Primary causes of decreased mitochondrial oxygen consumption ...

Am J Physiol Regulatory Integrative Comp Physiol 282: R372–R382, 2002; 10.1152/ajpregu.00401.2001.

Primary causes of decreased mitochondrial oxygen consumption during metabolic depression in snail cells TAMMIE BISHOP, JULIE ST-PIERRE, AND MARTIN D. BRAND Medical Research Council, Dunn Human Nutrition Unit, Cambridge CB2 2XY, United Kingdom Received 12 July 2001; accepted in final form 1 October 2001

Bishop, Tammie, Julie St-Pierre, and Martin D. Brand. Primary causes of decreased mitochondrial oxygen consumption during metabolic depression in snail cells. Am J Physiol Regulatory Integrative Comp Physiol 282: R372–R382, 2002; 10.1152/ajpregu.00401.2001.—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

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 samMANY ORGANISMS SURVIVE

Address for reprint requests and other correspondence: T. Bishop, The Henry Wellcome Building of Genomic Medicine, Roosevelt Dr., Oxford, OX3 7BN, UK (E-mail: [email protected]). R372

pling 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 deThe 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.

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METABOLIC DEPRESSION IN HEPATOPANCREAS CELLS FROM HELIX ASPERSA

Fig. 1. Energy metabolism in isolated cells. The whole of cellular metabolism is divided into 3 modules: the first module, substrate oxidation, produces a mitochondrial membrane potential (⌬␺m), which is then consumed by the second module, proton leak, and the third module, ATP turnover. The system can also be divided into 2 modules: the first module, the ⌬␺m producers, consists of substrate oxidation, the second module, the ⌬␺m consumers, consists of both proton leak and ATP turnover.

creases 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. MATERIALS AND METHODS

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 ␮g 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/[3HTPMP⫹]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) ⌬␺m ⫽ 59 log

冏再

冎冏

vmac 关Cl ⫺ 兴tot关TPMP ⫹ 兴totac共vc ⫹ vm兲 ⫺1 vcam 关Cl ⫺ 兴e关TPMP ⫹ 兴eaevc

(1)

where v is volume; a is apparent TPMP⫹ activity coefficient or TPMP⫹ binding correction; and subscripts c, m, e, and tot are cytoplasmic ⫹ nuclear, mitochondrial, extracellular, and total intracellular, respectively. 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 ␮Ci/ml) and 1 ␮M carrier TPMP⫹, 0.1 ␮Ci/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 ␮M carbonyl cyanide p-trifluoromethoxyphenylhydrazone (FCCP), oligomycin at 2 ␮g/106 cells, and 2.5 ␮M 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 3HTPMP⫹, equivalent to cell volume multiplied by TPMP⫹ accumulation ratio). Cell volume. Cell volume was measured according to Brand (5). 3H2O (1 ␮Ci/ml) and 0.1 ␮Ci/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 under-

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estimated cell volume). Mitochondrial volumes (vm) were calculated using the cell volume and the mitochondrial-tocellular 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 ␮Ci/ml) and 1 ␮Ci/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 ␮M 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 ␮Ci/ml 3HTPMP⫹ (for ⌬␺m) or an equivalent volume of H20 (for respiration), plus 1 ␮M carrier TPMP⫹ as well as various inhibitors: To determine the kinetics of substrate oxidation, ⌬␺m was changed by addition of the uncoupler FCCP (0, 3.5, and 7 ␮M). 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 ␮M) in the presence of oligomycin (an inhibitor of F1FO-ATP synthase and, therefore, of ATP turnover) at 2 ␮g/106 cells. To determine the kinetics of the ⌬␺m consumers, ⌬␺m was changed by addition of myxothiazol (0 and 2.5 ␮M). FCCP, myxothiazol, and oligomycin were dissolved in DMSO; a total of 3 ␮l 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 ␮M 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. Oligomycin inhibits the ⌬␺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.

Table 1. Cell properties used for calculation of ⌬␺m in hepatopancreas cells from active and estivating snails Snail

Mitochondrial volume [vm/(vm ⫹ vc)], % of cellular volume Cell volume, ␮l/106 cells Resting plasma membrane potential (⌬␺p, mV) Mitochondrial TPMP⫹ binding correction (am) Nonmitochondrial TPMP⫹ binding correction (ac) External TPMP⫹ binding correction (ae)

