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Medical Research Council, Dunn Human Nutrition Unit, Cambridge CB2 2XY, United Kingdom
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ABSTRACT |
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 TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES
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Cells isolated from the hepatopancreas of estivating snails (Helix aspersa) have strongly depressed mitochondrial respiration compared with controls. Mitochondrial respiration was divided into substrate oxidation (which produces the mitochondrial membrane potential) and ATP turnover and proton leak (which consume it). The activity of substrate oxidation (and probably ATP turnover) decreased, whereas the activity of proton leak remained constant in estivation. These primary changes resulted in a lower mitochondrial membrane potential in hepatopancreas cells from estivating compared with active snails, leading to secondary decreases in respiration to drive ATP turnover and proton leak. The respiration to drive ATP turnover and proton leak decreased in proportion to the overall decrease in mitochondrial respiration, so that the amount of ATP turned over per O2 consumed remained relatively constant and aerobic efficiency was maintained in this hypometabolic state. At least 75% of the total response of mitochondrial respiration to estivation was caused by primary changes in the kinetics of substrate oxidation, with only 25% or less of the response occurring through primary effects on ATP turnover.
hepatopancreas; substrate oxidation; proton leak; adenosine triphosphate turnover
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INTRODUCTION |
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TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES
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MANY ORGANISMS SURVIVE harsh environmental conditions, such as lack of oxygen, food, and water, by depressing their metabolic rate (24, 26, 27, 30). This decreases the rate at which they use the environmental supplies that are in shortage, thus extending their ability to survive. The land snail Helix aspersa undergoes metabolic depression (estivation) in response to desiccating conditions during the dry summer months. Viable cells have been isolated from the hepatopancreas of H. aspersa (25). Respiration rates of cells from estivating snails were 33% of those from active snails, when compared at their respective physiological pH and PO2 (25). Isolated hepatopancreas cells provide an excellent model in which to study metabolic depression, because they are more representative of the in vivo conditions than isolated mitochondria yet do not invoke problems, such as repeated homologous sampling and adequate perfusion, often encountered in isolated tissues.
Traditionally, understanding the mechanisms of metabolic depression involves studying a particular enzyme or process. However, if metabolic depression is to occur homeostatically with unchanged concentrations of intermediates such as ATP, then both producers and consumers of such intermediates must be coordinately downregulated. As a result, metabolic depression is unlikely to be caused by changes in a single enzyme or process. Moreover, the study of a single process does not allow the relative importance of producers and consumers of intermediates, such as ATP, to metabolic depression to be assessed. Compiling published results may help in identifying a greater range of metabolically depressed enzymes, but it may be hard to assemble results from different model systems that are subjected to different environmental stresses.
We are therefore trying to obtain an overview of the range of enzymes or processes that play a role in metabolic depression in one model system: hepatopancreas cells isolated from active and estivating H. aspersa. This can be achieved by dividing the whole of cellular metabolism into modules and then measuring the kinetics of the modules to see where the primary and secondary changes lie during estivation. Bishop and Brand (3) divided oxidative metabolism into nonmitochondrial and mitochondrial respiration based on their sensitivity to specific inhibitors of the mitochondrial electron transport chain. Proportional decreases in mitochondrial and nonmitochondrial respiration were found to account for the metabolic depression seen during estivation. The drop in mitochondrial respiration was mainly due to changes intrinsic to the cell, with pH playing only a minor role and PO2 having no effect (3).
The aim of the present study was to understand the primary and
secondary causes of decreased mitochondrial respiration in hepatopancreas cells from estivating H. aspersa. This was
achieved by further dividing mitochondrial respiration into the
producers (substrate oxidation) and consumers (ATP turnover and proton
leak) of the mitochondrial membrane potential (
m; see
Fig. 1). We show that respiration rate
decreases as a result of decreased substrate oxidation activity. The
resulting drop in m causes proton leak rate to
decrease without a change in its activity, and ATP turnover to
decrease, with a reduction in the activity of ATP turnover possibly
playing a role. At least 75% of the total response of mitochondrial
respiration to estivation occurs through primary changes in the
kinetics of substrate oxidation, with the remaining 25% or less
occurring through changes in the kinetics of ATP turnover.
