Preservation of metabolic reserves and function after storage of myocytes in hypothermic UW solution

Julia O. Hegge, James H. Southard, and Robert A. Haworth

Department of Surgery, University of Wisconsin, Madison, Wisconsin 53792


    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Isolated rat myocytes cold stored anaerobically up to 24 h in University of Wisconsin solution lost 95% of their ATP and 100% of their glycogen. They underwent contracture when rewarmed in a Krebs-Henseleit (KH) medium that contained Ca unless Ca addition was delayed. In the latter case, cell function, measured by stimulation-induced cell shortening, was surprisingly well retained. Aerobically stored cells were resistant to Ca on rewarming, although 96% of glycogen was still lost, along with 46% of ATP. Cells that were incubated for 48 h aerobically with the substrates glucose and pyruvate at pH 6.2 retained 77% of their ATP and 59% of their glycogen, with good cell morphology. At pH 6.2, the demand for ATP was only 55% of that at pH 7.4. However, after rewarming, these cells functioned no better than anaerobically stored cells, although their inotropic response to isoproterenol was improved. We conclude that 1) aerobic conditions with substrates at low pH preserve myocyte metabolic reserves well for 48 h, partly by reducing the demand for ATP; 2) rewarming conditions are critical for anaerobically stored cells with metabolic stores that are severely depleted; and 3) unloaded cell function is surprisingly insensitive to a period of severe metabolic deprivation.

glycogen; adenosine 5'-triphosphate; morphology; cell shortening; reperfusion conditions; University of Wisconsin solution


    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

CLINICALLY, HEART PRESERVATION for transplantation is limited to 4-6 h. This limitation may possibly reflect a need for maintaining some level of ATP, which is used by the myocytes to maintain intracellular homeostasis and a functional contractile apparatus. Unlike other organs, if ATP is severely depleted, the heart can develop an irreversible contracture during preservation or upon reperfusion. During cold storage, tissue PO2 falls quickly, and the heart must rely on glycolysis and glycogen stores for ATP production. Cold storage solutions like University of Wisconsin (UW) solution are thought to prevent cell swelling and minimize oxidative stress, but because of inherent limitations of oxygen supply, there is little that can be done during ischemic cold storage to improve energy reserves. To extend preservation and maintain adequate ATP levels by oxidative phosphorylation, it could be advantageous to use continuous perfusion during preservation. Experimental and clinical studies have shown that machine perfusion is superior to cold storage for kidneys (2), and recent studies suggest that this may be true for the heart (8, 34, 36).

Because function is closely linked to an adequate supply of energy from metabolism, metabolic impairment results in functional impairment of the heart (39). Transplanted hearts are required to function immediately. It is therefore reasonable to think that the ideal preservation solution for machine perfusion would minimize the use of energy reserves so that these are available to meet the demands of reperfusion and mechanical work. Previous (unpublished observations) studies in this laboratory have shown that even under aerobic storage conditions with adequate substrates, organs such as kidneys and livers lose their glycogen stores. Evidence comes from several studies indicating that conditions that attenuate the use of glycogen during preservation could result in better graft function. Lagerstrom et al. (28) showed that rabbit hearts depleted of glycogen before ischemia recovered poorly compared with hearts in which glycogen was preserved. Another study by Cross et al. (10) showed that glycogen was, on one hand, detrimental because its depletion resulted in acidosis, but on the other hand, beneficial because it delayed the onset of ischemic contracture. Thus high glycogen can be beneficial because it extends the time to onset of contracture, but it can also be detrimental in postcontracture hearts because the degree of acidosis is more severe. Hence, the impact of high glycogen depends on the time of ischemia. Finally, Ichihara et al. (25) found that glycogen, when well preserved during hypothermia, provided an important source of energy early during reperfusion. There can also be some correlation between ATP level and function in hearts reperfused after hypothermic storage (47) or normothermic global ischemia (12). This may, however, be more a reflection of the damage suffered than an ATP requirement for function. Studies in which ATP levels are depleted by treatment of hearts with 2-deoxyglucose show little correlation between ATP level and function and suggest, rather, that the ability to maintain a high phosphate potential may be more important (22, 26).

