand long-term molecular changes underlying skeletal muscle fatigue on systemic and organ levels in future studies [116]. If pathological acidemia occurs, the contractility of skeletal muscle can be further impaired by additional long-term structural and functional changes in the muscle that involve degradation of proteins and impaired energy supply due to mitochondrial dysfunction [117].

Cardiac muscle. Many of those effects described above apply to cardiac myocytes as well. There are, however, some notable differences: At normal pH_o, there is no net accumulation of K⁺ in T-tubules during contractions with high frequencies, because K⁺ removal via Na⁺/K⁺-ATPase pump activity is more efficient in cardiac myocytes than in skeletal muscle cells [88]. Unlike in the working skeletal muscle, in which glycolysis is the main source for ATP, the majority (>90%) of cardiac ATP stems from aerobic processes [118] and the main source for intracellular acidification is CO₂ released by mitochondria. CO₂ can leave the cell by diffusion across the membrane, or it reacts with H₂O to bicarbonate and protons, a reaction that is catalyzed by CAs [119]. It is assumed that under physiological conditions, the rate of protons generated by hydration of CO₂ is in balance with transport rates of proton export mechanisms [120] (see below). However, significant intracellular acidosis causes a strong depression in the contractility of cardiac Purkinje fibers [121]. It is frequently observed during cardiac ischemia, which is characterized by a fall in pH_o and pH_i, a reduced rate of ATP synthesis, negative inotropy [122,123,124] and, surprisingly, by a rise in [Ca²⁺]_CYT [123]. Intracellular acidosis has the following effects on cardiac ECC: (1) in isolated ventricular myocytes, protons affected the gating mechanism of L-type currents, and the amplitude of currents were 2.5 times smaller at pH_i = 6.2 as compared to pH_i = 7.2 [125], which reduces the efficiency of electrical coupling (Figure 1, box 1, 2b) [126]. In addition, low pH_i prevents PKA-mediated phosphorylation of LTCCs following β-adrenergic stimulation, which normally yields to larger Ca²⁺ currents and a gain in ECC efficiency [127]. The same mechanism blunts the compensatory positive inotropic effect that sympathetic nerve stimulation normally has on the human heart once the pump function turns weaker [128]. (2) Lipid bilayer experiments revealed an inhibition of RyR2s by cytoplasmic protons [129]. In intact and skinned cardiac myocyte preparations, the same effect caused less Ca²⁺ release from the SR per AP (Figure 2, 1), reflected by systolic Ca²⁺ transients with amplitudes that can drop by 50% at the beginning of acidosis [130,131,132]. However, during prolonged periods of acidosis the Ca²⁺ transient amplitudes recover [130], which is caused by an increase in diastolic [Ca²⁺]_CYT and subsequent increase in SR Ca²⁺ load [35,123]. (3) Experiments using either papillary muscle or cardiac Purkinje fibers revealed that the increase in diastolic [Ca²⁺]_CYT is partially caused by release of Ca²⁺ ions from internal buffers due to competition of H⁺ and Ca²⁺ ions for common binding sites (discussed in Section 5) [133,134] and by the following modifications of ion transport processes: the sodium–proton exchanger (NHE) is stimulated by intracellular protons (Figure 2, 11) [135] and NHE activation will lead to accumulation of internal Na⁺. Simultaneous inhibition of the Na⁺/K⁺-ATPase (Figure 2, 6), which would normally remove excessive Na⁺, further contributes to the accumulation of internal Na⁺ at low pH_i [136]. This reduces the Na⁺ gradient across the membrane that provides energy for other Na⁺-dependent transporters, as shown in inside-out patch-clamp experiments on ventricular myocytes [137]. Na⁺ accumulation affects Ca²⁺ extrusion by NCX: at low, physiological Na⁺ concentrations, allosteric inhibition of NCX by cytoplasmic Na⁺ fine-tunes the transport rate of NCX in forward mode and thus, regulates cardiac excitability and beat-to-beat Ca²⁺ changes in isolated myocytes and in intact hearts [138]. However, at higher internal Na⁺ concentrations, NCX will start to remove excessive Na⁺ at the cost of transporting Ca²⁺ into the cell (reverse mode of NCX, discussed below) [103]. In addition to its modulation by Na⁺, NCX itself is directly inhibited by protons as described above (Figure 2, 5) [104], an effect that is sensitized by intracellular Na⁺ when assessed in excised patches from ventricular sarcolemma [139]. In ventricular myocytes, the reduced transport rates of NCX in its forward mode caused a net elevation of diastolic [Ca²⁺]_CYT, which stimulates SERCA and increases SR Ca²⁺ load by more than 100 µM [123]. The stimulatory effect of Ca²⁺ on SERCA seems to compensate for the three-times slower transport rates (V_max) induced by protonation of cardiac SERCA alone, which was observed in experiments using SR vesicle preparations (Figure 2, 4) [100]. Following cardiac ischemia, the reverse mode of NCX causes severe damage of cardiac tissue. Due to the low pH_o during the ischemic period, the transport rate of NHE is reduced. Once reperfusion occurs, pH_o recovers quickly and NHE begins to rapidly transport protons out of the cell. This causes fast accumulation of Na⁺ (Figure 2, 11) and, in turn, rapid Ca²⁺ influx as NCX tries to remove the excessive Na⁺ in its reverse operation mode. In hearts from animal models and humans, this unusual large influx of Ca²⁺ via NCX represents a substrate for cardiac arrhythmias and causes more damage of the myocardium (reperfusion injury) [140,141]. A recent study suggests that inhibition of cardiac NCX1 by protons may contribute to Ca²⁺ overload and reperfusion injury in the same way as the Na⁺-driven regulation: during experimentally-induced acidosis (pH_i = 6.5), ventricular myocytes from genetically modified mice expressing a proton-insensitive mutant of NCX1 (H165A) did not show increases in Ca²⁺ transient amplitudes, suggesting that NCX was not inhibited and hearts from transgenic mice displayed reduced tissue damage following an ischemia–reperfusion protocol in comparison to wild-type mice [142]. This suggests, in turn, that the modulations of NCX by Na⁺ and H⁺ may induce Ca²⁺ overload in a synergistic fashion. (4) Like in skeletal muscle, in isolated papillary muscle the negative inotropic effect of intracellular acidosis is dominated by the reduced Ca²⁺ sensitivity of the cardiac contractile apparatus (pCa₅₀ is shifted by +0.1 unit/per fall of 0.1 pH unit) [124]. In experiments using intact myocytes, contractility remained depressed during acidosis, although the systolic Ca²⁺-transient amplitudes increased above control levels [130]. (5) A rise in [H⁺]_CYT induces proton uptake into mitochondria via different transporter systems in exchange for Ca²⁺, Na⁺ or K⁺ ions (Figure 2, 8) [59]. Measurements using isolated mitochondria from ventricular myocytes showed that acidification of mitochondria reduces the electrochemical gradient that the mitochondrial uniporter uses for Ca²⁺ transport and limits mitochondrial Ca²⁺ uptake [143]. This effect may add to the elevation of [Ca²⁺]_CYT during acidosis. Interestingly, changes in pH_i in cardiac myocytes are not homogenous, as one would assume due to the high diffusion speed of protons in aqueous solutions [144], but may rather occur as pH gradients: intracellular pH measurements in isolated ventricular myocytes revealed that there is a subcellular heterogeneity of pH due to local interaction of protons with intracellular buffers, which slows down the diffusion speed of protons [145,146,147,148]. Consequently, pH gradients may locally enhance those inhibitory effects that protons have on Ca²⁺ handling proteins. If systemic acidemia cannot be treated, e.g., during chronic sepsis or kidney failure, then the pump function of the heart is impaired via two mechanisms. First, metabolic acidosis causes a fall in pH_o, which reduces organ perfusion, causing tissue hypoxia [149]. Second, intracellular acidosis depresses the contractile function of cardiac myocytes by the mechanisms described above. On the level of the human heart, this translated into reduced left ventricular contractility and reduced cardiac output in patients with systemic acidemia (pH_o < 7.28) as compared to a cohort of patients with normal blood pH (pH_o > 7.28). Among the functional parameters that were affected by intracellular acidosis were stroke volume (reduced by ~40%) and ejection fraction (reduced by ~30%) [150]. Both parameters directly determine cardiac output. Thus, alterations of cardiac pump functions due to intracellular acidosis may contribute to the poor survival rate that is associated with chronic acidemia [151].