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MATERIALS AND METHODS |
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TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES
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Animals. Garden snails (H. aspersa), fresh mass ~7 g, from Blades Biological Systems (Kent, UK), were washed and given water, lettuce, and carrot mix (containing carrots, bran, milk powder, and calcium carbonate) three times each week (25). Snails were kept in plastic tanks in a Sanyo Versatile Environmental Test Chamber MLR-350HT (Sanyo, Loughborough, UK), maintained at 25°C, 90% relative humidity, under 120-W fluorescent light on a 14:10-h light-dark cycle starting at 9:00 A.M. After 2 wk, one-half of the snails were maintained as before (controls) and one-half of were kept without food or water and at 30% relative humidity. After a further 16 days, these snails were fully estivating (8, 25) and were used in this state for up to 2 mo.
Isolation of hepatopancreas cells. Cells were isolated at 20-25°C from active or estivating H. aspersa as described by Guppy et al. (25), using mechanical disruption and collagenase digestion of the hepatopancreas followed by differential centrifugation in Sorvall SS-34 and MSE bench centrifuges. They were resuspended in incubation medium (10 mM HEPES, 90 mM NaCl, 5 mM KCl, 5 mM NaH2PO4, 2 mM MgCl2, 1 mM CaCl2, 5 mM glucose, 1 mM acetate, 10 mg bovine serum albumin/ml, 20 µg gentamicin/ml at pH 7.8 for active or 7.3 for estivating snails) to 4 × 106 cells/ml and used within 6 h.
Calculation of the electrical potential across the mitochondrial
inner membrane.
The electrical potential across the mitochondrial inner membrane
(m) was calculated using Eq. 1
(46), which describes the relationship between the
accumulation of triphenylmethylphosphonium (TPMP+) into the
cells and the m at 25°C. To determine
m, the total accumulation of TPMP+ into
the cells
([3H-TPMP+]tot/[3H-TPMP+]e)
had to be corrected for other factors that affect its accumulation. These factors are the mitochondrial and nonmitochondrial volumes of the
cell (vm and vc), the plasma membrane potential
(p), and the apparent TPMP+ activity
coefficients or TPMP+ binding corrections: mitochondrial
(am), nonmitochondrial (ac), and extracellular
(ae)
![&Dgr;&psgr;<SUB>m</SUB><IT>=</IT>59 log <FENCE><FR><NU>v<SUB>m</SUB>a<SUB>c</SUB></NU><DE>v<SUB>c</SUB>a<SUB>m</SUB></DE></FR><FENCE><FR><NU>[Cl<SUP><IT>−</IT></SUP>]<SUB>tot</SUB>[TPMP<SUP><IT>+</IT></SUP>]<SUB>tot</SUB>a<SUB>c</SUB>(v<SUB>c</SUB><IT>+</IT>v<SUB>m</SUB>)</NU><DE>[Cl<SUP><IT>−</IT></SUP>]<SUB>e</SUB>[TPMP<SUP><IT>+</IT></SUP>]<SUB>e</SUB>a<SUB>e</SUB>v<SUB>c</SUB></DE></FR><IT>−</IT>1</FENCE></FENCE>](/content/vol282/issue2/fulltext/R372/img001.gif)
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(1) |
Time course of TPMP+ equilibration.
Cell suspensions were incubated in 15-ml glass vials (1 ml of
cells/vial) at 25°C in a shaking water bath (130 cycles/min). 3H-labeled TPMP+ (0.2 µCi/ml) and 1 µM
carrier TPMP+, 0.1 µCi/ml [14C]polyethylene
glycol, and 0.1 mg/ml carrier polyethylene glycol were added to each
vial at t = 0 min; then samples were removed every 50 min and
[3H-TPMP+]tot/[3H-TPMP+]e
was measured as described in Brand (5). Polyethylene
glycol is a cell-impermeant marker used to correct for contamination of
the pellet by supernatant. At t = 150 min,
m was collapsed by adding 80 µM carbonyl cyanide
p-trifluoromethoxyphenylhydrazone (FCCP), oligomycin at 2 µg/106 cells, and 2.5 µM myxothiazol, all dissolved in
DMSO, to one set of vials; an equivalent volume of DMSO was added to a
parallel set of vials. TPMP+ accumulation into the cells
([3H-TPMP+]tot/[3H-TPMP+]e)
was measured every 50 min.