The preservation of glycogen during storage is determined in part by the supply and demand for ATP. Under aerobic conditions, an adequate supply of substrates for glycolysis and oxidative metabolism would theoretically ensure an ATP supply and reduce the need for breakdown of glycogen stores. Hypothermia and conditions that inhibit or reduce the rate of ATP hydrolysis would decrease the demand for ATP. In this study, we used isolated myocytes to investigate preservation conditions that would improve the supply of ATP and decrease the demand for ATP, with the aim of conserving metabolic reserves of glycogen. We found that glycogen reserves were well preserved only under aerobic storage conditions at low pH and with substrates.

To evaluate the impact of these metabolic reserves on myocyte function, the cells must first be restored to physiological conditions. A major concern is the pH paradox, which is manifested in neonatal rat cardiac myocytes as cell blebbing and death on restoration of normal pH after a period of normothermic metabolic deprivation at low pH (3). We therefore investigated the requirements for restoring hypothermically preserved cells to physiological conditions and the impact of well-preserved metabolic reserves on heart cell function, as measured by cell shortening in response to electric field stimulation. We found that the anaerobically stored cells, which had negligible metabolic reserves, were more susceptible to rounding up than aerobically preserved cells when restored to physiological conditions. Because Ca overload has been implicated in the pathophysiology of ischemia-reperfusion injury (27), we also investigated the impact of delaying the restoration of Ca. We found that by delaying the addition of Ca for 15 min while the cells rewarmed, the morphology and function of these cells could be successfully restored. Moreover, at longer preservation times, their function without catecholamines was as good as that of aerobically stored cells, although their inotropic reserve, measured by their response to isoproterenol, was impaired.

Isolated myocytes were used for these studies for three reasons. 1) Cell morphology provides a convenient way to access viability and contracture at the cellular level. 2) Preparations of myocytes from a single heart allow testing of multiple solutions and storage conditions. 3) Isolated myocytes are a good model for assessing the conditions required for good preservation of cardiac striated muscle in isolation from the added complication of preservation-induced vascular changes.


    MATERIALS AND METHODS
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

UW solution. UW solution is an organ preservation solution developed at the University of Wisconsin that is currently used worldwide (46). It is a high-K, low-Na solution with chloride replaced by lactobionate and also contains other additives, including hydroxyethyl starch (HES) for oncotic support (Table 1). The UW solution used in these studies did not contain HES because it interferes with the accurate measurement of glycogen. The amyloglucosidase enzyme used in the glycogen assay (see Glycogen content) reacts with HES to generate glucose units, which results in erroneously high glycogen measurements. When used with intact organs, HES is added to UW solution as a colloid to prevent endothelial cell swelling. In isolated myocytes, this is not a factor. In preliminary studies, we found no apparent difference in the cell morphology or ATP content of myocytes preserved with or without HES.

                              
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Table 1.   Composition of UW solution

HES-free UW solution was made in the laboratory with glutathione added just before each experiment and pH adjusted to 7.4. Additional pH adjustments were made by adding HCl for acidic conditions and NaOH for alkaline conditions. Adenine and ribose (where added) were dissolved in alkaline HES-free UW solution before pH adjustment. Other additions were made directly to HES-free UW solution after pH adjustment, and pH was checked thereafter.

Cell isolation. All animals in this study received humane care in accordance with the guidelines set forth by the National Institutes of Health (NIH) in the Guide for the Care and Use of Laboratory Animals (NIH Publication No. 86-23) and approved by the Animal Care Committee at the University of Wisconsin. Myocytes were isolated from male Sprague-Dawley rats (300 g each) as previously described (18) and modified (16). Cells were isolated from two rats for each experiment and pooled. Rats were purchased from Harlan Sprague Dawley. The perfusion medium was KH-HEPES (see KH-HEPES medium) at 37°C, adjusted to pH 7.0 with NaOH, plus basal amino acids. Hearts were perfused initially with 1 mM Ca for 5 min and then switched to Ca-free perfusate for 5 min before enzymes were added. Ca (1 mM) was added back to the recirculating perfusate 15 min after enzyme addition. Ten minutes later, the hearts were cut up, and the pieces were shaken for 30 min in medium with 1 mM Ca to free the cells. This procedure results in cells that tolerate Ca well. They are exposed only to zero (added) Ca medium for 20 min at 37°C, which limits Na loading, followed by 40 min with Ca to allow reestablishment of normal homeostasis (13, 15). At the completion of the cell isolation procedure, the suspension of myocytes was centrifuged, cells resuspended with ice-cold HES-free UW solution, centrifuged again, and resuspended in ice-cold HES-free UW solution. Protein concentration was determined by the biuret method and adjusted as needed. Previously, we found that 1 mg of protein corresponds to 192,070 ± 45,730 cells (18).