If intracellular acidosis is reversible, excessive protons are removed from the cytosol of ventricular myocytes by the following mechanisms: activation of the voltage-gated proton channel H_V1 (Figure 2, 9; see Section 7) [152]; export via MCT (Figure 2, 10); Na⁺/H⁺-exchange by NHE (Figure 2, 11) and reaction with the cytosolic CO₂/bicarbonate buffer system [153]. In the mammalian heart, intracellular bicarbonate is provided by Na⁺/bicarbonate cotransport (NBC, Figure 2, 12) [154] and by the CA reaction, which generates bicarbonate by hydration of CO₂ stemming from mitochondrial respiration [119,155]. Bicarbonate leaves the cell by Cl⁻/bicarbonate anion exchange (AE, Figure 2, 13) [156]. In striated muscle, at low pH_i values, CAs catalyze the reverse reaction of protons with bicarbonate to H₂O and CO₂ that leaves the cell by diffusion (Figure 2, 14) [157]. CAs are not restricted to the cytoplasm and have been localized to different compartments, such as plasma membrane, mitochondria and the SR, where they regulate local pH [158]. Extracellular CAs in capillaries and erythrocytes reverse this reaction and hydrate CO₂ to bicarbonate and protons. The interplay between intracellular and extracellular CAs is important to maintain the CO₂ gradient and diffusion of CO₂ across the plasma membrane [159]. The CO₂/bicarbonate system contributes to ~20% of the total proton buffer capacity in skeletal muscle [160] and has a slightly larger buffer capacity in cardiac myocytes [161]. The importance of the CO₂/bicarbonate buffering system for cardiac function was directly demonstrated by the observation that the contractility of isolated murine myocytes and whole-heart preparations declined over time in the absence of a bicarbonate buffer due to an accumulation of intracellular protons, an effect that reversed when the same preparation was bathed in a CO₂/bicarbonate buffer and pH_i was normalized [156]. Despite contributing to buffering of protons, a recent study demonstrated that bicarbonate may emerge as an important signaling molecule that stimulates mitochondrial ATP production in isolated, contracting cardiac myocytes via a mechanism that is independent from mitochondrial Ca²⁺ [162]. Dysregulation of the CO₂/bicarbonate buffer system or proton transport processes have been implicated in skeletal muscle or cardiac pathologies. For example, upregulation of CAs represents a biomarker for cardiac ischemia and heart failure in humans [163,164]. During cardiac hypertrophy, upregulation of NBCs to rectify pH_i caused Na⁺ accumulation and Ca²⁺ overload via NCX [165].

Chronic fatigue syndrome is characterized by a high sympathetic tone, to which skeletal muscles respond with secretion of vasodilators, including protons, to improve muscle perfusion (“sympathetic escape” [166]). One factor that contributes to this mechanism is an upregulation of NHE that may induce Na⁺ overload and Ca²⁺-overload, which further deteriorates the contractile function of an already weak muscle [167]. In line with this, the pharmacological inhibition of NHE prevented muscle degeneration in a mouse model of Duchenne’s muscle dystrophy [168]. Based on results from several animal studies, an inhibition of NHE1 has been suggested as a potential novel treatment option to reduce tissue hypoxia and cardiac damage during metabolic acidosis in humans [169]. Despite affecting Ca²⁺ handling proteins, a fall in pH_i will also alter the properties of Ca²⁺ buffers and Ca²⁺ storing organelles, which we will discuss below.