[3H-TPMP+]tot/[3H-TPMP+]e
was also calculated as TPMP+ space (notional volume of the
cells determined by the distribution of
3H-TPMP+, equivalent to cell volume multiplied
by TPMP+ accumulation ratio).
Cell volume. Cell volume was measured according to Brand (5). 3H2O (1 µCi/ml) and 0.1 µCi/ml [14C]polyethylene glycol and 0.1 mg/ml carrier polyethylene glycol were added to cell suspensions. Controls showed that cell volume did not change during the course of the incubation (250 min) and that polyethylene glycol was not metabolized by, and did not bind to, the cells over time (which would result in an underestimated cell volume). Mitochondrial volumes (vm) were calculated using the cell volume and the mitochondrial-to-cellular volume ratio from T. Bishop, A. Ocloo, and M. D. Brand (unpublished observations), and nonmitochondrial volumes (vc) were calculated by subtraction.
Plasma membrane potential.
Plasma membrane potential (p) was measured using
36Cl
distribution across the plasma membrane
(5). 36Cl (0.1 µCi/ml) and 1 µCi/ml 3H20 were added to cell suspensions in
parallel to cell volume measurements. It was assumed that snail
hepatopancreas cells have chloride channels in their plasma membranes;
they have been reported in rodent hepatocytes (37, 53).
Plasma membrane potentials were also measured in the presence of 0.5 µM valinomycin, which allows K+ diffusion across the
plasma membrane, thereby hyperpolarizing the plasma membrane potential.
A corresponding change in chloride distribution across the plasma
membrane was seen, showing that chloride distribution did act as an
indicator of the plasma membrane potential. Plasma membrane potentials
in isolated rat hepatocytes were unaffected by the addition of
myxothiazol or oligomycin (37) and snail hepatopancreas
cells were assumed to be similar.
TPMP+ binding corrections (am,
ac, and ae).
TPMP+ binding corrections or activity coefficients describe
the amount of TPMP+ that is free (not bound to proteins or
other molecules) as a fraction of the total amount of
TPMP+. The liver mitochondrial TPMP+ binding
correction (am) does not vary greatly between species (10, 40). The value of 0.44 for rat hepatocytes
(37) was, therefore, used for snail hepatopancreas cells.
The calculated mitochondrial membrane potential is very insensitive to
errors in am: even a twofold error only alters the value by
~18 mV. In addition, the absolute difference between
m in active and estivating snails will be unaffected
by the value of am chosen. The nonmitochondrial TPMP+ binding correction (ac) was calculated as
the ratio of the theoretical and observed TPMP+ spaces in
the absence of m. The extracellular TPMP+
binding correction (ae) was calculated from Nobes et al.
(37).
Kinetics of substrate oxidation, proton leak, and ATP turnover.
Cells (1 ml/15-ml glass vial) were incubated in a shaking water bath
(130 cycles/min) at 25°C in parallel for m and
respiration rate measurements with 0.2 µCi/ml
3H-TPMP+ (for m) or an
equivalent volume of H20 (for respiration), plus 1 µM
carrier TPMP+ as well as various inhibitors:
m
was changed by addition of the uncoupler FCCP (0, 3.5, and 7 µM). To determine the kinetics of proton conductance, m was
changed by addition of myxothiazol (an inhibitor of complex III and
therefore of substrate oxidation; 0, 0.25, 0.5, and 2.5 µM) in the
presence of oligomycin (an inhibitor of
F1FO-ATP synthase and, therefore, of ATP
turnover) at 2 µg/106 cells. To determine the kinetics of
the m consumers, m was changed by
addition of myxothiazol (0 and 2.5 µM). FCCP, myxothiazol, and
oligomycin were dissolved in DMSO; a total of 3 µl DMSO was added to
each vial.
