Survey
* Your assessment is very important for improving the workof artificial intelligence, which forms the content of this project
* Your assessment is very important for improving the workof artificial intelligence, which forms the content of this project
Heat exchanger wikipedia , lookup
Dynamic insulation wikipedia , lookup
Copper in heat exchangers wikipedia , lookup
Solar air conditioning wikipedia , lookup
R-value (insulation) wikipedia , lookup
Heat equation wikipedia , lookup
Cogeneration wikipedia , lookup
Bioscience Reports, Vol. 21, No. 2, April 2001 ( 2001) MINI REVIEW Role of the Sarcoplasmic Reticulum Ca2+-ATPase on Heat Production and Thermogenesis Leopoldo de Meis1 Receiûed October 26, 2000 The sarcoplasmic reticulum of skeletal muscle retains a membrane bound Ca2+-ATPase which is able to interconvert different forms of energy. A part of the chemical energy released during ATP hydrolysis is converted into heat and in the bibliography it is assumed that the amount of heat produced during the hydrolysis of an ATP molecule is always the same, as if the energy released during ATP cleavage were divided in two non-interchangeable parts: one would be converted into heat, and the other used for Ca2+ transport. Data obtained in our laboratory during the past three years indicate that the amount of heat released during the hydrolysis of ATP may vary between 7 and 32 kcal兾mol depending on whether or not a transmembrane Ca2+ gradient is formed across the sarcoplasmic reticulum membrane. Drugs such as heparin and dimethyl sulfoxide are able to modify the fraction of the chemical energy released during ATP hydrolysis which is used for Ca2+ transport and the fraction which is dissipated in the surrounding medium as heat. KEY WORDS: Ca2+-ATPase; Ca2+ transport; energy interconversion; ATP hydrolysis; heat production; sarcoplasmic reticulum THERMOGENESIS Heat generation and burning calories are implicated in the regulation of several physiological processes including body temperature, metabolism, body weight, energy balance and cold acclimation. At least two different systems are known to be involved in the process of nonshivering thermogenesis and these are the uncoupling proteins (UCP) and the sarcoplasmic reticulum Ca2+-ATPase of skeletal muscle. In both systems the process of heat dissipation is initiated by the leakage of ions through the membrane, protons in the case of the UCPs and Ca2+ in the case of the Ca2+-ATPase [1–7]. Different uncoupling protein (UCP) isoforms have already been identified [4– 7]. These include UCP1 specific for brown adipose tissue, UCP2 found in most tissues and UCP3 which is highly expressed in skeletal muscle. From the three isoforms, only UCP1 is clearly involved in heat production. The physiological role of UCP2 and UCP3 is still controversial [4–7]. The different UCP isoforms promote 1 Instituto de Ciências Biomédicas, Departamento de Bioquı́mica Médica, Universidade Federal do Rio de Janeiro, Cidade Universitária, RJ 21941-590, Brazil. 113 0144-8463兾01兾0400-0113$19.50兾0 2001 Plenum Publishing Corporation 114 de Meis the dissipation of the proton electrochemical gradient formed across the inner mitochondrial membrane during respiration. In order to restore the gradient and to prevent the decrease of the cytosolic ATP concentration, the proton leakage promoted by the UCP leads to increased mitochondrial respiration, increase fatty acid oxidation and heat production. Skeletal muscle is by far the most abundant tissue of the human body and accounts for over 50% of the total oxygen consumption in a resting human and up to 90% during very active muscular work. Calorimetric measurements of rat soleus muscle indicate that 25–45% of heat produced in resting muscle is related to Ca2+ recirculation between sarcoplasm and sarcoplasmic reticulum [8]. Ca2+ leakage from the reticulum is accompanied by an increase of the Ca2+-ATPase activity needed to pump back Ca2+ into the reticulum. In steady state conditions the Ca2+-ATPase is able to synthesize ATP from ADP and Pi using the energy derived from the Ca2+ gradient [9–11] but the amount of ATP synthesized is much smaller than the amount of ATP cleaved and therefore an increase of Ca2+ leakage from the reticulum ultimately leads to increased mitochondrial respiration to maintain the cytosolic ATP concentration [3, 4]. Conditions that promote a change of the rate of heat production in animals are usually associated with changes of expression of both, the sarcoplasmic reticulum Ca2+-ATPase and UCP proteins. Thus, during cold adaptation, UCP1, and UCP2, but not UCP3 are overexpressed [6, 7, 12–14]. Similarly, in cold-acclimated ducklings, there is a 30–50% increase of both sarcoplasmic reticulum Ca2+-ATPase and ryanodine-sensitive Ca2+ release channels. In cold-acclimated ducklings 70% of the total heat production is derived from muscle [15, 16]. The expression of both sarcoplasmic reticulum Ca2+-ATPase and UCP1 are decreased in hypothyroid rats. In these animals, the injection of the thyroid hormone 3,5,3′-triiodo L-thyronine (T3) increases the expression of both Ca2+-ATPase and UCP1 [12, 17–20]. A curious system that highlights the importance of the sarcoplasmic reticulum Ca2+-ATPase in heat production is the heater tissues of billfishes. In marlin and swordfish, ocular muscles are transformed into specialized heater tissues [3]. During the daily fluctuations in temperature, the swordfish reduces the temperature changes experienced by the brain and retina by warming these tissues with the heater organ. The heater tissues are composed of modified muscle cells in which the contractile filament is virtually absent and the cell volume is packed with mitochondria and a highly developed sarcoplasmic reticulum. Activation of thermogenesis seems to be associated with the ATP-dependent cycling of Ca2+ at the sarcoplasmic reticulum. Mitochondrial respiration is then stimulated by cytosolic ADP generated by the Ca2+-ATPase, the result being increased heat production. Alteration of thermogenesis is observed in different pathological conditions as for instance obesity and the hypothermia noted during ischemia. Obesity results from a chronic imbalance between energy intake (feeding) and energy expenditure. Ischemia and hypoxia elicits hypothermia as a compensatory mechanism to oxygen deprivation. Obesity affects more than one-third of the U.S. population and is a major public health concern because it is associated with diabetes, hypertension and cardiovascular disease. In different studies it was shown that some humans resist fat gain with overeating, whereas others readily store excess fat [21–23]. The resistance Heat Production by the Ca2+-ATPase 115 to fat gain was found to vary up to 10-fold in volunteers that underwent a supervised overfeeding [24]. The interindividual variation in weight gain suggests that a thermogenic mechanism involving an increase of heat production could be activated in some individuals to prevent weight gain or obesity [21–24]. A different mechanism of temperature control is activated during ischemia or hypoxia. In this condition there is a temperature decrease in the cells deprived of oxygen. The strategy seems to be to decrease the oxygen requirement by lowering the metabolic activity of the tissue. The hypothermia induced by hypoxia can be promoted in one specific organ, particularly in oxygen sensitive organs such as the heart and brain [25–27], or it can lead to a decrease of the whole body temperature when animals are exposed to an environment having a low oxygen concentration. This is observed in different animal species and in mice, severe hypoxia may lead to a decrease of body temperature of more than 5°C [28, 29]. In this review we will focus on the mechanism of heat production by the Ca2+ATPase of the sarcoplasmic reticulum. ENERGY INTERCONVERSION BY THE SARCOPLASMIC RETICULUM Ca2+-ATPase The Ca2+-ATPase found in the membrane of the sarcoplasmic reticulum of skeletal muscle is able to interconvert different forms of energy (Fig. 1). The Ca2+ATPase translocates Ca2+ from the cytoplasm to the lumen of the reticulum by using the chemical energy derived from ATP hydrolysis (route 1 in Fig. 1). After that Ca2+ is accumulated inside the reticulum, and a Ca2+ gradient is formed across the membrane and this promotes the reversal of the catalytic cycle of the enzyme during which Ca2+ leaves the reticulum in a process coupled with the synthesis of ATP from ADP and Pi (route 2). During reversal, the osmotic energy derived from the gradient is transformed by the enzyme back into chemical energy. In conditions similar to Fig. 1. Ca2+ translocation through the membrane of (A) intact and (B) leaky vesicles. In the figure (1) represents Ca2+ uptake, (2) Ca2+ efflux coupled to the synthesis of ATP and (3) uncoupled Ca2+ efflux. 