5. Effect of Low pH_i on Ca²⁺ Buffers and Ca²⁺-Storing Organelles

In most cells Ca²⁺ ions are bound to ~99% to intracellular buffers [23]. To give an example, calculations for cardiac myocytes suggest that a change in free cytosolic Ca²⁺ from resting levels of 100 nM to ~1 μM during systole will require the net mobilization of as much as 100 μM Ca²⁺ from internal stores in the presence of internal buffers [31]. The simplest interference between protons and Ca²⁺ ions in the cytoplasm is their competition for the same binding sites at intracellular buffers [25]. The most relevant cytosolic Ca²⁺ buffer in type II (fast twitch) skeletal muscle fibers is the protein parvalbumin [170]. Parvalbumin is considered a slow buffer because it binds both Ca²⁺ and Mg²⁺ thus its Ca²⁺ buffering properties depend on competition with Mg²⁺ [171,172]. Considering this, Ca²⁺ buffering by parvalbumin does not limit the rapid rising phase and amplitude of Ca²⁺ transients during single twitches, but affects the decay of the transient and thus, the relaxation time of skeletal muscle [173]. In contrast, there is no relevant role for parvalbumin in type I (slow twitch) muscle fibers due to their slow relaxation [170]. In heart tissue, parvalbumin expression was species-specific [174], with low expression levels in human and larger animals and higher expression in small animals, according to the relaxation speed [175]. The most relevant Ca²⁺ buffers in cardiac cells are the Ca²⁺ binding sites of troponin C [176] and SERCA [177]. Computer models suggest that simple binding (buffering) of Ca²⁺ ions to troponin C and SERCA, followed by active transport of Ca²⁺ into the SR shape Ca²⁺ transients in the decay phase of the Ca²⁺ signal [177]. Because protonation lowers the K_d of buffers for Ca²⁺, it is plausible that intracellular acidification will liberate a substantial amount of Ca²⁺ into the cytosol, which may affect [Ca²⁺]_CYT during acidosis [178]. The effect that acidosis lowers the Ca²⁺ buffer capacity of parvalbumin has been shown in dorsal root ganglia neurons [179]; however this effect has not been established for striated muscle. For cardiac troponin C protonation decreases its affinity for Ca²⁺ [180], which should result in liberation of Ca²⁺ from contractile filaments during acidosis and impaired contractility.

Mitochondria are organelles that can buffer transient changes in cytoplasmic Ca²⁺ via Ca²⁺ diffusion through the mitochondrial uniporter (Figure 1, 5) [181,182] and can release mitochondrial Ca²⁺ into the cytosol by Na⁺/Ca²⁺ exchange [181,183]. In cardiac muscle, it is a matter of debate whether mitochondrial Ca²⁺ uptake occurs fast on a beat-to-beat basis or in an integrative manner as slow increase over time, dependent on the frequency of contractions [184]. It has been suggested that mitochondrial Ca²⁺ uptake may contribute to feedback mechanisms that couple cytoplasmic ATP demand with mitochondrial ATP production by activating Ca²⁺-sensitive enzymes that control the tricarboxylic acid cycle [55,185,186]. At the same time mitochondria maintain the proton gradient across the inner mitochondrial membrane by pumping out the protons via complexes I, III and IV of the respiratory chain, to provide, in combination with the mitochondrial membrane potential, the energy for ATP synthesis [187]. There is some experimental evidence that cytosolic pH_i and mitochondrial Ca²⁺ signaling can interfere: acidification of the cytoplasm was accompanied by proton uptake via Na⁺/H⁺ exchanger, K⁺/H⁺-exchanger and Ca²⁺/H⁺ exchanger and proton leak [188,189,190], leading to acidification of mitochondria, which inhibited mitochondrial Ca²⁺ uptake by the mitochondrial uniporter in mitochondria isolated from cardiac cells [143] (Figure 2, 7). In isolated, electrically paced cardiac myocytes, oscillatory changes in pH_i (“pH transients”) displayed kinetics that resembled the beat-to-beat changes in cytosolic Ca²⁺ transients [191]. The authors suggested as an underlying mechanism that Ca²⁺ uptake by mitochondria will induce simultaneous release of protons into the cytosol, which caused the observed fluctuations in pH_i. Those fluctuations were small (±0.1 pH units) and it remains to be established if they contribute to intracellular acidosis.