After incubation for 250 min, m and respiration rates
were measured simultaneously. m was measured
(5) using the correction factors in Table
1. Respiration rates were measured using
two 0.5-ml Clark oxygen electrodes (Rank Brothers, Bottisham,
Cambridge, UK) thermostatted at 25°C and connected to a Kipp and
Zonen dual-channel chart recorder, assuming 479 nmol O/ml at air
saturation (41). Rates were expressed as a function of
cell number, determined by an average of 10 counts (each of 50 to 100 cells) in 0.1% (wt/vol) neutral red in an improved Neubauer
hemocytometer. Respiration rates were measured at 80% air saturation,
at which point 5 µM myxothiazol was added and the
myxothiazol-insensitive nonmitochondrial respiration rate was
subtracted to give the mitochondrial respiration rate. Mitochondrial
respiration does not vary with oxygen tension (3).
Respiration rates were therefore measured at high oxygen tensions
instead of the physiological oxygen tensions of cells from active and
estivating snails (42 and 29% air saturation, respectively). This
minimized the incubation time of cells within the electrode at the cost
of an increase in the ratio of nonmitochondrial to mitochondrial
respiration.
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m consumer
F1Fo-ATP synthase, and, therefore, typically
causes m to increase in mammalian hepatocytes (40). In snail hepatopancreas cells, however, oligomycin
caused m to decrease (possibly because substrate
oxidation is inhibited by the drop in ATP levels on addition of
oligomycin), so it was necessary to extrapolate the proton leak
kinetics to the resting m to calculate the cellular
proton leak rates. The best exponential and linear fits were applied to
the active and estivating proton leak data, taken together, to allow
estimation of the resting proton leak rate.
The kinetics of the ATP consumers were calculated by subtracting the
respiration rates driving proton leak from the respiration rates
driving the m consumers (for which a straight line
fit was taken to the 2 points) at the same m. This
was carried out using both the best exponential and straight line fits
for proton leak. ATP turnover rates were not calculated below the
m at which the apparent rate of the
m consumers was less than the proton leak rate. This
effect probably occurred because, at low m, ATP
hydrolysis maintained m (36) in the
absence of oligomycin.
Relative contributions of proton leak and ATP turnover to resting
cellular respiration.
The rates of respiration driving proton leak and ATP turnover
(calculated as mitochondrial respiration minus respiration rate driving
proton leak) are shown in Fig. 5. Respiration to drive cellular proton
leak was calculated in three different ways. In Fig. 5, A
and B, oligomycin-inhibited respiration was used. In Fig. 5,
C and D, proton leak rate was obtained from the
exponential fit to the proton leak data. In Fig. 5, E and
F, it was obtained from the straight line fit to the proton
leak data. In the first case, proton leak was underestimated, due to
the drop in m to 89 ± 3% of the resting
m in cells from active snails and to 85 ± 9%
of the resting m in cells from estivating snails on addition of oligomycin. In the last two cases, proton leak rate was
obtained at the resting m but required an assumption
about the shape of the proton conductance curve. As all three cases were flawed in different ways, all three were considered to obtain an
overview of the relative contributions of proton leak and ATP turnover
to resting cellular respiration in hepatopancreas cells from active and
estivating snails.
Statistics.
The depression of substrate oxidation rate during estivation and of
m on addition of oligomycin were calculated by
dividing the average depressed value by the average control value. The contributions of proton leak and ATP turnover to total mitochondrial respiration were calculated by dividing the average respiration driving
proton leak or ATP turnover by the average mitochondrial respiration.
SE values for the proton leak rate at a defined potential were
estimated from the errors at all experimental points.
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RESULTS |
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TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES
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Time course of TPMP+ equilibration.
m measurements required TPMP+ to be fully
equilibrated across both the plasma membrane and the mitochondrial
inner membrane, so time courses for equilibration of TPMP+
into hepatopancreas cells from active and estivating snails were measured. The accumulation of TPMP+ into the cells leveled
off after 250 min for cells from both active (Fig.
2A) and estivating snails
(Fig. 2B). Cells were, therefore, incubated with
TPMP+ for 250 min in subsequent experiments.