116 de Meis those found in the living cell at rest, a steady state is established during which the Ca2+ concentration is high inside the reticulum and low in the cytosol and the pump operates forward and backwards, cleaving and synthesizing ATP continuously. In the bibliography, the simultaneous synthesis and hydrolysis of ATP measured in steady state conditions is referred to as the ATP ↔Pi exchange reaction [30–35]. During the ATP↔Pi exchange only part of the Ca2+ efflux is coupled with the synthesis of ATP [36–44]. The other part leaks through the Ca2+-ATPase without promoting the synthesis of ATP, a process referred to as an uncoupled efflux (route 3 in Fig. 1). The rates of the coupled and the uncoupled Ca2+ effluxes can be modified by different drugs [42–44]. Only a part of the chemical energy released during the hydrolysis of ATP is converted into other forms of energy such as osmotic energy. The other part is converted into heat, and this is used by the cell to maintain a constant and high body temperature. Nonshivering thermogenesis is a key component of temperature regulation in animals having little or no brown adipose tissue. During nonshivering thermogenesis most of the heat is derived from resting muscles but the mechanism of heat production is still unclear. It has been proposed that Ca2+ leaks from the sarcoplasmic reticulum and heat would then be derived from the hydrolysis of the extra amount of ATP needed to maintain a low myoplasmic Ca2+ concentration. In this formulation it is assumed that the amount of heat produced during the hydrolysis of an ATP molecule is always the same and is not modified by the formation of the gradient, as if the energy released by ATP hydrolysis were to be divided in two non-interchangeable parts: one would be converted into heat, and the other used for Ca2+ transport [2–4]. Data obtained in our laboratory during the past three years [45–49] indicate that the amount of heat produced during the hydrolysis of each ATP molecule varies depending on whether or not a Ca2+ gradient is formed across the reticulum membrane. In presence of the gradient the heat produced during the hydrolysis of each ATP molecule is two to three times larger than that measured in absence of a gradient and at least a part of this extra heat seems to be associated with the uncoupled Ca2+ efflux. The coupled and the uncoupled Ca2+ efflux may therefore represent two distinct routes of energy conversion, both mediated by the Ca2+-ATPase in which the osmotic energy derived from the Ca2+ gradient is either used to synthesize ATP (coupled Ca2+ efflux—route 2 in Fig. 1) or is dissipated into the medium as heat (uncoupled Ca2+ efflux—route 3 in Fig. 1). HEAT PRODUCTION AND ATP SYNTHESIS BY THE Ca2+-ATPase A transmembrane Ca2+ gradient is formed when intact vesicles derived from the sarcoplasmic reticulum of rabbit white muscle are incubated in a medium containing ATP and Mg2+. This is not observed with leaky vesicles because the Ca2+ transported across the membrane readily diffuses back to the assay medium (Fig. 1B and Table 1). Both in the presence and absence of a Ca2+ gradient the amount of heat produced during the hydrolysis of ATP was found to be proportional to the amount of ATP hydrolyzed (Fig. 2A). However, in the presence of the gradient, the amount of heat released after the hydrolysis of each ATP molecule was found to be larger than that measured with leaky vesicles. As a result the value of the calorimetric enthalpy for Heat Production by the Ca2+-ATPase 117 Table 1. Ca2+ Uptake, ATP Hydrolysis, ATP Synthesis and Heat Production by Rabbit Sarcoplasmic Reticulum Vesicles Additions Leaky (no gradient) LeakyC20% DMSO Intact (gradient) IntactC20% DMSO IntactC3 µg兾ml heparin n Ca2+ uptake, µmol兾mg ATP hydrolysis µmol兾mg ATP synthesis µmol兾mg Heat released mcal兾mg ∆Hcal, kcal兾mg 5 5 5 6 4 None None 3.4J0.5 4.9J0.2 0.9J0.2 28.4J1.6 8.0J1.1 17.8J3.0 9.5J1.9 9.8J0.5 None None 0.5J0.1 0.8J0.1 0.3J0.1 286.5J27.6 91.8J8.7 391.7J36.1 102.5J11.3 297.1J21.8 −10.2J1.3 −11.4J1.1 −21.9J1.5 −10.8J1.4 −30.2J2.1 The reaction was performed at 35°C and the incubation time was 20 min. The assay medium composition was 50 mM MOPS兾Tris buffer, pH 7.0, 0.1 mM CaCl2 , 1 mM ATP, 4 mM MgCl2 , and 10 mM Pi . The medium was divided into four samples. One was used for heat measurements. To the other three samples trace amounts of either 45Ca, [ γ -32P]ATP or 32Pi were added for measurement of Ca2+ uptake, ATP hydrolysis and ATP synthesis, respectively. The four reactions were started simultaneously by the addition of vesicle protein (20 µg兾ml). The values in the table are the average ±SE. In the table (n) is the number of experiments and DMSO is dimethyl sulfoxide. The calorimetric enthalpy (∆Hcal) was calculated by dividing the amount of heat released by the amount of ATP cleaved by the vesicles. For experimental details see Refs. 45 and 47. ATP hydrolysis (∆Hcal) measured within intact vesicles was more negative than that measured with leaky vesicles (Fig. 2B and Table 1 compare ∆Hcal values of leaky and intact vesicles in the absence of DMSO). The ∆Hcal value was calculated by dividing the amount of heat released by the amount of ATP cleaved by the vesicles. A negative value indicates that the reaction was exothermic and a positive value Fig. 2. Heat release during ATP hydrolysis in presence and absence of a transmembrane Ca2+ gradient. The assay medium and experimental conditions were as in Table 1 using (䊊) intact vesicles that accumulated Ca2+ and formed a transmembrane Ca2+-gradient and (●) leaky vesicles which were not able to retain Ca2+ inside the vesicles. The figure shows a typical experiment where the amount of heat released (A) and the ∆Hcal values (B) were plotted as a function of the amount of ATP cleaved at different incubation intervals. The calorimetric enthalpy (∆Hcal) was calculated by dividing the amount of heat released by the amount of ATP cleaved by the vesicles. Note that the more negative the values of ∆Hcal, the more heat was released during the hydrolysis of ATP. For experimental details see Ref. 45. 118 de Meis indicates that it was endothermic. This difference suggests that the vesicles were able to convert a part of the osmotic energy derived from the gradient into heat and the possibility was raised that the conversion could be mediated by the uncoupled leakage of Ca2+ through the ATPase (route 3 in Fig. 1A). According to this reasoning it would be expected that drugs which change the rate of the uncoupled Ca2+ efflux should also change the amount of ATP synthesized and the amount of heat produced during ATP hydrolysis i.e., the ∆Hcal of ATP hydrolysis. Dimethyl sulfoxide (DMSO) is able to arrest the uncoupled Ca2+ efflux [11, 38, 40, 50]. As a result, more Ca2+ is accumulated and less ATP is cleaved by the vesicles. On the other hand, the enhancement of Ca2+ uptake results in an increase of the transmembrane Ca2+ gradient, a condition that favors the reversal of the catalytic cycle of the enzyme and therefore, an increase of the rate of ATP synthesis from ADFP and Pi . We observed [45, 47] that the decrease of the uncoupled Ca2+ efflux promoted by DMSO is associated with a decrease of heat production and the ∆Hcal value increases to the same value as that measured with leaky vesicles (Table 1). Note that DMSO did not change the ∆Hcal measured with leaky vesicles in the presence of 0.1 mM CaCl2 (Table 1). Heparin is an uncoupling drug that inhibits both the synthesis and the hydrolysis of ATP and increases the uncoupled leakage of Ca2+ through the Ca2+ATPase [43–45, 51]. In the presence of 3 µg兾ml heparin the vesicles still retained a small amount of Ca2+ and despite the significant decrease of the ATPase activity, the heat released during the different incubation intervals was similar to that measured with the control without heparin, i.e., more heat was produced for each ATP cleaved. Thus, the ∆Hcal measured with 3 µg兾ml heparin was significantly more negative than that of the control (Table 1). The degree of leakage increased when the heparin concentration was raised to 10 µg兾ml and although the vesicles were still able to hydrolyze ATP, they were no longer able to accumulate Ca2+ and form a transmembrane Ca2+ gradient. This promoted a decrease in the ∆Hcal to the same value as that measured with leaky vesicles [45, 47]. CONTROL OF HEAT PRODUCTION IN ABSENCE OF A TRANSMEMBRANE Ca2+ GRADIENT The Ca2+-ATPase seems to be able to modulate the ∆Hcal of ATP hydrolysis even in the absence of a transmembrane Ca2+ gradient. In this case however, the amount of heat produced during ATP hydrolysis was always smaller than that measured with intact vesicles (without DMSO in Table 1). Leaky vesicles can catalyze both the hydrolysis and the synthesis of ATP when the Ca2+ concentration in the medium is raised to a level similar to that found inside the vesicles when a gradient is formed (about 2 mM) [33, 35, 52–54]. This promotes both a decrease of the rate of ATP hydrolysis and activation of the synthesis of ATP (Fig. 