Lysosomes are traditionally seen as organelles involved in protein turnover and degradation of biomaterial (“waste removal”) [192,193] but not recognized as cellular compartments involved in Ca²⁺ signaling, even though they display high proton and Ca²⁺ contents [20,194]. Interference of protons with Ca²⁺ ions occurs during transport processes across the lysosomal membrane. Lysosomes maintain a low luminal pH ~4.5 to create a pH optimum for acid hydrolases [195]. In addition, they contain a considerably large amount of luminal Ca²⁺ (~500 μM) [194], which is partially released during vesicle fusion processes [196]. It has been suggested that lysosomes contribute to Ca²⁺ buffering in skeletal muscle cells and cardiac myocytes [197]. Lysosomal V-ATPases pump protons from the cytosol into the lumen of lysosomes [198]. This adds positive charges to an organelle with a very small lumen and to maintain the electrochemical gradient across the lysosomal membrane, any proton influx must be compensated either by import of anions, such as Cl⁻, or by export of cations, including Ca²⁺, via lysosomal ion channels and exchangers [199,200]. Even though the overall volume of lysosomes per cell volume is very small (~3%) [194], local release of lysosomal Ca²⁺ via type 2 two-pore channels has been implicated in triggering larger cytoplasmic Ca²⁺ release events from intracellular Ca²⁺ stores via CICR by modifying IP₃-Rs in non-excitable cells [201] or cardiac RyR2 signaling in atrial and ventricular myocytes [202]. In skeletal muscle cells, lysosomes form contact sites with the SR, mitochondria and the nuclear membrane, but the effector proteins of lysosomal signaling at those sites are not well defined [203].

Another organelle that can contribute to transport of protons into the cytosol is the SR. The pump cycle of Ca²⁺-ATPases requires charge compensation: whenever SERCA pumps Ca²⁺ ions into the SR, the added charge is compensated by antiport of two protons to the cytosol for each Ca²⁺ ion that enters the SR lumen [204]. Whether acidification affects [Ca²⁺]_CYT and ECC by Ca²⁺ release from buffers or cell organelles in striated muscle cells has not been addressed yet.

This complexity of local and global variations of pH requires the use of appropriate methods that are sensitive and specific to measure Ca²⁺ and pH in different cellular compartments with sufficient temporal and spatial resolution. The next chapter will give a short overview of established and recently developed optical tools that can be used to measure Ca²⁺ and pH in living cells.

6. Optical Methods to Measure Ca²⁺_CYT and pH_i in Living Cells

To study how changes in pH will affect Ca²⁺ signaling, fluorescent indicators have been used to directly monitor changes in pH or [Ca²⁺] in living cells. Fluorescent Ca²⁺ indicators are available as chemical compounds (Ca²⁺ dyes) that are loaded into cells or as genetically encoded biosensors based on fluorescent proteins, which are expressed by the cells. For both types of indicators, intensity-based and ratiometric variants are available [205,206]. There are advantages for the choice of either type for experiments: chemical indicators are well-established [207], require less experimental intervention and offer a wide range of affinities for Ca²⁺ ions [208]. The major advantage of genetically encoded probes is the possibility to target them to cell organelles, which allows simultaneous Ca²⁺ measurements in the cytosol and in organelles, such as ER or mitochondria [209]. In general, sensors with high affinity for Ca²⁺ are used in the cytosol, whereas sensors with low affinity for Ca²⁺ can be used to measure Ca²⁺ in organelles with high Ca²⁺ content, such as the SR or lysosomes. Popular organelle-specific Ca²⁺ biosensors based on fluorescent proteins are the indicators of the GECO family [209,210,211,212]. During experiments at constant pH, changes in fluorescence of the Ca²⁺ indicator reflect changes in [Ca²⁺]. However, the assessment of [Ca²⁺]_CYT during dynamic changes of pH_i is challenging, because most Ca²⁺ indicators are pH-sensitive: for example, in situ calibrations demonstrated that the K_d for Ca²⁺ of the widely used indicator fura-2 was approximately two-times lower at pH = 7.2 than at pH = 6.6, which means that a considerably larger Ca²⁺ concentration is required to cause a similar change in the emission ratio of fura-2 [30]. Being proteins, genetically encoded Ca²⁺ sensors bind protons as well and the intensity of emitted fluorescence depends on pH, a phenomenon that is either caused by direct quenching of the fluorophore [213] and/or due to competition of protons with Ca²⁺ ions at Ca²⁺ binding domains of those sensors. Thus, careful pH-calibration of Ca²⁺ sensors may be required for fluorescence measurements at different pH_i values. On the other hand, the generation of fluorescence intensity/pH curves established specific pKa values for each type of fluorescent protein used in biosensors [214]. This allows one to design fluorescent biosensors for specific applications, such as Ca²⁺ sensors that are bright enough at the very low pH values in lysosomes [215]. Comprehensive reviews on chemical Ca²⁺ indicators and Ca²⁺ biosensors are given by Paredes and colleagues [208] and by Mank and Griesbeck [216], respectively.