KH-HEPES medium. Basal KH medium contained (in mM) 118 NaCl, 4.8 KCl, 25 HEPES, 1.2 KH2PO4, 1.2 MgSO4, and 11 glucose, adjusted to pH 7.4 with NaOH.

Lactate production: rate of ATP utilization. Myocytes for these measurements were stored in HES-free UW solution without glucose, pyruvate, or lactate additions. Oxidative phosphorylation was inhibited by the addition of 3 µM rotenone at time zero. Myocytes were stored at 6°C as shown, and samples were taken at time intervals for the measurement of lactate, ATP, and %rods. Lactate was measured on perchloric acid extracts of aliquots of cell suspensions as previously described (20). Under these experimental conditions (no glucose added), 1 mol of lactate produced is equivalent to 1.5 mol of ATP synthesized. Each mole of glucose 6-phosphate uses 1 mol of ATP to give 1 mol of fructose 1,6-bisphosphate, and this ultimately yields 4 mol of ATP and 2 mol of lactate. Hence, the calculation used for the rate of ATP utilization was
ATP (nmol<IT>·</IT>mg<SUP>−1</SUP><IT>·</IT>min<SUP>−1</SUP>)<IT>=</IT>[lactate produced (nmol/mg)<IT>×</IT>1.5<IT>+</IT>ATP lost (nmol/mg)<IT>+</IT>CPr lost (nmol/mg)]/time interval (minutes)
The time interval used to calculate the initial rate of ATP utilization was from 0 to 60 min at 6°C. This interval was chosen because the rate of lactate production over this interval was linear and unaffected by glycogen exhaustion.

Glycogen content. Glycogen was determined by the method of Carr and Neff (9). Briefly, 1.5-ml aliquots of cell suspension were spun in a high-speed bench centrifuge, and the supernatants were discarded. Two hundred fifty microliters of citrate buffer (100 mM, pH 5.0) were added to the cell pellet, and the samples were sonicated and boiled for 5 min. The resulting slurry was centrifuged, and the background glucose was determined using a Yellow Springs Instrument glucose analyzer. After background glucose was measured, these samples were treated with 25 µl of 0.5% amyloglucosidase (giving 30 U/ml), vortexed, and incubated at 55°C for 2 h. After being incubated, the enzyme-treated samples were centifuged, and the glucose was determined by the same method. Background glucose corrections were made, and glycogen was expressed as micromoles of glucose equivalents per milligram of protein.

ATP and phosphocreatine content. ATP was measured in acid extracts of myocytes using a modification of the method by Sellevold et al. (42). Aliquots of cells were mixed with an equal volume of 1 M perchloric acid. These samples were centrifuged, and 300 µl of supernatant were neutralized with 150 µl of 2 M Trizma base to pH 7.3 and analyzed by high-performance liquid chromatography. Samples were eluted from a Supelcosil 25-cm × 4.6-mm, 5-µm particle column with an isocratic buffer that contained 175 mM KH2PO4, 1.85 mM tetrabutylammonium hydrogen sulfate, and 8% acetonitrile (pH 6.25 with KOH). For samples in which ATP alone was measured, the flow rate was 1.0 ml/min, and absorbance was measured at 254 nm. For samples in which both phosphocreatine (PCr) and ATP were measured, the flow rate was 0.8 ml/min, and absorbance was measured at 216 nm. Data were analyzed using a Waters Millennium software system.

Morphology. Aliquots of cell suspensions were mixed with an equal volume of 2.5% glutaraldehyde and examined with a light microscope. A total of 300 cells from each sample was counted as either rod shaped or nonrod shaped. Relaxed cells with a sarcomere length in the region of 1.8 µm were counted as rod shaped (%rod) (19).