3 and Table 2). MSO is known to propitiate the reversal of the catalytic cycle [50, 55], decreasing the ratio between the velocities of ATP hydrolysis and of ATP synthesis. With the use of leaky vesicles a small decrease of the ∆Hcal was detected when the Ca2+ concentration in the medium was raised from 0.1 to 2.0 mM (Fig. 3 and Table 2). The decrease was more pronounced when the fraction of ATP synthesized from ADP and Pi was enhanced due to the addition of DMSO to the assay medium. This finding suggests that part of the chemical energy derived from the hydrolysis of ATP is retained by Heat Production by the Ca2+-ATPase 119 Fig. 3. Heat release (A) and ATP resynthesis (B) during ATP hydrolysis. The assay medium composition was 50 mM MOPS兾Tris buffer pH 7.0, 1 mM ATP, 0.1 mM ADP, 4 mM MgCl2 , 10 µM A23187, 20% dimethyl sulfoxide, 2 mM Pi and either (䊊) 0.1 or (●) 2.0 mM CaCl2 . The reaction was started by the addition of rabbit leaky vesicles. The reaction was performed at 25°C. The values represent the averageJSE of four experiments. For experimental details see Ref. 46. Table 2. Heat Released and ATP Synthesis in the Absence of Ca2+ Gradient ATP, µmol兾mg · 20 min−1 Condition (a) 0.1 mM CaCl2 (b) 2.0 mM CaCl2 (c) 2.0 mM CaCl2 without added Pi Hydrolysis Synthesis ∆Hcal (kcal兾mol Pi ) 5.34J0.68 2.31J0.29 0 0.37J0.03 −12.25J0.25 −7.78J0.23* 2.43J0.36 0 −11.93J0.18 The reactions were performed at 25°C with rabbit leaky vesicles. The assay medium composition was 50 mM MOPS-Tris pH 7.0, 1 mM ATP, 0.1 mM ADP, 4 mM MgCl2 , 10 µM A23187, 20% (v兾v) DMSO and either with 2 mM Pi (a and b) or without added Pi (c). Other conditions were as described in the legend to Fig. 1. The values shown in the table represent the average ±SE of either seven (a, b) or three experiments (c). Note that negative values of ∆Hcal indicates that the reaction was exothermic. The differences between the ∆Hcal values for ATP hydrolysis measured with 0.1 and 2.0 CaCl2 (*) was significant (t-test) with pF0.001. For experimental details see Ref. 46. the enzyme and can be used to either synthesize more ATP or it can be dissipated as heat and the selection between the two routes would be determined by both the Ca2+ concentration in the medium and by the presence of Pi , in one of the substrates needed for the synthesis of ATP. EFFECT OF TEMPERATURE: Ca2+ TRANSPORT AND HEAT PRODUCTION BY ENDOTHERMIC (RABBIT) AND POIKILOTHERMIC (TROUT) ANIMALS This was explored using vesicles derived from the sarcoplasmic reticulum vesicles of rabbit white muscle and trout muscle [47]. The activity of the two vesicle 120 de Meis Table 3. Energy Interconversion by the Rabbit and Trout Ca2+-ATPase at Different Temperatures Ca2+ uptake µmol兾mg ATP hydrolysis µmol兾mg · min−1 ATP synthesis µmol兾mg · min−1 ∆Hcal kcal兾mol Pi Rabbit 35°C 25°C 3.25J0.41 (6) 1.54J0.10 (9) 0.89J0.15 (6) 0.42J0.05 (9) 0.03J0.08 (6) 0.09J0.01 (9) −20.78J1.33 (41) −11.53J0.54 (17) Trout 25°C 15°C 1.42J0.14 (12) 0.94J0.11 (5) 0.67J0.10 (12) 0.40J0.06 (5) 0.028J0.02 (12) 0.010J0.001 (5) −21.7J1.15 (18) −11.1J0.69 (9) Animal and temperature Values are meansJSE of the number of experiments shown in parentheses. Assay medium composition and other experimental conditions were as described in Table 1. Ca2+ uptake values are not initial velocities but steady-state level reached after 40 min incubation. For experimental details see Ref. 47. preparations increases with the temperature and after 40 min incubation at 25°C the amounts of Ca2+ retained by the rabbit and trout vesicles are practically the same (Table 3). The trout Ca2+-ATPase is unstable at temperatures higher than 25°C and is inactivated after a few minutes incubation at 35°C [56]. The rabbit ATPase however, is stable for more than one hour at 35°C. The physiological body temperature of the trout varies between 20° and 25°C while the rabbit is 37°C. Thus, in spite of the fact that the two enzymes can pump similar amounts of Ca2+ at 25°C, at the physiological body temperature the rabbit sarcoplasmic reticulum is able to pump more Ca2+ (Table 3) and at a faster rate than the reticulum of the trout. After formation of the gradient both the rabbit and the trout Ca2+-ATPases are able to synthesize a small amount of ATP and in all conditions tested, the rate of synthesis is 25 to 45 times smaller than the rate of ATP hydrolysis (Table 3). Both, in the presence and in the absence of a Ca2+ gradient, the amount of heat released is proportional to the amount of ATP hydrolyzed. This can be visualized plotting either the heat released as a function of the amount of ATP hydrolyzed (Fig. 2) or calculating the ∆Hcal at each incubation interval (Table 3 and Fig. 4). The heat released for each ATP molecule hydrolyzed varies depending on the temperature of the assay and the source of the vesicles used. For the rabbit, the value of ∆Hcal measured at 35°C with intact vesicles is double that measured with leaky vesicles. This difference is no longer detected when the temperature is decreased to 25°C as if, in the rabbit, the mechanism that converts osmotic energy into heat production would be turned off when the temperature is decreased to a level far away from the physiologic body temperature (Table 3 and Fig. 4). For the trout vesicles (poikilotherm), formation of a transmembrane Ca2+ gradient at 25°C leads to a change of the ∆Hcal for ATP hydrolysis to a value similar to that measured with the rabbit vesicles at 35°C. The difference of ∆Hcal values measured with trout vesicles in the presence and absence of a Ca2+ gradient is also abolished when the temperature of the medium is decreased but in this case, to a value below 17°C. The ∆Hcal measured with leaky vesicles did not vary with the temperature nor with the source of the vesicles used (Fig. 4). These data indicate that the amount of heat produced during ATP hydrolysis by the Ca2+ATPase increases when a gradient is formed across the sarcoplasmic reticulum membrane regardless of whether trout or rabbit were used. The gradient dependent heat production however, seems to be arrested when the temperature of the medium is Heat Production by the Ca2+-ATPase 121 Fig. 4. Effect of gradient and temperature on the ∆Hcal of ATP hydrolysis. The assay media and experimental conditions were as in Table 3 using trout (■, 䊐) and rabbit (䊊, ●) vesicles and either intact vesicles (solid symbols) or leaky vesicles (open symbols). For experimental details see Ref. 47. decreased more than 5°C below the physiological body temperature, i.e., below 30°C for the rabbit and below 20°C for the trout. The enhancement of heat production associated with the gradient could therefore play a physiological role in the maintenance of the body temperature but would not be a good emergency system to raise the body temperature after rapid cooling of the animal to an extreme point that leads to a large variation of the body temperature. Ca2+ TRANSPORT AND HEAT PRODUCTION BY DIFFERENT Ca2+-ATPase ISOFORMS Three distinct genes encode the sarco兾endoplasmic reticulum Ca2+-ATPases (SERCA) isoforms, but the physiological meaning of isoform diversity is not clear. The SERCA 1 gene is expressed exclusively in fast skeletal muscle whereas blood platelets and lymphoid tissues express SERCA 3 and SERCA 2b genes [57–60]. The catalytic cycle of the different SERCA can be reversed after a Ca2+ gradient has been formed across the vesicles membrane and all of them are able to synthesize ATP from ADP and Pi during Ca2+ transport [30–35, 48, 52]. The vesicles derived from blood platelets endoplasmic reticulum are able to accumulate a smaller amount of Ca2+ than the vesicles derived from muscle (Table 4). During transport the two vesicle preparations catalyze simultaneously the hydrolysis and the synthesis of ATP from ADP and Pi . In both muscle and blood platelet vesicles, the rate of synthesis was several fold slower than the rate of hydrolysis. Using the same experimental conditions as those described in Table 4 and in presence of 1 µM free Ca2+, the rates of ATP synthesis for platelets and muscle vesicles were 0.08J0.01 [6] and 2.57J0.22 [4] µmole of ATP兾mg protein · 30 min−1 respectively. These values are the average ±SE of the number of experiments shown in parentheses. As for the muscle vesicles 122 de Meis Table 4. Energy Interconversion by the Ca2+-ATPase of Rabbit Sarcoplasmic Reticulum and Human Blood Platelets Endoplasmic Reticulum Ca2+ uptake µmol兾mg ATP hydrolysis µmol兾mg Heat release, mcal ∆Hcal kcal兾mol Pi Skeletal muscle Ca2+, zero 1 µM 10 µM — 1.85J0.16 (5) 2.65J0.43 (5) 2.1J0.1 (11) 40.3J2.5 (5) 46.1J2.2 (5) 20.6J2 (11) 1270.4J62.9 (5) 1054.1J147.4 (5) −10.20J1.38 (11) Blood platelets Ca2+, zero 1 µM 10 µM — 0.14J0.02 (3) 0.22J0.02 (9) 0.5J0.1 (7) 1.8J0.3 (4) 1.6J0.2 (15) 4.2J0.9 (7) 15.9J0.75 (4) 17.1J2.6 (15) −12.30J0.71 (7) −9.91J1.93 (4) −10.99J1.09 (15) Vesicles and Ca2+ concentration −31.88J1.22 (5) −22.67J2.14 (5) The incubation time at 35°C was 