Download Role of the Sarcoplasmic Reticulum Ca2+-ATPase

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).