Aerobic and anaerobic storage conditions. After being isolated, myocytes were suspended in ice-cold HES-free UW solution for 5 min, and baseline samples for measurement of ATP, glycogen, and %rods were taken. For the anaerobic conditions, a concentrated cell suspension (10 mg protein/ml) was used. One-milliliter aliquots of cells were placed in 1.5-ml microcentrifuge tubes, and the cells were allowed to settle and remain undisturbed during preservation at 6°C. Under these anaerobic conditions, the myocytes formed a pellet creating an environment in which HES-free UW solution and oxygen were not available. After storage, anaerobic cells were diluted to 2 mg/ml with cold UW solution and gently mixed before samples for ATP, glycogen, and cell morphology analysis were taken. For aerobic storage, cell suspensions were diluted to a protein concentration of 2 mg/ml, and a 5-ml aliquot of cells was placed in a flat-bottom flask (50-ml Erlenmeyer). The cells were allowed to settle without agitation at 6°C. In preliminary experiments, we tried shaking these cells to provide additional oxygenation but found that diluted cells that were not shaken maintained better ATP levels. From this, we concluded that unshaken cells formed a thin diffuse layer in the bottom of each flask, which created an environment in which preservation solution and oxygen were apparently readily available to each myocyte. We also concluded that shaking introduced other factors detrimental to myocyte preservation. Thus for our aerobic condition, we did not shake the cells. In preliminary experiments, we also established that there was no change in the percentage of cells that took up trypan blue under any of these conditions, suggesting that the integrity of the cell membrane was unchanged. After being stored, aerobic cells were gently mixed before samples for ATP, glycogen, and cell morphology analysis were taken.

Rewarming protocols. For studies on the effects of rewarming preserved myocytes, two different rewarming protocols were used. For both rewarming protocols, myocytes were centrifuged for 90 s at 29 g at 4°C, washed once with ice-cold KH-HEPES medium, pH 7.4 (see KH-HEPES medium), and finally resuspended to a volume of 5 ml. Protein measurements indicated that no significant amount of cells was lost during centrifugation, so no selection for rod-shaped cells occurred during this step. Cell suspensions were then transferred to 25-ml flasks, placed in a 37°C shaker bath, and allowed to rewarm for 30 min. In one protocol (immediate Ca), the rewarming medium also contained CaCl2 (1 mM) so that myocytes were exposed to Ca immediately after preservation. In the other protocol (delayed Ca), the Ca addition was delayed until myocytes had rewarmed for 15 min at 37°C, and then CaCl2 (1 mM) was added: Ca was added as 3 × 12.3-µl aliquots of 0.1 M CaCl2 to 3.7-ml cell suspension, added over a period of 90 s. Samples for measurement of morphology, ATP, and glycogen were taken at the end of preservation, after resuspension in ice-cold KH-HEPES, and after 15 and 30 min of rewarming at 37°C. Cell viability on rewarming was not examined because the preservation of cell morphology and function are more exacting preservation criteria (13).

Cell shortening. Cell shortening was measured by a photodiode array device as described by Boyett et al. (6) using a Nikon diaphot microscope. Cells were superfused at 37°C with KH-HEPES medium, 1 mM Ca, and 50 µM diethylenetriaminepentaacetic acid and field stimulated at 0.5 Hz with 2- to 3-ms biphasic pulses (20). Cells were attached to the dish with laminin; we have previously shown that such attachment does not affect the shortening parameters of cell populations (17). Only rod-shaped cells were measured, although both rod-shaped and round cells would attach. At least 10 rod-shaped cells were measured in each group in each experiment. Normal cell shortening measurements were made first, followed by 25 nM isoproterenol stimulation. Cells used for the shortening measurements without isoproterenol were not the same cells used for the shortening measurements with isoproterenol. Cell shortening was determined in a time period from 30 min to 2 h after the end of the rewarming protocol.

Statistics. All data shown are the mean values from three experiments ± SD between experiments. The data were analyzed with analysis of variance (ANOVA). When multiple treatments were applied to the same initial sample, the data were analyzed with a split-plot design to account for within-sample correlations. All analyses were performed with SAS Proc Mixed (SAS Institute, Cary, NC). If the ANOVA indicated a significant treatment effect, pairwise comparisons were performed by comparing population marginal means (least-squares means); this procedure is very similar to Fisher's protected least significant differences test. Synergy was tested for by first performing a linear contrast test for nonadditivity. For an effect to be declared synergistic, significant nonadditivity was needed, and the estimated effect had to be superadditive.