30 min. The assay medium composition was 50 mM MOPS兾Tris buffer (pH 7.0), 4 mM MgCl2 , 100 mM KCl, 1 mM ATP, 5 mM NaN3 , 10 mM Pi , and 5 mM EGTA (zero Ca2+) or 0.1 mM EGTA and either 0.063 or 0.112 CaCl2 . The calculated free Ca2+ concentration with these different mixtures of EGTA and CaCl2 were 1 and 10 µM respectively. Values are the meanJSE of the number of experiments shown in parentheses. For experimental details see Ref. 48. (SERCA 1), Ca2+ transport by the vesicles derived from blood platelets endoplasmic reticulum (SERCA 3 and 2b) is exothermic and the amount of heat released during the different incubation intervals was proportional to the amount of ATP cleaved [48]. This could be visualized calculating the ∆Hcal using the values of heat release and Pi produced at different incubation intervals (Fig. 5). Two different ATPase activities can be distinguished in both platelet and muscle vesicles. The Mg2+-dependent activity requires only Mg2+ for its activation and is measured in the presence of EGTA to remove contaminant Ca2+ from the assay medium. The ATPase activity Fig. 5. Effect of Ca2+ gradient on the ∆Hcal values measured with platelet (A) and skeletal muscle (B) vesicles. The experimental conditions were as described in Table 4. The free Ca2+ concentrations were (䉭) zero, (●) 1 µM and (䊊) 10 µM. For experimental details see Ref. 48. Heat Production by the Ca2+-ATPase 123 which is correlated with Ca2+ transport requires both Ca2+ and Mg2+ for full activity [33, 34]. In both vesicle preparations, the Mg2+-dependent ATPase activity represents a small fraction of the total ATPase activity measured in presence of Mg2+ and Ca2+. The amount of heat produced during the hydrolysis of ATP by the Mg2+-dependent ATPase was the same regardless of whether muscle or platelet vesicles were used and the ∆Hcal value calculated in the two conditions (Table 4) was the same as that previously measured with soluble F1 mitochondrial ATPase [46] and soluble myosin at pH 7.2 [61]. For the vesicles derived from muscle (SERCA 1) the formation of a Ca2+ gradient increased the yield of heat production during ATP hydrolysis. This was not observed with the use of platelet vesicles (SERCA 2b and 3) where the yield of heat produced during ATP cleavage was the same in presence and absence of a transmembrane Ca2+ gradient (Fig. 5 and Table 4). For the muscle vesicles there was no difference in the ∆Hcal value of the Mg2+-dependent ATPase and the Ca2+-ATPase when the vesicles were rendered leaky (compare values of no gradient in Table 1 and zero Ca2+ in Table 4). With intact vesicles, the ∆Hcal value was more negative, i.e., more heat was produced during the hydrolysis of each ATP molecule when the free Ca2+ concentration in the medium was decreased from 10 to 1 µm (Fig. 5 and Table 4). During transport, the Pi available in the assay medium diffuses through the membrane of both muscle and platelet vesicles, to form Ca2+ phosphate crystals inside the vesicles. These crystals operate as a Ca2+ buffer that maintains the free Ca2+ concentration inside the two vesicles constant (∼5 mM) at the level of the solubility product of calcium phosphate [10, 62]. The energy derived from the gradient depends on the difference between the Ca2+ concentrations inside and outside the vesicles. The different values of ∆Hcal measured with the muscle vesicles with 1 and 10 µM Ca2+ suggest that when the free Ca2+ concentration in the medium is lower, the gradient formed across the vesicles membrane is steeper; thus more osmotic energy would be available, more heat was produced and a more negative value of the ∆Hcal for ATP hydrolysis is observed. With vesicles derived from blood platelets, there is no extra heat production during Ca2+ transport regardless of the free Ca2+ concentration in the medium (Table 4). During transport, the free Ca2+ concentration in the lumen of the platelet vesicles is the same as that of the muscle (∼5 mM). Thus, the Ca2+ gradient formed across the membrane in the presence of 1 and 10 µM Ca2+ should be the same in the two vesicles preparations, but only in muscle vesicles the Ca2+ gradient increases the yield of heat production during ATP hydrolysis. These findings indicate that different from the muscle, the Ca2+-ATPase of blood platelets is not able to convert the osmotic energy derived from the gradient into heat. UNCOUPLED Ca2+ EFFLUX Kinetic experiments indicate that a part of the extra heat measured after the formation of a gradient in muscle vesicles is related to the uncoupled Ca2+ efflux mediated by the Ca2+-ATPase [45, 47, 49]. This can be measured arresting the pump by the addition of an excess EGTA to the medium (Fig. 6). In this condition, the free calcium available in the medium is chelated but ATP and other reagents remain at the same concentration as those used for measurements of ATP hydrolysis and 124 de Meis Fig. 6. Ca2+ release from skeletal muscle vesicles. The assay medium composition was 50 mM MOPS兾Tris pH 7.0, 2 mM MgCl2 , 1 mM ATP, 0.1 mM CaCl2 , 20 mM Pi . The reaction was started by the addition of vesicles at 35°C. (䊊) control without additions. The arrow indicates the addition of either 5 mM EGTA (∆) or 5 mM EGTA plus 1 M thapsigargin (&). For experimental details see Ref. 48. heat production. The uncoupled efflux can also be measured diluting vesicles previously loaded with Ca2+ in a medium containing only buffer and EGTA. In the absence of either Mg2+, ADP or Pi , the ATPase cannot synthesize ATP (Fig. 7). For the muscle vesicles, the efflux measured in presence of excess EGTA decreases when thapsigargin, a specific inhibitor of the Ca2+-ATPase [63, 64], is added to the medium simultaneously with EGTA. The difference between the total efflux and the efflux measured in presence of thapsigargin represents the uncoupled efflux mediated by the Ca2+-ATPase [48, 65] and in muscle vesicles it represents about 70% of the total Ca2+ efflux (Table 5). The Ca2+ efflux of platelet vesicles is slower than that of muscle Fig. 7. Ca2+ efflux from platelet (A) and skeletal muscle (B) vesicles. The vesicles were preloaded with 45 Ca and diluted to a final concentration of 30 µg of protein兾ml into a medium containing 50 mM MOPS兾 Tris pH 7.0 and 0.1 mM EGTA either in the absence (●) or presence (䊊) of 1 µM thapsigargin. For experimental details see Ref. 48. Heat Production by the Ca2+-ATPase 125 Table 5. Ca2+ Efflux from Skeletal Muscle and Blood Platelets Vesicles Vesicles and PAF addition n Total efflux (A) nmol兾mg · min−1 5 µM TG (B) nmol兾mg · min−1 TG-sensitive (A–B) Muscle Without PAF 4 µM PAF 7 4 203J26 228J38 63J19 97J20 140J22 130J38 Platelets Without PAF 4 µM PAF 6 4 40J3 H273J9 41J6 61J3 0 H212J10 The assay medium composition and experimental conditions were as described in Fig. 7. In the table, TG refers to thapsigargin and n to the number of experiments. Values are averageJSE. For experimental details, see Ref. 48. and is not impaired by thapsigargin, regardless of the method used to measure the efflux (Fig. 7 and Table 5). These data suggest that Ca2+ leaks through the SERCA 1 of skeletal muscle but not through the SERCA 2B and 3 found in blood platelets. Therefore, the difference of heat production measured in muscle and platelet vesicles after formation of a transmembrane gradient (Table 4) could be due to the absence of uncoupled Ca2+ leakage through the Ca2+-ATPase in platelet vesicles (thapsigargin sensitive efflux in Table 5). PLATELET ACTIVATING FACTOR Some of the lipids derived from the breakdown of membrane phospholipids are able to increase the uncoupled efflux mediated by the Ca2+-ATPase of skeletal muscle sarcoplasmic reticulum [66]. We therefore tested different lipids in platelet vesicles in search of a compound that could promote a thapsigargin sensitive Ca2+ efflux. The reasoning was that if we could promote the leakage of Ca2+ through the platelet Ca2+-ATPase then, similar to the muscle vesicles, the platelet vesicles should become able to convert osmotic energy into heat. In the course of these experiments we found that DL-α -phosphatidylcholine, β -acetyl-γ -O-hexadecyl could promote such an efflux in platelets but not in muscle vesicles. This phospholipid belongs to a family of acetylated phospholipids known as platelet activating factor (PAF) which are produced when cells involved in the inflammatory process are activated. PAF was found to inhibit the Ca2+ uptake of both platelet and muscle vesicles (Table 6). With the two vesicles, half maximal inhibition is obtained with 4 to 6 µM PAF. In contrast with the Ca2+ uptake, the ATPase activity of the two vesicle preparations was not inhibited by PAF (48). The discrepancy between Ca2+ uptake and ATPase activity suggests that the decrease of Ca2+ accumulation is promoted by an increase of Ca2+ efflux and not by an inhibition of the ATPase. The amount of Ca2+ retained by the vesicles is determined by the differences between the rates of Ca2+ uptake and of Ca2+ efflux. The higher the efflux, the smaller the amount of Ca2+ retained by the vesicles. The addition of PAF during the course of Ca2+ uptake promoted the release of Ca2+ until a new steady state level of Ca2+ retention was achieved (Fig. 8). With both preparations, the higher the concentration of PAF added, the lower the new steady state level of Ca2+ filling. The release of Ca2+ promoted by PAF was not 126 de Meis Table 6. Effect of PAF on the Ca2+ Uptake and ∆Hcal of ATP Hydrolysis Skeletal muscle vesicles Blood platelet vesicles Ca , µM PAF, µM Ca uptake µmol兾mg ∆H , kcal兾mol Pi Ca uptake µmol兾mg ∆Hcal kcal兾mol Pi 1 0 4 1.83J0.21 (4) 0.45J0.19 (4) −32.99J2.90 (4) −25.69J1.71 (4) 0.11J0.03 (3) 0.03J0.01 (3) −12.58J1.29 (5) −20.04J0.37 (3) 10 0 6 2.66J0.44 (3) 0.68J0.35 (5) −22.92J2.24 (3) −16.91J1.50 (5) 0.20J0.04 (3) 0.06J0.01 (3) −10.70J1.01 (3) −23.90J1.06 (3) 2+ 2+ cal 2+ The assay medium and experimental conditions were as in Table 4. The values in the table are the averageJSE of the number of experiments shown in parentheses. The differences between the ∆Hcal values measured in the absence and in the presence of PAF with skeletal muscle were significant (t-test) with pF0.05 both with 1 and 10 µM Ca2+ and with blood platelets were significant pF0.005 (1 µM Ca2+) and pF0.001 (10 µM Ca2+). For experimental details see Ref. 48. accompanied by a burst of ATP synthesis. On the contrary, PAF inhibited the synthesis of ATP driven by the coupled Ca2+ efflux [48]. This indicates that the Ca2+ release promoted by PAF was not promoted by an increase of the reversal of the pump. A major difference between the muscle and platelet vesicles was found when thapsigargin was added to the medium together with PAF. For the platelet vesicles, the rate of Ca2+ release measured after the addition of PAF was greatly decreased in presence of thapsigargin (Fig. 8 and Table 5) indicating that most of the Ca2+ left the vesicles through the ATPase as an uncoupled Ca2+ efflux. This could be better seen after the initial minute of incubation. In fact, the rate of release in platelet Fig. 8. Ca2+ release after the addition of PAF. The assay medium composition was 50 mM MOPS兾 Tris pH 7.0, 2 mM MgCl2 , 10 mM Pi , 40 µM CaCl2 , 100 mM KCl and 3 mM ATP. The reaction was started by the addition of either platelets (A) or muscle (B) vesicles at 35°C (䊊) control without additions. The arrow indicates the addition of either 6 µM PAF (䉭) or 6 µM PAF plus 1 M thapsigargin (&). For experimental details see Ref. 48. Heat Production by the Ca2+-ATPase 127 vesicles was so fast that we could not measure the initial velocity of release with the method available in our laboratory. Thus, the values with PAF in Table 5 differ from the other values in that it does not reflect a true rate, but only the parcel of Ca2+ released during the first incubation minute. In muscle, the rate of Ca2+ efflux measured after the addition of PAF was slower than that measured with platelet vesicles (compare Figs. 8A and B) and the proportion between the Ca2+ effluxes sensitive and insensitive to thapsigargin measured with PAF was practically the same as that measured when the pump was arrested with EGTA (Table 5). Having found a compound that induces the release of Ca2+ through the pump, we then measured the heat produced during ATP hydrolysis in the presence and absence of PAF (Table 6). The PAF concentrations selected were sufficient to enhance the rate of efflux without completely abolishing the retention of Ca2+ by the vesicles, i.e., without abolishing the formation of a Ca2+ gradient through the vesicles membrane. In such conditions PAF enhances the amount of heat produced during the hydrolysis of ATP by the blood platelets. In muscle vesicles however, PAF decreases the amount of heat produced during ATP hydrolysis. The ∆Hcal values measured with PAF and muscle vesicles were less negative than those measured in absence of PAF, but still more negative than the values measured in absence of Ca2+ gradient. CONVERSION OF OSMOTIC ENERGY INTO EITHER CHEMICAL ENERGY OR HEAT DURING THE THAPSIGARGIN SENSITIVE Ca2+ EFFLUX The experiments described above were performed in steady state conditions, during which Ca2+ is translocated both inward and outward the vesicles and the Ca2+-ATPase catalyze simultaneously the hydrolysis and the synthesis of ATP. Thus, the heat measured during steady state represents in fact the sum of these different reactions. We therefore decided to measure the caloric yield during the unidirectional efflux of Ca2+ [49]. In these experiments it was found that heat is absorbed from the medium if ATP is synthesized during the Ca2+ efflux. However, if the synthesis of ATP is impaired, then the Ca2+ efflux mediated the ATPase leads to the reduction of a significant amount of heat. The synthesis of ATP was determined diluting vesicles previously loaded with Ca2+ in a medium containing ADP, Pi , Mg2+ and EGTA. In this condition Ca2+ leaves the vesicles at a fast rate, ATP is synthesized from ADP and Pi and heat is absorbed from the environment (Fig. 9 and Table 7). The amount of heat absorbed from the medium was proportional to both the amounts of ATP synthesized and the amount of thapsigargin sensitive Ca2+ efflux (compare Figs. 9B and C). The ∆Hcal values of Table 7 shows that 5 kcal were absorbed for each mol of Ca2+ released from the vesicles. The rate of thapsigargin sensitive Ca2+ efflux decreased and ATP was no longer synthesized when Mg2+ was not added and EDTA was included in the medium in order to chelate the small amount of Mg2+ introduced in the medium together with the Ca2+ loaded vesicles. In this condition the synthesis of ATP was impaired and the Ca2+ efflux was exothermic. Now, the amount of heat released was proportional to the amount of Ca2+ released through the thapsigargin sensitive route and 30 kcal were produced for each mole of Ca2+ released by the vesicles (Fig. 9 and Table 7). 128 de Meis Fig. 9. Unidirectional Ca2+ efflux (A), ATP synthesis (B) and heat release (C). The assay medium composition was 50 mM MOPS兾Tris buffer (pH 7.0), 5 mM EGTA and (䊊) 0.1 mM ADP, 10 mM Pi , 4 mM MgCl2 , 100 mM KCl, 5 mM NaN3 , 10 µM P 1, P 5-di(adenosine 5′) pentaphosphate, 20 mM glucose and 10 units兾ml hexokinase; (●) same as in (䊊) but without MgCl2 and with 5 mM EDTA; (B) same as in (●) but with 1 µM thapsigargin. The reaction was started by the addition of Ca2+ loaded vesicles, total of 20 µg兾ml. The values shown in the figure are meanJSE of 6 experiments. Table 7. Energy Transduction During the Thapsigargin Sensitive Ca2+ Efflux Additions to efflux media ADP, Pi , Mg2+ ADP, Pi , EDTA n TG sensit. Ca2+ efflux µmol兾mg ATP synthesis µmol兾mg Heat, mcal兾mg Ca2+ efflux ∆Hcal, kcal兾 Ca2+ mol 11 14 0.77J0.09 0.11J0.03 0.62J0.06 None +3.86J1.01 −3.30J0.48 +5.01J1.05 −30.00J1.62 The values of Ca2+ effluxes are the difference between the rates measured with and without 1 µM thapsigargin (TG). Other additions to the assay medium were 50 mM MOPS兾Tris buffer pH 7.0, 100 mM KCl, 5 mM EGTA, 5 mM sodium azide, 20 mM glucose and 11 units hexokinase兾ml. The concentrations of ADP, Pi , MgCl2 and EDTA were 0.1 mM, 10 mM, 4 mM and 5 mM respectively. The reaction time at 35°C was 9 min. The values are meanJSE. In the table, n refers to the number of experiments. For experimental details see Ref. 49. THE CYCLE OF ENERGY INTERCONVERSION The catalytic cycle of the Ca2+-ATPase varies depending on whether or not a Ca gradient is formed across the vesicle membrane (Figs. 10 and 11). The basic sequence (no gradient) was first proposed in 1976 and since then has been widely confirmed in different laboratories using different experimental approaches [10, 11, 33, 67–69]. During catalysis the enzyme cycles through two different conformations, E1 and E2 (originally E and *E, Ref. 67). The enzyme form E1 binds Ca2+ with high affinity (Ks 1 µm at pH 7.0) on the outer surface of the vesicles (reaction 1 in Fig. 10) and can be phosphorylated by ATP forming an acyl phosphate residue (phosphoaspartate) at the catalytic site of the enzyme (reaction 2). In the form E1 the enzyme cannot be phosphorylated by Pi . The enzyme form E2 binds Ca2+ with low affinity (Ks 2 mM at pH 7.0) on the inner surface of the vesicles (reaction 4) and 2+ Heat Production by the Ca2+-ATPase 129 Fig. 10. The catalytic cycle of the Ca2+-ATPase in the absence of a transmembrane Ca2+ gradient. Fig. 11. The catalytic cycle of the Ca2+-ATPase after formation of a transmembrane Ca2+ gradient. can be phosphorylated by Pi but not by ATP (reaction 5 backwards). The key feature of this cycle is the mechanism by which energy is transduced. The sequence of events proposed in earlier models of active transport assumed that energy was released at the moment of hydrolysis of the phosphate compound. The energy would then be absorbed by the enzyme and used to perform work. In this view, the energy of hydrolysis of phosphate