Rat

Active

Estivating

22 4 ⫺32 0.44 0.21 0.7

2.3 ⫾ 0.1 (6) 1.9 ⫾ 0.1 (6) ⫺34 ⫾ 1 (5) 0.44 0.36 ⫾ 0.03 0.8

2.4 ⫾ 0.2 (6) 1.7 ⫾ 0.3 (4) ⫺40 ⫾ 2 (3)* 0.44 0.66 ⫾ 0.11 0.8

Values are means ⫾ SE; n, numbers in parentheses. Rat data were from Porter and Brand (40); snail mitochondrial volume data were from T. Bishop, A. Ocloo, and M. D. Brand (unpublished observations); rat cell volume was converted from ␮l/mg dry mass of cells using the value in Berry et al. (2); all mitochondrial TPMP⫹ binding correction data were from Nobes et al. (37), with the snail data assumed; and snail nonmitochondrial TPMP⫹ binding corrections were calculated from snail hepatopancreas cell volumes, resting plasma membrane potentials, and TPMP⫹ spaces in cells whose ⌬␺m has been collapsed (Fig. 2). * Significantly different from active (P ⬍ 0.05). AJP-Regulatory Integrative Comp Physiol • VOL

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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. Student’s t-test was used to test for significant differences between active and estivating snail hepatopancreas cells. P values ⬍0.05 were considered significant. RESULTS

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. ⌬␺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).

Fig. 2. Equilibration of triphenylmethylphosphonium (TPMP⫹) into Helix aspersa hepatopancreas cells from active (A) and estivating (B) snails. 3H-labeled TPMP⫹ was added at 0 min; at 150 min (arrow) 2.5 ␮M myxothiazol, oligomycin at 2 ␮g/106 cells, and 80 ␮M carbonyl cyanide p-trifluoromethoxyphenylhydrazone (FCCP; all in DMSO) were added; an equivalent volume of DMSO was added to controls, as described in MATERIALS AND METHODS. Values are means ⫾ range, n ⫽ 2 (A), or means ⫾ SE, n ⫽ 3 (B). 䊐, Control; ■, inhibited with myxothiazol, oligomycin, and FCCP after 150 min.

Neither mitochondrial nor cellular volume differed significantly between cells from active and estivating snails (Table 1). ⌬␺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


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Figure 4A shows that there was a large decrease in the activity of the substrate oxidation module in cells from estivating snails. This is indicated by a large decrease in the rate of substrate oxidation at any value of ⌬␺m. As proton leak rate decreases with decreasing ⌬␺m, the drop in ⌬␺m in cells from estivating snails (Figs. 3

Fig. 3. Modular kinetics of oxidative phosphorylation in H. aspersa hepatopancreas cells from active (A) and estivating (B) snails. Cells prepared from active and estivating snails were incubated as described in MATERIALS AND METHODS. Kinetics of substrate oxidation (䊐) were established by changing ⌬␺m by titration with FCCP (0, 3.5, and 7 ␮M), and a line was applied using linear regression. Kinetics of proton leak (■) were established by changing ⌬␺m by titration with myxothiazol (0, 0.25, 0.5, and 2.5 ␮M) in the presence of oligomycin at a concentration of 2 ␮g/106 cells, and both an exponential curve and a linear regression were applied. Kinetics of the ⌬␺m consumers were established by changing ⌬␺m by titration with myxothiazol (0 and 2.5 ␮M, * and E, respectively), and a line was applied to the 2 points (dotted line). Respiration rates were measured at 80% air saturation, after which 5 ␮M 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).

(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. The titrations in Fig. 3 were used to calculate the modular kinetics shown in Fig. 4. Figure 4A shows the kinetics of substrate oxidation in cells from active and estivating animals, Fig. 4B shows the kinetics of proton leak, and Fig. 4C shows the kinetics of ATP turnover.

Fig. 4. Kinetics of substrate oxidation (A), proton conductance (B), and ATP turnover (C) in hepatopancreas cells from active and estivating H. aspersa. Data for substrate oxidation (A) was taken directly from Fig. 3, A and B. Data for proton conductance (B) was taken from Fig. 3, A and B, with a single exponential (solid line) or linear fit (dotted lines) to the combined data for cells from estivating and active snails. ATP turnover (C) was calculated by subtracting the proton leak rate in B [from either exponential (solid lines) or linear fits (dotted lines)] from the rate of the ⌬␺m consumers (from Fig. 3, A and B) at the same ⌬␺m. *Standard condition of the cells without inhibitors. Values are means ⫾ SE, n ⫽ 6 (awake) or 5 (estivating) for A and B and means n ⫽ 6 (awake) or 5 (estivating) for C. 䊐, active; ■, estivating.