There is also a wide range of chemical indicators or protein-based biosensors for the optical assessment of pH_i available. One simple way to measure cytoplasmic pH is the use of enhanced YFP (eYFP, pKa = 7.1), which is pH-sensitive and gets dimmer when protonated [217]. As for Ca²⁺ measurements, using targeted fluorescent proteins allows measurements of pH changes within cell organelles [217]. An example of a chemical dye to measure pH_i is the widely used compound BCECF (pKa~7.0), which is a dual-excitation, single emission ratiometric indicator suitable for intracellular pH changes from pH = 6.5–7.5 [218]. Newer pH-sensitive fluorescent proteins include red-shifted variants of fluorescent proteins or ratiometric sensors and are engineered with enhanced quantum yield, improved dynamic range and tailored to match the specific pH values of organelles, such as the ER, the Golgi apparatus, mitochondria or lysosomes. A recent overview on protein-based pH-biosensors is given by Li and colleagues [214] and chemical pH indicators are described in detail by Han and Burgess [218]. For some experiments it may be useful to monitor pH_i and [Ca²⁺]_CYT simultaneously. This can be achieved by using combinations of Ca²⁺ indicators and pH indicators with different spectral properties, such as the Ca²⁺ indicators fura-2 or indo-1 in combination with the pH indicator SNARF, which does not show a high affinity for Ca²⁺ ions [219,220]. The advantage of this approach is that one can distinguish the kinetics of Ca²⁺ changes from the kinetics of pH changes in real time. These tools are necessary to quantify the degree of acidosis during physiological activity and pathological states of striated muscle cells and to correlate pH changes with functional changes. Transgenic animal models expressing genetically encoded biosensors can be used for long-term studies of acidosis in striated muscle cells.

7. Discussion and Outlook

Intracellular acidosis is a strong inductor of negative inotropy in striated muscle cells. We have introduced general mechanisms on how protons may alter the function of Ca²⁺ handling proteins and the contractile apparatus. Many molecular aspects of this phenomenon are still controversially discussed for the two types of striated muscle cells. Studying the effects of protons on Ca²⁺ signaling can be challenging because properties of Ca²⁺ indicators are sensitive to pH, which requires careful interpretation of acquired data. Moreover, data on skeletal muscle contractility seem to be highly sensitive to experimental conditions: some results obtained from experiments using isolated muscle cells (skinned or intact fibers), such as the degree of negative inotropy that individual metabolites will induce, could not be confirmed in intact skeletal muscle preparations [221,222]. Storage of tissue samples for future experiments may also affect the contractile properties of muscle fibers: a recent paper demonstrated that storage in glycerol, a commonly used freezing agent, affected the passive stiffness of skeletal fibers [223]. Furthermore, it seems critical whether results were obtained at room temperature, which is common for fiber studies, or at body temperature, which is used for muscle preparations or studies in vivo [224]. This has led to some skepticism about how well single cell data may be extrapolated to the situation in intact muscle [221,222], thus further investigations will be required to address ischemia in the whole organ. During cardiac ischemia, excessive production of ROS affects cardiac excitability and adds to the negative inotropic effect as many Ca²⁺ handling proteins are redox-sensitive [113,114]. Experimentally, it may be difficult to separate proton-induced effects from ROS-mediated alterations of Ca²⁺ handling proteins in intact organ preparations.