compounds would be the same regardless of whether the phosphate compound was in solution in the cytosol or bound to the enzyme surface [10, 11]. In aqueous solutions an acyl phosphate residue has a high energy of hydrolysis, the equilibrium constant for the hydrolysis (Keq ) being practically the same as that of ATP (Table 8). In 1974 it was found that the ATPase could be spontaneously phosphorylated by Pi forming a phosphoaspartate without the need 130 de Meis Table 8. Variability of the Energy of Hydrolysis of Phosphate Compounds During the Catalytic Cycle of Energy Transducing Enzymes Solution or enzyme bound before work Enzyme Reaction Ca2+-ATPase, Na+兾K +-ATPase Aspartyl phosphate hydrolysis F1-ATPase myosin Inorganic pyrophosphatase Hexokinase ATP hydrolysis Pi hydrolysis ATPCglucose→ Glucose 6-PCADP Enzyme bound after work Keq (M) ∆G0, (kcal兾mol) Keq (M) (kcal兾mol) ∆G0 106 −8.4 1.0 0 106 104 2B103 −8.4 −5.6 −4.6 1.0 4.5 1.0 0 −0.9 0 For details, see reviews 11 and 35. for energy input into the system (reaction 5 backwards). This was only observed in the absence of Ca2+ when the enzyme is stabilized in the E2 form [70, 71]. The low energy E2AP could be converted back into the high energy form 2Ca: E1 ∼ P (reactions 4 and 3 backwards) when the Ca2+ concentration in the medium was suddenly raised to the range of 1 to 3 mM, i.e., to a range similar to that found in the vesicle lumen after formation of transmembrane Ca2+ gradient (∼5 mM at pH 7.0). The 2Ca: E1 ∼ P form (but not the E2AP) can transfer its phosphate to ADP (reaction 2 backwards) forming soluble ATP [46, 53, 54, 72]. The finding that the energy of hydrolysis of the phosphoenzyme varies during the catalytic cycle indicates that the energy needed for the translocation of Ca2+ through the membrane is used before the cleavage of the phosphoenzyme (reaction 3), i.e., it is during the conversion of 2Ca: E1 ∼ P into 2Ca: E2AP that the chemical energy is used to perform work. Simultaneously with the discovery of the low energy phosphoenzyme E2 ∼ P [71] in Paul Boyer’s laboratory [73, 74] it was found that ATP could be spontaneously formed at the catalytic site of mitochondrial F1 ATPase and in the subsequent years it became apparent that the conversion of a phosphate compound from ‘‘high’’ into ‘‘low’’ energy is a general feature found in different enzymes involved in processes of energy transduction (Table 8) such as the Na+兾K +-ATPase, myosin, inorganic pyrophosphate and hexokinase [11, 35, 69]. After the description of the catalytic cycle of the Ca2+-ATPase it was found that the energy change of the phosphoenzyme (reaction 3) was related to a change in water activity in the microenvironment of the catalytic site [50]. The energy of hydrolysis of different phosphate compounds is determined by the differences in solvation energy between the reactant and products. Thus, a change of water activity in the catalytic site of the enzyme leads to a change in solvation energy of the phosphoenzyme and to a change in its energy of hydrolysis. Experimental evidence supporting this possibility was obtained in various laboratories and were described in reviews previously published [11, 35, 69]. As for the phosphoaspartate at the catalytic site of the Ca2+-ATPase, the conversion of the ‘‘low energy’’ tightly bound ATP into the ‘‘high energy’’ loosely bound ATP seems to be promoted by a change of water activity at the catalytic site of the mitochondrial F1 ATPase [11, 35, 74]. Heat Production by the Ca2+-ATPase 131 The sequence shown in Fig. 10 is only valid for leaky vesicles in presence of Ca2+ concentrations similar to those found in the sarcoplasm (0.1 to 10.0 µm). In these conditions there is no Ca2+ accumulation in the vesicles lumen and after translocation through the membrane, the calcium ions bound to the different E2 enzyme form readily dissociate from the enzyme (Fig. 1B). In intact vesicles however, Ca2+ is retained by the vesicles and after a few catalytic cycles its concentration in the vesicles lumen rises to about 5 mM. In these conditions calcium remains bound to the enzyme as it cycles between the two forms E1 and E2 and the catalytic cycle now includes different steps not detected when the vesicles are leaky (reactions 4 to 7). With intact vesicles, a steady state of Ca2+ uptake is reached after a short incubation interval during which the rate of Ca2+ efflux equals the rate of Ca2+ uptake, the ATPase catalyzes simultaneously the hydrolysis and the synthesis of ATP and depending on the conditions used, the amount of heat produced for each ATP molecule increases two- to threefold (Table 1). Part of the Ca2+ efflux is coupled to the synthesis of ATP (ATP↔Pi exchange) and in the sequence this is accounted for by reactions 2 to 5 flowing forward and backward. There is no ATP ↔Pi and Cain ↔Caout exchange mediated by the ATP with leaky vesicles incubated in the presence of physiological Ca2+ concentrations because the Ca2+ concentration on the vesicles lumen is the same as that of the assay medium and far too small to permit the rebinding of Ca2+ to the E2 form. Therefore, in the sequence shown in Fig. 10, reaction 4 is irreversible and this does not allow the reversal of reactions 3 and 2. For leaky vesicles the chemical energy of ATP hydrolysis is divided into at least two parcels. One is converted into heat and corresponds to the ∆Hcal measured with leaky vesicles in Table 2 and Fig. 1. The second parcel is converted into work, i.e., the translocation of calcium from the outer surface to the inner surface of the vesicles and is accounted for reactions 1 to 4 forward in Fig. 10. In intact vesicles, the parcel of energy used to translocate Ca2+ through the membrane is not consumed in work but it is conserved in the form of osmotic energy. In steady state the parcel of Ca2+ which enter the vesicles bound to the enzyme returns to the outer surface of the vesicles also bound to the enzyme either through the reversal of reaction 2 to 4 or through reactions 5 and 6 and in both cases there is no net Ca2+ accumulation. The high Ca2+ concentrations available in the vesicle lumen allow for the reversal of reactions 5 to 2 during which osmotic energy is converted back into the chemical energy needed for the synthesis of ATP detected during the ATP ↔Pi exchange reaction (Table 1). The uncoupled Ca2+ efflux is accounted by reactions 5, 6 and by the sequence shown in Fig. 12. If the Ca2+ concentration in the outer surface of the vesicles is not sufficient to saturate the calcium binding sites of the enzyme form E1 , then part of the enzyme form 2Ca: E1 (reaction 1 Fig. 11) is converted into E1 which will then cycle through the membrane as shown in Fig. 12 promoting a net Ca2+ efflux during which, osmotic energy is converted into heat by the ATPase (Fig. 9 and Table 7). Another source of heat is probably derived from the hydrolysis of the enzyme form 2Ca: E ∼ P represented as reaction 7 in Fig. 11. Recently [75, 76] it has been shown that after formation of the Ca2+ gradient, the amount of ATP cleaved by the Ca2+-ATPase far exceeds the amount of ATP needed to pump back the Ca2+ that leaks from the vesicles. The kinetic evidence reported indicates the extra amount 132 de Meis Fig. 12. The uncoupled Ca2+ efflux. of ATP cleaved is derived from the hydrolysis of the enzyme form 2Ca: E1 ∼ P (reaction 7 in Fig. 11). The amount of heat produced during the hydrolysis of the 2Ca: E1 ∼ P enzyme should be higher than that of the enzyme form E2AP due to difference of the Keq for the hydrolysis of the two enzyme forms (Table 8) and because the cleavage of 2Ca: E1 ∼ P takes place before part of the energy of the system is converted into work or osmotic energy. Thus, both the Ca2+ efflux though the cycle of Fig. 12 and the hydrolysis of the enzyme form 2Ca: E1 ∼ P (reaction 7 Fig. 11) are probably responsible for the different values of ∆Hcal for ATP hydrolysis measured in presence and absence of a transmembrane Ca2+ gradient. ATP hydrolysis measured in presence and absence of a transmembrane Ca2+ gradient. Recently [77], it was shown that the amount of heat released during the cleavage of the phosphoenzyme form 2Ca: E1 ∼ P is much larger than that measured during the hydrolysis of the form E2AP and the difference of heat production noted during ATP hydrolysis in the presence and absence of a Ca2+ gradient is mostly related to the phosphoenzyme form preferentially cleaved during catalysis, 2Ca: E1 ∼ P in presence of a transmembrane Ca2+ gradient and E2AP in leaky vesicles. CONCLUSIONS The Ca2+-ATPase can regulate the interconversion of energy in such a way as to vary the fraction of the energy derived from ATP hydrolysis which is dissipated as heat. This can be observed both in the presence and in the absence of a transmembrane Ca2+ gradient and depending on the conditions used the ∆Hcal for ATP hydrolysis may vary from −7.8 (Table 2) up to −31.9 kcal兾mol Pi (Table 4). The experiments