Chemicals. All chemicals were purchased from Sigma-Aldrich Chemical, St. Louis, MO.


    RESULTS
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Anaerobic vs. aerobic storage for 24 h. The first series of experiments was designed to measure the loss of metabolic reserves in cells stored anaerobically in HES-free UW and to determine whether aerobic cold storage in HES-free UW solution resulted in better preservation of metabolic reserves than anaerobic cold storage. Freshly isolated cells for this study contained 79 ± 4% rod-shaped cells, 25.8 ± 6.1 nmol ATP/mg, and a glycogen content of 0.164 ± 0.034 µmol glucose/mg. Myocytes were stored under anaerobic or aerobic conditions for 24 h at 6°C. Morphology, ATP, and glycogen samples were taken before and after preservation. After 24 h in HES-free UW solution at 6°C, the number of rod-shaped cells showed a small but significant decrease (Fig. 1A) in both the anaerobically and the aerobically stored cells, and the glycogen content (Fig. 1C) of both was depleted to near zero. The main difference between the two storage conditions was that the aerobic cells still contained 54% of their ATP, whereas the anaerobic cells contained only 5% (Fig. 1B).


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Fig. 1.   Effect of 24-h preservation in University of Wisconsin solution without hydroxyethyl starch (HES-free UW solution) under aerobic or anaerobic conditions on cell morphology and metabolic reserves. Cells were stored under aerobic or anaerobic conditions as described in MATERIALS AND METHODS, and cell morphology (A), ATP (B), and glycogen (C) were measured. Even though morphology was well preserved, glycogen was depleted during preservation under both conditions; n = 3 experiments.

Rewarming after anaerobic or aerobic storage in HES-free UW solution for 24 h. Myocytes were stored as indicated earlier under anaerobic or aerobic conditions for 24 h at 6°C without added substrates. Samples were taken before and after preservation for assessment of morphology, ATP, and glycogen, and separate cell suspensions were used for rewarming studies. To study the effects of rewarming on cell viability, cells were rewarmed using the two protocols (with or without Ca initially) described in MATERIALS AND METHODS. Figure 2 shows the effect of the immediate Ca protocol on fresh and preserved cells. Even fresh cells showed some decline in glycogen (Fig. 2C) and ATP (Fig. 2B), and these changes were significant on transfer of the cells from cold HES-free UW solution to cold KH-HEPES medium when Ca was present in the KH-HEPES medium. We were surprised by these declines because we had not thought this to be a metabolically demanding change for the cells: being cold, their metabolic demand is expected to be low, and since they were aerobic and fresh, their ability to supply that demand should be more than sufficient. Resuspension of fresh cells in the presence of Ca showed some variable loss of %rod-shaped cells, even before the cells were rewarmed, and this trend was not evident when cells were resuspended without Ca (Fig. 2A). By contrast, the anaerobically preserved cells that were rewarmed in the presence of Ca all went into contracture when the cells were warmed to 37°C (Fig. 3A), whereas aerobic cells rewarmed with Ca were less susceptible (Fig. 4A) and showed a loss of %rods similar to freshly isolated cells (Fig. 2A). Morphology of both the anaerobic and aerobic cells (Figs. 3A and 4A) was well preserved when the cells were rewarmed for 15 min without Ca, followed by Ca restoration. Under these conditions, 61% of the cells in both the aerobic and anaerobic conditions were rod shaped after 30 min at 37°C. This was similar to fresh cells, which had 62% rod-shaped cells when rewarmed initially without Ca (Fig. 2A). Anaerobically stored cells regained ATP (Fig. 3B) and significant but small levels of glycogen (Fig. 3C) when rewarmed initially without Ca, whereas ATP levels in aerobically stored cells were stable during rewarming. Thus after rewarming without Ca, whether they were stored anaerobically or aerobically, cells were similar both morphologically (Figs. 3A and 4A) and metabolically (Fig. 3, B and C, and Fig. 4, B and C). By contrast, rewarming anaerobically stored cells with the immediate Ca protocol gave cells that had as much ATP and glycogen as cells rewarmed without Ca (Fig. 3, B and C), but their morphology was completely different: cells rewarmed with immediate Ca had all rounded up, whereas cells rewarmed with the delayed Ca protocol retained good morphology (Fig. 3A). It should be noted, however, that after either rewarming protocol, ATP levels were only about one-half of starting levels (Figs. 2B and 3B), and glycogen levels were merely 10-15% of starting levels (Figs. 2C and 3C). Thus glycogen levels do not rebound quickly under these conditions, and ATP is only partially restored, whether or not the cells have retained good morphology. Because there was no loss of %rod-shaped cells under these rewarming conditions, these reduced levels must reflect changes in ATP and glycogen content in these rod-shaped cells.