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m was abolished using FCCP, oligomycin, and
myxothiazol to reverse the mitochondrial accumulation of
TPMP+ (Fig. 2, A and B).
Mitochondrial density in snail hepatopancreas cells was low (Table 1),
so mitochondrial TPMP+ represented only a small fraction of
the total (Fig. 2, A and B), and only a small
fraction of the total TPMP+ was released on mitochondrial
depolarization. The remainder equilibrated into the cells according to
the plasma membrane potential and nonmitochondrial TPMP+
binding and was used to calculate the nonmitochondrial
TPMP+ binding correction.
Correction factors for the calculation of m.
To calculate m from the accumulation of
TPMP+, corrections had to be made for mitochondrial and
cellular volumes; resting plasma membrane potential
(p); and mitochondrial, nonmitochondrial and
extracellular binding (am, ac, and
ae).
p, however, was significantly (P < 0.01) greater (more negative) in hepatopancreas cells from estivating
snails (Table 1). This might have been due either to an increase in the
leakiness of the plasma membrane to K+ or, more likely, to
a decrease in leakiness of the plasma membrane to Na+ in a
phenomenon known as channel arrest (29). Sodium leak
decreases, for example, in turtle brain during anoxia
(39). The nonmitochondrial TPMP+-binding
correction was higher in cells from estivating snails (Table 1); this
was probably an experimental artifact. The modular kinetics in cells
from active snails were calculated using both active and estivating
correction factors, with similar results.
Kinetics of substrate oxidation, proton leak, and ATP turnover in
hepatopancreas cells from active and estivating snails.
Figure 3 shows the titrations of
respiration in cells from active (Fig. 3A) and estivating
snails (Fig. 3B). Mitochondrial respiration rate in cells
from estivating snails decreased significantly (P < 0.0005) to 35 ± 7% of control values in agreement with Bishop and Brand (3). Figure 3 shows that there was also a
significant (P < 0.005) decrease in the resting
m, from 97 to 76 mV.
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m.
As proton leak rate decreases with decreasing m, the
drop in m in cells from estivating snails (Figs. 3
and 4A) results in a decrease in the respiration rate
driving proton leak. The proton leak kinetics in cells from active and
estivating snails overlapped over the measured range of
m (Fig. 4B) and we assumed they also did
so at normal cellular m. Therefore, the decrease in
respiration driving cellular proton leak in cells from estivating snails was caused only by the drop in m and not by
any change in proton conductance.
As ATP turnover rate also decreases with decreasing
m, the drop in m resulted in a
decrease in the respiration rate driving ATP turnover during
estivation. The activity of the ATP turnover module, determined using
either an exponential or a straight line fit for proton conductance
(Fig. 4C; solid line and dotted line, respectively),
appeared to decrease during estivation. However, it is hard to make
conclusive statements, because the kinetics of ATP turnover hardly
overlapped in cells from active and estivating snails. Therefore, the
respiration driving ATP turnover decreased during estivation as a
result of the drop in m. In addition, the kinetics of
ATP turnover may have changed during estivation to further decrease ATP turnover.
When the electron transport chain was completely inhibited using
myxothiazol, cells from both active and estivating snails maintained a
higher m than they did when oligomycin was also present (Fig. 3). This was because the F1Fo-ATP
synthase acted in reverse as an oligomycin-sensitive ATPase using
glycolytically produced ATP to maintain m. This
m was significantly (P < 0.01) lower
in cells from estivating snails (52 mV) than in those from active
snails (76 mV). This may have been because glycolysis was turned down
during estivation, reducing the supply of ATP, or because the ATPase
itself was inhibited during estivation.
Relative contributions of proton leak and ATP turnover to resting
cellular respiration in hepatopancreas cells from active and estivating
snails.
The respiration used to drive both proton leak and ATP turnover
decreased during estivation. This was true whether the cellular proton
leak was calculated from oligomycin inhibition of respiration (Fig.
5A), from the exponential fit
to proton conductance (Fig. 5C; but note the large errors),
or from the straight line fit to proton conductance (Fig.
5E).