An interesting outlook for future studies lies in alternative signaling pathways that cells can use to sense changes in pH: many cell types that experience sudden intracellular acidification can rectify intracellular acidosis by activation of proton channels [82]. Proton channel currents have been identified in skeletal myotubes [225] and adult skeletal muscle cells [226], but their physiological role is not well defined. The expression of proton channel mRNA in cardiac tissue has been described for multiple species [227], and there is some indication for upregulation of cardiac proton channel transcripts during heart failure [228]. Recently, functional currents of the voltage-gated proton channel H_V1 were recorded in adult canine ventricular myocytes and pharmacological blockade of this channel in the absence of NHE activity lowered pH_i from 7.1 to 6.13 in beating myocytes [152]. Thus, H_V1 contributes to the regulation of pH_i in ventricular myocytes when [H⁺]_CYT rises (Figure 2, 10), an alternative pathway to proton exchange by NHE [229,230]. The advantage of a voltage-gated proton channel, compared to a transporter like NHE, lies in its exclusive ability to conduct protons out of the cell, driven by the electrochemical gradient. In all primary mammalian cells measured to date, it does not permit proton influx [231]. Thus, during acidification, it requires no additional energy while counteracting the drop in cytosolic pH. This is particularly beneficial for excitable cells, as the channel expels protons from the cytosol during depolarization, which also serves as the gating mechanism for the channel. Depolarization typically occurs just before a rise in energy demand in muscle cells, when metabolically generated protons begin to accumulate. The mechanism underlying this unique biophysical property—functioning as a proton “overpressure valve”, exclusively conducting protons out of the cell—is still under investigation [232]. Nonetheless, voltage-gated proton channels have been identified in various mammalian cell types [233]. Another notable feature is the absolute selectivity for protons, with no other ions conducted [234]. A frequently overlooked capability of the voltage-gated proton channel is its ability to sense both intracellular and extracellular pH, effectively making it a cellular “pH meter.” In line with the topic of this review, it is worth noting that the first scientists to voltage-clamp this channel also investigated the relationship between intracellular calcium and pH in excitable cells [235].

Other membrane proteins that cells use to sense pH_o include Ca²⁺-conducting cation channels (ASIC- and TRP channels) [236], and the rather novel, but small class of proton-sensing, G-protein-coupled receptors (GPCRs) [237]. Some of those receptors couple to heterotrimeric G_q proteins and activate phospholipase C (PLC), which controls intracellular Ca²⁺ signals by activating IP₃-mediated Ca²⁺ release from the SR [15]. The PLC-IP₃-Ca²⁺ signaling axis controls gene transcription [238,239] and could potentially contribute to long-term adaptation processes of muscle cells to acidification. In skeletal muscle cells, the functional roles of conventional GPCRs or proton-sensing GPCRs are not well investigated, but there is evidence that some orphan receptors could belong to the class of proton-sensing GPCRs [240]. In the heart transcripts of GPR68, a proton-sensing GPCR which activates PLC, have been identified in neonatal ventricular myocytes [236] and adult tissue. Thus, proton-sensing GPCRs have been suggested as novel drug targets to treat ischemic heart diseases [241].

8. Conclusions

In conclusion, although the functional consequences of intracellular acidosis on the contractility of striated muscle cells are well established, the molecular aspects of proton biochemistry underlying acidosis remain unclear. There are interesting, unexplored signaling pathways that could link acidification during ischemia with Ca²⁺ signaling and gene transcription in striated muscle cells. The continuous development of fluorescent biosensors to assess Ca²⁺ and pH_i in living cells will prove useful to investigate those aspects in the future.