described suggest the following sequences of energy conversion: (i) Ca2+ gradient—from the total chemical energy released during the ATP hydrolysis (∼30 kcal兾mol Pi ), about 31 is converted into heat (∼10 kcal) provided that the Ca2+ concentration on both sides of the membrane is kept in the micromolar range. This was observed with the use of leaky vesicles in Tables 1 and 2. The rest of the energy ( ∼20 kcal兾mol Pi ) is probably Heat Production by the Ca2+-ATPase 133 used to translocate Ca2+ across the membrane (work). However, if the membrane is intact, then the energy used for the translocation of Ca2+ is converted into osmotic energy (Ca2+ gradient) and the Ca2+-ATPase can use this energy to either synthesize back a small part of the ATP previously cleaved or to produce heat. The balance between these two routes would be determined by the ratio between the coupled and uncoupled enzyme units. In one extreme (dimethylsulfoxide in Table 1), there would be a high degree of energy conservation, most of the energy derived from the hydrolysis of ATP being conserved by the vesicles as osmotic energy and practically all the Ca2+ that leaves the vesicles is used to synthesize back a part of the ATP previously cleaved. In the other extreme (3 µg兾ml heparin), the SR operates as if it was a ‘‘furnace,’’ a small amount of Ca2+ is retained by the vesicles and most of the energy derived from ATP hydrolysis is dissipated into the medium as heat. (ii) No gradient—previous studies demonstrated that both leaky vesicles and the soluble Ca2+-ATPase are able to retain part of the energy derived from ATP hydrolysis even after both ADP and Pi dissociate from the enzyme and the energy retained can be used by the enzyme for the synthesis of a new ATP molecule from ADP and Pi . This is promoted by the binding of Ca2+ to a low affinity site of the Ca2+-ATPase located in a region of the protein facing the vesicles lumen [4, 25, 26, 30–32]. The data of Table 2 indicates that the synthesis of ATP in leaky vesicles is associated with an increase of the ∆Hcal value for ATP hydrolysis, suggesting that the binding of Ca2+ to the low affinity site of the enzyme may regulate the fraction of the energy that dissipates as heat and that which can be used for the synthesis of ATP. (iii) It seems that osmotic energy cannot be transformed spontaneously into heat and that a device is needed for this conversion. For the sarcoplasmic reticulum the device is probably the Ca2+-ATPase itself, that in addition to interconverting chemical into osmotic energy, can also convert osmotic energy into heat (Table 1). (iv) It is generally assumed that the energy released during the hydrolysis of ATP by the Ca2+-ATPase can be divided in two non-interchangeable parts, one is converted into heat and the other is used to pump Ca2+ across the membrane. This was observed with the platelet vesicles before the addition of PAF (Table 6). The finding that the SERCA 1 can convert osmotic energy into heat revealed an alternative route that increases two- to threefold the amount of heat produced during ATP hydrolysis therefore permitting the maintenance of the cell temperature with a smaller consumption of ATP. The data obtained with the blood platelet vesicles show that not all the SERCA isoforms are able to readily convert osmotic energy into heat. These vesicles however, can be converted by PAF into a system capable of increasing the heat production during ATP hydrolysis (Tables 5 and 6), suggesting that the mechanism capable of providing additional heat production can be turned on and off and this could represent a mechanism of thermoregulation specific of the cells expressing SERCA 2b and 3. Both 134 de Meis in muscle and platelet vesicles there is a Ca2+ efflux which is not inhibited by thapsigargin. We do not know through which membrane structure this Ca2+ flows, but the data obtained with platelets before the addition of PAF indicate that during this efflux, osmotic energy is not converted into heat. In platelets, PAF promoted simultaneously the appearance of thapsigargin sensitive efflux and extra-heat production during ATP hydrolysis. These observations corroborate with the notion that the conversion of osmotic energy into heat cannot be promoted by any kind of Ca2+ leakage and that a device is needed for this conversion. ACKNOWLEDGMENT This work was supported by grants from PRONEX–Financiadora de Estudos e Projetos (FINEP), Conselho Nacional de Desenvolvimento Cientı́fico e Tecnológico (CNPq) and by Fundação de Amparo à Pesquisa do Estado do Rio de Janeiro (FAPERJ). REFERENCES 1. Nicholls, D. and Locke, R. M. (1984) Thermogenic mechanisms in brown fat. Physiol. Reû. 64:1– 64. 2. Clausen, T., van Hardeveld, C., and Everts, M. E. (1991) Significance of cation transport in control of energy metabolism and thermogenesis. Physiol. Reû. 71:733–774. 3. Block, B. A. (1994) Thermogenesis in muscle. Annu. Reû. Physiol. 56:535–577. 4. Janský, L. (1995) Humoral thermogenesis and its role in maintaining energy balance. Physiol. Reû. 75:237–259. 5. Skulachev, V. P. (1998) Uncoupling: new approaches to an old problem of bioenergetics. Biochem. Biophys. Acta 1363:100–124. 6. Boss, O., Muzzin, P., and Giacobino, J. P. (1987) The uncoupling proteins, a review. Eur. J. Endocrinol 139:1–9. 7. Lowell, B. B. and Spiegelman, B. M. (2000) Toward a molecular understanding of adaptive thermogenesis. Nature 404:652–660. 8. Chinet, A. E., Decrouy, A., and Even, P. C. (1992) Ca2+ dependent heat production under basal and near-basal conditions in the mouse soleus muscle. J. Physiol. Lond. 455:663–678. 9. Hasselbach, W. (1978) Reversibility of the sarcoplasmic calcium pump. Biochim. Biophys. Acta 515:23–53. 10. de Meis, L. (1981) The sarcoplasmic reticulum: Transport and energy transduction, Vol. 2, L. Bittar (ed.), Wiley, New York. 11. de Meis, L. (1989) Role of water in the energy of hydrolysis of phosphate compounds—Energy transduction in biological membranes. Biochim. Biophys. Acta 973:333–349. 12. Bianco, A. C. and Silva, J. E. (1987) Optimal response of key enzymes and uncoupling protein to cold BAT depends on local T3 generation. Am. J. Physiol. 253:E-255–E263. 13. Klingenspor, M., Ivemeyer, M., Wiesinger, H., Haas, K., Heldmaier, G., and Wiesner, J. (1996) Biogenesis of thermogenic mitochondria in brown adipose tissue of Djugarian hamsters during cold adaptation. Biochem. J. 316:607–613. 14. Boss, O. et al. (1998) Uncoupling protein-3 expression in rodent skeletal muscle is modulated by food intake but not by changes in environmental temperature. J. Biol. Chem. 273:5–8. 15. Dumonteil, E., Barré, H., and Meissner, G. (1993) Sarcoplasmic reticulum Ca2+-ATPase and rymanodine receptor in cold-acclimated ducklings and thermogenesis. Am. J. Physiol. 265 (Cell Physiol. 34): C507–C513. Heat Production by the Ca2+-ATPase 135 16. Dumonteil, E., Barré, H., and Meissner, G. (1995) Expression of sarcoplasmic reticulum Ca2+ transport proteins in cold-acclimating ducklings. Am. J. Physiol. 269 (Cell Physiol. 38):C955–C960. 17. Gong, D. W., Yufang, H., Karas, M., and Reitman, M. (1997) Uncoupling protein-3 is a mediator of thermogenesis regulated by thyroid hormone, β 3-adrenergic agonists, and leptin. J. Biol. Chem. 272:24129–24132. 18. Van der Linden, C. G. et al. (1996) Fiber-specific regulation of Ca2+-ATPase isoform expression by thyroid hormone in rat skeletal muscle. Am. J. Physiol. 271:C1908–C1919. 19. Simonides, W. S., Brent, G. A., Thelens, M. H. M., van der Linden, C. G., Larsen, P. R., and van Hardeveld, C. (1996) Characterization of the promoter of the rat sarcoplasmic endoplasmic reticulum Ca2+-ATPase1 gene and analysis of thyroid hormone responsiveness. J. Biol. Chem. 271:32048–32056. 20. Hämälainen, N. and Pette, D. (1997) Coordinated fast-to-slow transitions of myosin and SERCA isoforms in chronically stimulated muscles of euthyroid and hyperthyroid rabbits. J. Muscle Res. Cell Motil. 18:545–554. 21. Ethan, A. H. and Danforth, Jr., E. (1987) Expenditure and storage of energy in man. J. Clin. Inûest. 79:1019–1025. 22. Bouchard, C. et al. (1990) The response of long-term overfeeding in identical twins. New Engl. J. Med. 322:1477–1482. 23. Diaz, E. O., Prentice, A. M., Goldberg, G. R., Murgatroyd, P. R., and Coward, W. A. (1992) Metabolic response to experimental overfeeding in lean and overweight healthy volunteers. Am. J. Clin. Nutr. 56:641–655. 24. Levine, J. A., Eberhardt, N. L., and Jensen, M. D. (1999) Role of nonexercise activity thermogenesis in resistance to fat gain in humans. Science 283:212–214. 25. Busto, R., Dietrich, W. D., Globus, M., Valdés, I., Scheinberg, P., and Ginsberg, M. (1987). Small differences in intraschemic brain temperature critically determine the extent of ischemic neuronal injury. J. Cereb. Blood Flow Metab. 7:129–138. 26. Dietrich, D. L. L. and Elzinga, G. (1993) Heat produced by rabbit papillary muscle during anoxia and reoxygenation. Circ. Res. 73:1177–1187. 