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Fig. 2.   Effect of the rewarming protocol on fresh cells. Fresh cells were suspended in ice-cold HES-free UW solution for 15 min before resuspension in cold Krebs-Henseleit (KH)-HEPES. Cells were rewarmed using the immediate or delayed Ca protocols as described in MATERIALS AND METHODS, and changes in cell morphology (A), ATP (B), and glycogen (C) were measured. Some loss of glycogen was evident on transfer of cells to cold Ca-containing KH-HEPES; n = 3 experiments.



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Fig. 3.   Effect of the rewarming protocol on anaerobically stored cells. Anaerobically preserved cells (see MATERIALS AND METHODS) were stored in cold HES-free UW solution for 24 h before being resuspended in cold KH-HEPES. Cells were rewarmed using the immediate or delayed Ca protocol, and changes in cell morphology (A), ATP (B), and glycogen (C) were measured. Cells stored anaerobically rounded up with the immediate Ca protocol but not with the delayed Ca protocol; n = 3 experiments.



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Fig. 4.   Effect of the rewarming protocol on aerobically stored cells. Aerobically preserved cells (see MATERIALS AND METHODS) were stored in cold HES-free UW solution for 24 h before being resuspended in cold KH-HEPES. Cells were rewarmed using the immediate or delayed Ca protocol, and changes in cell morphology (A), ATP (B), and glycogen (C) were measured. Cells stored aerobically had similar morphology and ATP to fresh cells subjected to similar protocols; n = 3 experiments.

Because in clinical practice, hearts are preserved under anaerobic conditions, it is of interest to measure how well cell function is preserved under these conditions and to compare it with the function of aerobically stored cells whose metabolic reserves have not suffered such extreme variations. Thus measurements of cell shortening in response to electric field stimulation were made on the anaerobically and aerobically stored cells (Fig. 5, A and B). These cells were rewarmed initially without Ca, since this condition best preserved rod-shaped morphology (Fig. 4A). Freshly isolated myocytes served as controls and shortened to 11.4 ± 2.2% of resting length (Fig. 5A). After a 24-h storage, the %shortening had significantly decreased by 34%, but there was no difference between anaerobic and aerobic cells (7.8 ± 0.7% and 7.3 ± 2.7%, respectively). To gain a measure of the inotropic reserves of the cells, their response to isoproterenol, a mixed beta -adrenergic agonist, was also measured. With 25 nM isoproterenol, shortening increased by a factor of 1.5 in fresh cells (15.9 ± 0.5%) and by a similar factor in cells stored for 24 h (anaerobic = 10.9 ± 0.7%, aerobic = 11.8 ± 2.7%).


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Fig. 5.   Effect of preservation condition on cell contractile parameters. Preserved cells were restored to physiological conditions using the delayed Ca protocol. Cell shortening was measured in either fresh cells, 24-h aerobically preserved cells, or in 24-h anaerobically preserved cells. A: %shortening. B: shortening profile of a fresh cell with an isoproterenol (Isoprel)-stimulated cell superimposed. Cells stored by either protocol showed significant depression of contractile function either with or without isoproterenol; n = 3 experiments.

These results show that anaerobically stored cells, which have severely depleted metabolic reserves, are very susceptible to Ca overload on rewarming; however, a rewarming protocol that presumably prevents Ca overload results in cells with morphology, metabolic reserves, and unloaded shortening function essentially as good as that of aerobically stored cells.