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Metabolic control analysis to quantify the effects of estivation
acting through m producers and m
consumers.
Mitochondrial respiration was divided into two modules:
m producers (substrate oxidation) and
m consumers (ATP turnover and proton leak), see Table
2. Coefficients that quantify control and
regulation were calculated as described by Brand (6),
Korzeniewski et al. (31), and Ainscow and Brand
(1).
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m describe
how strongly a small change in m affects their rates:
they encapsulate the kinetics of each module at defined
m. The elasticity coefficients of the
m producers were similar in cells from active and
estivating snails. The elasticity coefficient of the
m consumers, however, was higher in cells from active
snails than in cells from estivating snails, because the
m consumers in cells from active snails were more
responsive to changes in m than those from estivating snails.
Control coefficients of m producers or consumers over
mitochondrial respiration describe how strongly small changes in the activities of the modules affect mitochondrial respiration rate: they
quantify the control each module exerts over the rates in the system.
Table 2 shows that control over respiration rate was shared between the
m producers and the m consumers.
The m producers had slightly more control in cells
from active snails. The balance shifted in cells from estivating
snails, where the m consumers had slightly more control.
The proportional activation coefficient (31) is shown in
Table 2. It describes how much the intrinsic activity of the
m consumers is changed by estivation relative to the
intrinsic activity change of the m producers; it is
the ratio of the integrated elasticity coefficients of the modules to
estivation. Expressed in terms of the percentage of the total
inhibition of the modules, 69% occurred through the
m producers and 31% through the m consumers.
Partial integrated response coefficients describe how much the change
in mitochondrial respiration due to estivation is caused by changes in
each module. They take into account both the intrinsic changes in the
activities of the m producers and consumers in estivation, as well as the control that each module exerts over mitochondrial respiration. The partial integrated response coefficients (Table 2) showed that 75% of the response of mitochondrial respiration in hepatopancreas cells to estivation was through the
m producers. Only 25% of the response occurred via
the m consumers (in this case, via ATP turnover only).
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DISCUSSION |
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TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES
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This study demonstrates for the first time the subcellular basis
for the intrinsic decrease in mitochondrial respiration during metabolic depression in an isolated cell system: hepatopancreas cells
from H. aspersa. The activity of the m
producers (substrate oxidation) decreased in response to estivation.
This contributed to a threefold decrease in respiration and lowered
m. In turn, the drop in m caused
the respiration used to drive both the proton leak and ATP turnover to
decrease. This did not involve a change in the kinetics of proton leak,
but may have involved a decrease in the activity of ATP turnover.
Although the respiration to drive proton leak and ATP turnover
decreased, they did so to the same extent as the overall decrease in
mitochondrial respiration, so they accounted for similar proportions of
mitochondrial respiration before and during estivation, thus
conserving metabolic efficiency in the hypometabolic state. At least
75% of the response of mitochondrial respiration to estivation was due
to primary changes in the kinetics of substrate oxidation, with the
remaining 25% probably being due to primary changes in the kinetics of
ATP turnover.
Kinetics of substrate oxidation. The activity of substrate oxidation decreased in cells from estivating H. aspersa compared with active controls, contributing to an almost threefold reduction in respiration. This is similar to isolated mitochondria from frogs (49) where the proton conductance was measured, and in ground squirrels (14, 15, 17, 21), hamsters (42), and gophers (16), where the uncoupled rates were measured and the activity of substrate oxidation was found to decrease in hibernation.