27. Caputa, M., Folkow, L., and Blix, A. S. (1998) Rapid brain cooling in diving ducks. Am. J. Physiol. 275:R363–R371. 28. Gordon, C. (1988) Temperature regulation in laboratory mammals following acute toxic insult. Toxicology 53:161–178. 29. Wood, S. C. (1991) Interaction between hypoxia and hypothermia. Annu. Reû. Physiol. 53:71–85. 30. Hasselbach, W. and Makinose, M. (1961) Die Calcium pumpe der ‘‘Erschlaffungsgrana’’ des Muskels und ihre Abhängigkeit vor der ATP-spaltung. Biochem. Z. 333:518–528. 31. Barlogie, B., Hasselbach, W., and Makinose, M. (1971) Activation of calcium efflux by ADP and inorganic phosphate. FEBS Lett. 12:267–268. 32. Makinose, M. and Hasselbach, W. (1971) ATP synthesis by the reverse of sarcoplasmic reticulum pump. FEBS Lett. 12:271–272. 33. de Meis, L. and Vianna, A. L. (1979) Energy interconversion by the Ca2+-transport ATPase of Sarcoplasmic Reticulum. Annu. Reû. Biochem. 48:275–292. 34. Inesi, G. (1985) Mechanism of Ca2+ transport. Annu. Reû. Physiol. 47:573–601. 35. de Meis, L. (1993) The concept of energy-rich phosphate compounds: water, transport ATPases and entropic energy. Arch. Biochem. Biophys. 306:287–296. 36. Gould, G. W., McWhirter, J. M., East, J. M., and Lee, A. G. (1987) A fast passive Ca2+ efflux mediated by the (Ca2+CMg2+)-ATPase in reconstituted vesicles. Biochem. Biophys. Acta 904:45–54. 37. Gould, G. W., McWhirter, J. M., and Lee, A. G. (1978) A fast passive Ca2+ efflux mediated by the (Ca2+CMg2+)-ATPase in reconstituted vesicles. Biochim. Biophys. Acta 904:45–54. 38. Inesi, G. and de Meis, L. (1989) Regulation of steady state filling in sarcoplasmic reticulum. Roles of back-inhibition, leakage, and slippage of the calcium pump. J. Biol. Chem. 264:5929–5936. 39. Galina, A. and de Meis, L. (1991) Ca2+ translocation and catalytic activity of the sarcoplasmic reticulum ATPase; Modulation by ATP, Ca2+ and Pi . J. Biol. Chem. 266:17978–17982. 40. de Meis, L., Suzano, V. A., and Inesi, G. (1990) Functional interactions of catalytic site and transmembrane channel in the sarcoplasmic reticulum ATPase. J. Biol. Chem. 265:18848–18851. 41. de Meis, L. (1991) Fast efflux of Ca2+ mediated by the sarcoplasmic reticulum Ca2+-ATPase. J. Biol. Chem. 266:5736–5742. 136 de Meis 42. de Meis, L. and Inesi, G. (1992) Functional evidence of a transmembrane channel with the Ca2+ transport ATPase of sarcoplasmic reticulum. FEBS Lett. 299:33–35. 43. de Meis, L. and Suzano, V. A. (1994) Uncoupling of muscle and blood platelets Ca2+ transport ATPase by heparin: regulation by K +. J. Biol. Chem. 269:14525–14529. 44. Wolosker, H. and de Meis, L. (1995) Ligand-gated channel of the sarcoplasmic reticulum Ca2+ transport ATPase. Biosci. Rep. 15:365–376. 45. de Meis, L., Bianconi, M. L., and Suzano, V. A. (1997) Control of energy fluxes by the sarcoplasmic reticulum Ca2+-ATPase: ATP hydrolysis, ATP synthesis and heat production. FEBS Lett. 406:201– 204. 46. de Meis, L. (1998) Control of heat produced during ATP hydrolysis by the sarcoplasmic reticulum Ca2+-ATPase in the absence of a Ca2+ gradient. Biochem. Biophys. Res. Commun. 243:598–600. 47. de Meis, L. (1998) Control of heat production by the Ca2+-ATPase of rabbit and trout sarcoplasmic reticulum. Am. J. Physiol. 274 (Cell Physiol. 43):C1738–C1744. 48. Mitidieri, F. and de Meis, L. (1999) Ca2+ release and heat production by the endoplasmic reticulum Ca2+-ATPase of blood platelets: effect of the platelets activating factor. J. Biol. Chem. 274:28344– 28350. 49. de Meis, L. (2000) ATP synthesis and heat production during Ca2+ efflux by sarcoplasmic reticulum Ca2+-ATPase. Biochem. Biophys. Res. Commun. 276:35–39. 50. de Meis, L., Martins, O. B., and Alves, E. W. (1980) Role of water, hydrogen ions, and temperature on the synthesis of adenosine triphosphate by the sarcoplasmic reticulum adenosine tryphosphatase in the absence of a calcium ion gradient. Biochem. 19:4252–4261. 51. Rocha, J. B., Wolosker, H., Souza, D. O., and de Meis, L. (1996) Alteration of Ca2+ fluxes in brain microsomes by K + and Na+ : Modulation by sulfated polysaccharides and trifluoperazine. J. Neurochem. 55:772–778. 52. de Meis, L. (1998) Approaches to study mechanisms of ATP synthesis in sarcoplasmic reticulum. Methods Enzymol. 157:190–206. 53. de Meis, L. and Carvalho, M. G. (1974) Role of the Ca2+ concentration gradient in the adenosine 5′triphosphate. Inorganic phosphate exchange catalyzed by sarcoplasmic reticulum. Biochem. 13:5032–5038. 54. de Meis, L. and Sorenson, M. M. (1975) ATP–Pi exchange and membrane phosphorylation in sarcoplasmic reticulum vesicles: Activation by silver in the absence of a Ca2+ concentration gradient. Biochem. 14:2739–2744. 55. de Meis, L. and Inesi, G. (1985) Intrinsic regulation of substrate fluxes and energy conservation in Ca2+-ATPase. FEBS Lett. 185:135–138. 56. Chini, E. N., Toledo, F. G. S., Albuquerque, M. C., and de Meis, L. (1993) The Ca2+-transporting ATPases of rabbit and trout exhibit different pH- and temperature dependence. Biochem. J. 293:469– 473. 57. MacLennan, D. H., Brandl, C. J., Korczak, B., and Green, N. M. (1985) Amino acid sequence of Ca2+, Mg2+-dependent ATPase from rabbit muscle sarcoplasmic reticulum, deduced from its complementary DNA sequence. Nature 316:696–700. 58. Lytton, J., MacLennan, D. H. (1988) Molecular cloning of cDNAs from human kidney coding for two alternatively spliced products of the cardiac Ca2+-ATPase gene. J. Biol. Chem. 263:15024–15031. 59. Wuytack, F. et al. (1994) A sarco兾endoplasmic reticulum Ca2+-ATPase3-type Ca2+ pump is expressed in platelets, in lymphyoid cells, and in mast cells. J. Biol. Chem. 269:1410–1416. 60. Lytton, J., Westin, M., Burk, S. E., Shull, G. E., MacLennan, D. H. (1992) Functional comparisons between isoforms of the sarcoplasmic reticulum family of calcium pumps. J. Biol. Chem. 267:14483– 14489. 61. Gajewski, E., Steckler, D. K., and Goldberg, R. N. (1986) Thermodynamics of the hydrolysis of adenosine 5′triphosphate to adenosine 5′-diphosphate. J. Biol. Chem. 261:12733–12737. 62. de Meis, L., Hasselbach, W., and Machado, R. D. (1974) Characterization of calcium oxalate and calcium phosphate deposits in sarcoplasmic reticulum vesicles. J. Cell. Biol. 62:505–509. 63. Thastrup, O., Foder, B., and Scharff, O. (1987) The calcium mobilizing and tumor promoting agent, thapsigargin elevates the platelet cytoplasmic free calcium concentration to a higher steady state level. A possible mechanism of action for the tumor promotion. Biochem. Biophys. Res. Comm. 142:654– 660. Heat Production by the Ca2+-ATPase 137 64. Sagara, Y., Fernandez-Belda, F., de Meis, L., and Inesi, G. (1992) Characterization of the inhibition of intracellular Ca2+ transport ATPase by thapsigargin. J. Biol. Chem. 267:12606–12613. 65. Wolosker, H., and de Meis, L. (1994) pH dependent inhibitory effects of Ca2+, Mg2+ and K + on the Ca2+ efflux mediated by the sarcoplasmic reticulum ATPase. Am. J. Physiol. 266 (Cell Physiol. 35): C1376–C1381. 66. Cardoso, C. M. and de Meis, L. (1993) Modulation by fatty acids of Ca2+ fluxes in sarcoplasmic reticulum vesicles. Biochem. J. 296:49–52. 67. Carvalho, M. G., Souza, D. G., and de Meis, L. (1976) On a possible mechanism of energy conservation in sarcoplasmic reticulum membrane. J. Biol. Chem. 251:3629–3636. 68. Tanford, C. (1984) Twenty questions concerning the reaction cycle of the sarcoplasmic reticulum calcium pump. CRC Crit. Reû. Biochem. 17:123–151. 69. Wolosker, H., Engelender, S., and de Meis, L. (1998) Reaction mechanism of the sarcoplasmic reticulum Ca2+-ATPase. Adûances in Molecular and Cell Biology 23A:1–31. 70. Masuda, H. and de Meis, L. (1973) Phosphorylation of the sarcoplasmic reticulum membrane by orthophosphate. Inhibition by calcium ions. Biochemistry 12:4581–4585. 71. de Meis, L. and Masuda, H. (1974) Phosphorylation of the sarcoplasmic reticulum membrane by orthophosphate through two different reactions. Biochem. 13:2057–2061. 72. de Meis, L. and Tume, R. K. (1977) A new mechanism by which Ca2+ and H + concentration gradient drives the synthesis of ATP. pH jump and ATP synthesis by the Ca2+-dependent ATPase of the sarcoplasmic reticulum. Biochem. 16:4455–4463. 73. Boyer, P. D., Cross, R. L., and Momsen, W. (1973) A new concept for energy coupling in oxidative phosphorylation based on molecular explanation of the oxygen exchange reaction. Proc. Nat. Acad. Sci. USA 70:2837–2839. 74. Boyer, P. D. (1998) Nobel Lecture 1997—Energy, life and ATP. Biosci. Rep. 18:97–117. 75. Yu, X. and Inesi, G. (1995) Variable stoichiometric efficiency of Ca2+ and Sr 2+ transport by the sarcoplasmic reticulum ATPase. J. Biol. Chem. 270:4361–4367. 76. Fortea, M. I., Soler, F., and Fernandez-Belda, F. (2000) Insight into the uncoupling mechanism of sarcoplasmic reticulum ATPase using the phosphorylating substrate UTP. J. Biol. Chem. 275:12521– 12529. 77. de Meis, L. (2001) Uncoupled ATPase activity and heat production by the sarcoplasmic reticulum Ca2+-ATPase. J. Biol. Chem. 276 (in press).