Because the objective of our study was to develop conditions of hypothermic preservation that allowed metabolic reserves to be maintained, we investigated the effectiveness of modifications to the HES-free UW solution designed to conserve these reserves. Aerobic conditions were used because anaerobic conditions offer such limited potential for the preservation of metabolic reserves.

Effect of additives and pH on aerobic myocyte preservation. Glucose (11 mM) was added as a substrate for glycogen synthesis and glycolysis. Lactate (5 mM) was chosen as an exogenous substrate for oxidative phosphorylation, which has also been found to promote glycogen synthesis (29). Adenine (5 mM) plus ribose (5 mM) were also tested as precursors for adenine nucleotide synthesis, which has been found effective in the preservation of ATP levels in hypothermically perfused dog kidney (31) and rabbit heart (40). Because acidosis has been shown to be protective against normothermic anoxia and reoxygenation injury in a variety of cell types, including myocytes (4), we also investigated the effect of pH on hypothermic myocyte preservation. Cells were stored in the modified HES-free UW solution containing glucose, lactate, and adenine plus ribose, over a range of pH values from pH 6.0 to pH 8.0. The preservation time for these studies was increased to 48 h to better delineate differences between groups. At the end of preservation, %rods, glycogen, and ATP were measured.

After 48 h of aerobic storage, cell morphology and glycogen were well maintained at the acidic pH levels. The best results were obtained from myocytes stored at pH 6.6 or lower (Fig. 6, A and C). Glycogen and %rods significantly declined above pH 7.0, and this decline became more dramatic as the pH increased. ATP was well preserved at all pH levels (Fig. 6B). The highest ATP levels were found near pH 6.8 (98% remaining) and the lowest at pH 8.0 (63% remaining).


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Fig. 6.   Effect of HES-free UW solution pH on cell morphology and metabolic reserves after 48-h aerobic preservation in HES-free UW solution with additives. HES-free UW solution additionally contained 11 mM glucose, 5 mM lactate, 5 mM adenine, and 5 mM ribose. Changes in cell morphology (A), ATP (B), and glycogen (C) were measured as described in MATERIALS AND METHODS. Cells at low pH showed the best preservation of morphology and glycogen; n = 3 experiments.

In the previous experiments, the modified HES-free UW solution with a low pH provided the best overall preservation conditions. We then investigated which additives were having a beneficial effect and how pH may alter this effect. In addition, we tested the use of pyruvate as a substitute for lactate. For these experiments, glucose (11 mM), lactate (5 mM), or pyruvate (5 mM) and adenine plus ribose (5 mM each) were added individually or in combination with HES-free UW solution at either pH 6.2 or pH 7.4. Myocytes were stored aerobically for 48 h, and at the end of preservation, %rods, glycogen, and ATP were measured.

We found that myocytes stored at pH 6.2, in any of the solutions, showed only a small but significant change in morphology (Fig. 7A). In HES-free UW with no additives, the %rods were well maintained, even though the glycogen (Fig. 7C) and ATP (Fig. 7B) were severely depleted. In contrast, most of the myocytes stored at pH 7.4 had gone into contracture even when additional substrates were added. The ATP level (Fig. 7B) of cells stored in HES-free UW with no additives at pH 7.4 was also severely depleted, as it was at pH 6.2. At pH 7.4, the addition of glucose, lactate, or pyruvate either alone or in combination increased ATP levels significantly (P < 0.01). At pH 6.2, pyruvate alone or in combination with glucose also significantly increased ATP levels (P < 0.05) to the same extent as pH 7.4. At pH 6.2, adenine and ribose surprisingly had no significant effect on ATP or glycogen when added alone (data not shown) or when combined with other substrates (Fig. 7, B and C). Although additional substrates (glucose, lactate, or pyruvate) improved ATP content, the amount of ATP preserved was not very pH dependent. After 48 h, there were large differences in the amount of glycogen remaining in myocytes stored at pH 7.4 vs. pH 6.2 (Fig. 7C). In the absence of substrates, glycogen was severely depleted at both pH values (Fig. 7C). Glycogen was consistently low in myocytes stored at pH 7.4, whether or not substrates were added. The best results at pH 7.4 were obtained from myocytes stored with multiple substrates and ATP precursors, but even these cells had only 17% of their glycogen remaining. In contrast, at pH 6.2, pyruvate alone significantly increased glycogen, while glucose with pyruvate gave a further significant increase. A similar trend was also seen with lactate. After 48 h, 42-60% of the glycogen was still remaining when lactate or pyruvate was combined with glucose (Fig. 7C). The combined effect of glucose plus either pyruvate or lactate was beyond additive: a statistical test showed it to be synergistic (P < 0.02).