The decrease in the activity of substrate oxidation may have been due to a decrease in the level of substrates that are endogenous to the cells, such as glycogen and fatty acids. However, the survival of animals during metabolic depression strongly correlates with their ability to reduce the rate of substrate use, so this mechanism appears unlikely. Alternatively, the activity of the substrate oxidation module may have decreased through a decrease in the activities of some of its constituent enzymes and transporters. Indeed, the activities of citrate synthase and cytochrome-c oxidase, enzymes involved in aerobic respiration, decrease to ~65% of control values in mitochondria isolated from the hepatopancreas of estivating H. aspersa (Bishop et al., unpublished observations). These enzymes also decrease during estivation in the hepatopancreas of the related land snail Cepaea nemoralis (51, 52) and during hibernation in skeletal muscle of the frog Rana temporaria (48). NADH dehydrogenase and succinate dehydrogenase also decrease during hibernation in the frog (48), and succinate dehydrogenase decreases during hibernation in the ground squirrel Citellus tridecemlineatus (55). Thus it appears that the activity of several enzymes involved in aerobic respiration decrease in metabolic depression. This is in accordance with the concept that control of oxidative phosphorylation is distributed fairly evenly among its constituent enzymes (23). Many studies have also looked at the activities of enzymes that are involved in anaerobic metabolism or glycolysis. The activities of hexokinase, aldolase, glyceraldehyde-3-phosphate dehydrogenase, phosphofructokinase, pyruvate kinase, and others decrease during metabolic depression in many animals [for example, land snails (12, 13, 54), turtles (11), frogs (22)]. In summary, many enzymes that form the backbone of substrate oxidation (those involved in glycolysis, the tricarboxylic acid cycle, and the electron transport chain) decrease in activity in metabolic depression, possibly decreasing the activity of substrate oxidation coordinately, thereby ensuring homeostasis of intermediates. However, the relevance of particular steps to the changed kinetics of the substrate oxidation module remains to be investigated.Kinetics of proton leak. The kinetics of mitochondrial proton leak, as measured in isolated hepatopancreas cells, did not change as a result of estivation in H. aspersa. This is similar to isolated mitochondria from the skeletal muscle of the frog R. temporaria, which show no change in the kinetics of proton leak in response to hibernation (49). It is also similar to isolated mitochondria from the liver of the ground squirrel Spermophilus lateralis, in which the state 4 rate, an indicator of the proton leak, did not change during hibernation (35).
Despite the lack of change in the kinetics of proton leak, the respiration driving proton leak decreased, as a simple downstream effect of the drop inm (in turn caused by the
decreased activity of substrate oxidation). Proton leak decreased in
line with total mitochondrial respiration, so the proportion of
mitochondrial respiration driving proton leak was similar before and
during estivation. Proton leak makes up a substantial proportion of
metabolic rate in many different animals, perhaps 20-30%
(7, 9, 50). Despite this, the function of proton leak
remains elusive, although it may play a role in decreasing production
of reactive oxygen species (47). This may be important
during emergence from metabolic depression in hepatopancreas cells from
snails, where there is a sudden increase in extracellular oxygen
tension and maybe, therefore, in production of reactive oxygen species;
protection from such species might increase the chances of survival.
Indeed, the activities of antioxidant enzymes, such as superoxide
dismutase and catalase, increase during estivation in the land snail
Otala lactea (28).
Kinetics of ATP turnover.
The drop in m in estivation led to a decrease in the
respiration driving ATP turnover. In addition, the activity of the ATP
turnover module may have decreased, although this conclusion should be
treated with caution as the data are at the limits of the sensitivity
of the assay. The respiration to drive ATP turnover decreased to the
same extent as the drop in total mitochondrial respiration, leaving the
proportion unchanged.
m and ATP (20). Stable (or only
slightly decreased) levels of ATP have been reported in many of these
systems during metabolic depression (19, 33, 38),
suggesting that ATP turnover rate decreases at least in part by a
reduction in the activities of specific components of the ATP turnover
module. Matched depression of the activities of ATP supply and demand
would ensure homeostasis of ATP during metabolic depression. However,
in hepatopancreas cells from H. aspersa there is a marked
drop in m during metabolic depression, so it is most
unlikely that these cells maintain ATP levels.
Relative importance of changes in the kinetics of substrate oxidation, proton leak, and ATP turnover. Seventy-five percent of the response of mitochondrial respiration to estivation was due to a primary decrease in the activity of substrate oxidation. Twenty-five percent or less was due to a primary decrease in the kinetics of ATP turnover. The kinetics of proton leak did not change and, therefore, did not play a primary role in the response of mitochondrial respiration to estivation.