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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). The relative contributions of proton leak and ATP turnover to resting mitochondrial respiration, however, did not change greatly during estivation. This was true regardless of how proton leak was estimated (Fig. 5, B, D, F); the apparent changes using the straight line fit to proton conductance (Fig. 5F) were not significant. Overall, therefore, proton leak and ATP turnover each caused around one-half of mitochondrial respiration and no large changes in their relative contributions were apparent during estivation. 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

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2. Coefficients that quantify control and regulation were calculated as described by Brand (6), Korzeniewski et al. (31), and Ainscow and Brand (1). Elasticity coefficients of the modules to ⌬␺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). DISCUSSION

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


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Fig. 5. Processes contributing to the mitochondrial respiration rate of hepatopancreas cells from active and estivating H. aspersa. Proton leak and the corresponding ATP turnover were calculated using the oligomycin-inhibited respiration (A, B); using the exponential fit to proton conductance (C, D); and using the linear fit to proton conductance (E, F). A, C, E present absolute rates; B, D, F present rates expressed as a percentage of the total mitochondrial respiration. Data for A and B are from Fig. 3; data for C–F are from Fig. 4. Shaded bars, respiration driving proton leak; open bars, respiration driving 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 dehydroge-


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Table 2. Metabolic control analysis of mitochondrial respiration in hepatopancreas cells from active and estivating snails Mitochondrial Respiration (respirationm)

⌬␺m producers (substrate oxidation)

3⌬␺m 3

⌬␺m consumers (ATP turnover ⫹ proton leak) Active

Estivating

⫺3.14

⫺3.45

3.97

2.45

0.57

0.42

0.43

0.59

Elasticity coefficients ⌬␺ producer

ε⌬␺m m

⌬␺ consumer ␧⌬␺m m

Control coefficients respirationm producer

C⌬␺

m

respirationm C⌬␺ consumer m

Proportional activation ⌬␺ producer 䡠 ⌬␺mconsumer

m Pestivation

0.45

Partial integrated response coefficients ⌬␺mproducers

respirationm

IR estivation

⌬␺m consumers

respirationm

IRestivation

0.75 0.25

Calculations are described in the text. Average values (n ⫽ 6 active, n ⫽ 5 estivating) for ⌬␺m producers (substrate oxidation) and ⌬␺m consumers (ATP turnover ⫹ proton leak) from data in Fig. 3. Partial integrated response coefficients have been scaled to a sum of 1.0.

nase 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

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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 in ⌬␺m (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. ATP turnover in mammalian cells is responsible for ⬃75% of respiration. Approximately 20% of respiration is dedicated to protein synthesis and breakdown and 20% to sodium cycling (43, 45; reviewed in 44). The rate of protein synthesis and the activity of the Na⫹-K⫹ATPase have been well studied and shown to decrease during metabolic depression in hepatocytes isolated from turtles and goldfish (18, 32, 34). Decreased rates of various ATP consumers such as protein synthesis may be caused by a decrease in their activity or may simply be the downstream effect of a drop in ⌬␺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.


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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 hypoxiaor 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 (sub-

strate 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. We thank J. A. Stuart, S. J. Roebuck, and J. A. Buckingham for help and advice. 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. REFERENCES 1. Ainscow EK and Brand MD. Quantifying elasticity analysis: how external effectors cause changes to metabolic systems. Biosystems 49: 151–159, 1999. 2. Berry MN, Edwards AM, and Barritt GJ. Isolated Hepatocytes Preparation, Properties and Applications. Amsterdam, The Netherlands: Elsevier, 1991, p. 127. 3. Bishop T and Brand MD. Processes contributing to metabolic depression in hepatopancreas cells from the snail Helix aspersa. J Exp Biol 203: 3603–3612, 2000. 5. Brand MD. Measurement of mitochondrial protonmotive force. In: Bioenergetics—A Practical Approach, edited by Brown GC and Cooper CE. Oxford, UK: Oxford University Press, 1995, p. 39–62. 6. Brand MD. Top down metabolic control analysis. J Theor Biol 182: 351–360, 1996. 7. Brand MD, Bishop T, Boutilier RG, and St-Pierre J. Mitochondrial proton conductance, standard metabolic rate and metabolic depression. In: Life in the Cold, edited by Heldmaier G and Klingenspor M. Berlin: Springer-Verlag, 2000, p. 413–430. 8. Brand MD, Boutilier RG, Wass M, St-Pierre J, and Bishop T. Mitochondrial proton leak during metabolic depression. In: Molecular Mechanisms of Metabolic Arrest: Life in Limbo, edited by Storey KB. Oxford: BIOS Scientific, 2001, p. 59–74. 9. Brand MD, Chien LF, Ainscow EK, Rolfe DFS, and Porter RK. The causes and functions of mitochondrial proton leak. Biochim Biophys Acta 1187: 132–139, 1994. 10. Brookes PS, Buckingham JA, Tenreiro AM, Hulbert AJ, and Brand MD. The proton permeability of the inner membrane of liver mitochondria from ectothermic and endothermic vertebrates and from obese rats: correlations with standard metabolic rate and phospholipid fatty acid composition. Comp Biochem Biophys 119B: 325–334, 1998. 11. Brooks SPJ and Storey KB. Regulation of glycolytic enzymes during anoxia in the turtle Pseudemys scripta. Am J Physiol Regulatory Integrative Comp Physiol 257: R278–R283, 1989.


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