It was surprising that changes in the kinetics of substrate oxidation were at least three times more important than changes in the kinetics of ATP turnover for the response of mitochondrial respiration to estivation. This implies that changes in individual ATP consumers, including protein synthesis and sodium cycling, cannot be very important in causing metabolic depression. Control over the rate of ATP turnover is widely distributed among the ATP consumers: no single process has more than one-third of the control in rat thymocytes (20). This makes it unlikely that any one ATP consuming process is responsible for >10% of the response of mitochondrial respiration to estivation. Therefore, a decrease in any of the ATP consuming processes, such as protein turnover or channel arrest and decreased sodium cycling, does not appear to be very significant to the overall metabolic depression in this model system.Perspectives
Until recently, many studies on metabolic depression have focused on glycolysis and the mechanisms by which glycolytic flux is turned down. Historically, this is because important model systems involve hypoxia- or anoxia-induced hypometabolism. Many organisms, however, still rely on mitochondrial metabolism during metabolic depression. Does mitochondrial metabolism play a role in metabolic depression in these organisms, and, if so, how is it turned down?In mitochondria isolated from frogs (49), squirrels (14, 15, 17, 21), hamsters (42), and gophers (16), the activity of the substrate oxidation module decreases during metabolic depression. This suggests that decreased activity of substrate oxidation is general in metabolic depression. The decrease in the activity of substrate oxidation in isolated mitochondria from frogs results in a drop in membrane potential, thus decreasing proton leak rate; the kinetics of proton leak, however, remain constant (49). Does this still hold true in cells? This project has shown that, in cells isolated from snails, the activity of substrate oxidation (and probably ATP turnover) decreased and the kinetics of proton leak remained constant, suggesting that results obtained using isolated mitochondria may also apply to cells in general.
Two of the model systems for metabolic depression (cells isolated from
snails during estivation and mitochondria isolated from frogs during
hibernation) therefore appear to use a similar mechanism for metabolic
depression. Inhibition of the upstream module (substrate oxidation) is
the main cause of decreased respiration, and the consequent drop in
m lowers proton leak rate and ATP turnover as
secondary events. There is no primary change in one downstream module
(proton leak), but there may be a smaller primary change in the other
downstream module (ATP turnover). One might have predicted that to
maintain m (and prevent it from dropping so low as to
cause cell death), the activities of all three modules would be shut
down to the same extent. The fact that the kinetics of proton leak do
not change during metabolic depression suggests either that proton leak
is not readily decreased during entry into metabolic depression or that
it is not easily reversed during arousal. The mechanism of the basal
proton leak remains elusive, although simple diffusion of protons
across the unperturbed phospholipid bilayer is not sufficient to
explain proton leak and other attributes of the mitochondrial membrane
must be involved (50). For example, membrane proteins may
play a role, either through nonspecific effects or by acting as
catalysts. If the kinetics of proton leak were not changed because the
proton leak is not readily altered, however, this suggests that basal
proton leak is not catalyzed by a specific protein but that it occurs
through less easily regulated processes.
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ACKNOWLEDGEMENTS |
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We thank J. A. Stuart, S. J. Roebuck, and J. A. Buckingham for help and advice.
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FOOTNOTES |
|---|
Trinity Hall, Cambridge, the Cambridge Commonwealth Trust (to T. Bishop), and the Medical Research Council (to M. D. Brand and J. St-Pierre) provided financial support.
Address for reprint requests and other correspondence: T. Bishop, The Henry Wellcome Building of Genomic Medicine, Roosevelt Dr., Oxford, OX3 7BN, UK (E-mail: tammie{at}well.ox.ac.uk).
The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
10.1152/ajpregu.00401.2001
Received 12 July 2001; accepted in final form 1 October 2001.
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REFERENCES |
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TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
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, Control;
, inhibited with myxothiazol, oligomycin, and FCCP
after 150 min.
,
respectively), and a line was applied to the 2 points (dotted line).
Respiration rates were measured at 80% air saturation, after which 5 µM myxothiazol was added to the cells. Mitochondrial respiration
rates were calculated as the total cellular rate minus the
myxothiazol-inhibited rate at 80% air saturation. *Standard condition
of the cells without inhibitors. Values are means ± SE,
n = 6 (A) or 5 (B).