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Fig. 7.   Effect of additives on the preservation of morphology and metabolic reserves after 48-h aerobic preservation in HES-free UW solution. Additives were glucose (Gluc; 11 mM), lactate (Lact; 5 mM), pyruvate (Pyr; 5 mM), and adenine (Adn; 5 mM) plus ribose (Rib; 5 mM) in the combinations shown. Changes in cell morphology (A), ATP (B), and glycogen (C) were measured as described in MATERIALS AND MEthods. Glucose and either pyruvate or lactate acted synergistically to conserve cell glycogen but only at low pH. Open bars, pH 7.4; hatched bars, pH 6.2; n = 3 experiments; NS, not significant.

Effect of pH on the rate of ATP utilization. Because low pH reduced the rate of glycogen loss, we investigated how pH affected the rate of ATP utilization by cells. This was accomplished with a strategy used previously (14) in which the rate of ATP utilization could be measured from the rate of lactate production under conditions in which glycogen was the major source of ATP. This condition was achieved by eliminating oxidative phosphorylation by adding rotenone, so that glycolysis was the only source of ATP, and by excluding glucose from the medium, so that glycogen was the only substrate for glycolysis. Thus a fixed stoichiometry of 1.5 mol of ATP per mole of lactate could be assumed. Because ATP loss and PCr loss also contribute to the rate of ATP utilization, these also were measured and included in the calculation (see MATERIALS AND METHODS). These conditions are not intended to precisely emulate the conditions experienced by the cells under the anaerobic or aerobic incubation conditions of Fig. 1. They are designed, rather, to address the question of how the factor of reduced pH itself affects the rate at which the cells utilize ATP under conditions in which the supply of ATP is not rate limiting. Freshly isolated myocytes were stored aerobically in HES-free UW solution at pH 6.2 and pH 7.4. Adenine (5 mM) and ribose (5 mM) were included in the HES-free UW solution, but glucose, lactate, and pyruvate were not used. At the start of each experiment, 3 µM rotenone was added to the cell suspensions to block oxidative phosphorylation. Myocytes were stored aerobically at 6°C, and samples for lactate, ATP, and %rods were taken at time intervals during storage.

At pH 7.4, lactate production begins to plateau by 4 h (Fig. 8A), probably as the result of glycogen depletion (cf. Ref. 14). As this occurs, ATP is also degraded (Fig. 8B), and cells go into contracture (Fig. 8C). At pH 6.2, lactate production does not plateau, and 50% of the ATP is still remaining after 4 h. The best measure of the rate of ATP utilization comes from the initial linear rate of lactate production (Fig. 8A) before a significant number of cells have gone into contracture (Fig. 8C) and before ATP (Fig. 8B) has declined much. This is the period from 0 to 60 min for both pH values. In this period, the ATP loss at pH 6.2 was not significantly different from that seen at pH 7.4 (Fig. 8B, P > .05), and the same was true for PCr loss (22.8 ± 5.3 nmol/mg at pH 7.4, 16.6 ± 6.7 nmol/mg at pH 6.2, N = 3). However, the contributions of ATP loss and PCr loss to ATP utilization are small compared with that estimated from lactate production (Fig. 8A). Figure 8D shows that myocytes stored at pH 6.2 at 6°C had a total rate of ATP utilization approximately one-half that of myocytes at pH 7.4. This decreased rate of ATP utilization impacted cell configuration: after 4 h of storage with rotenone, myocytes at pH 6.2 still retained their rod shape (75% rods) while the majority of cells at pH 7.4 had gone into contracture (39% rods).


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Fig. 8.   Effect of pH on the rate of ATP utilization by cells. Rotenone (3 µM) was added at time 0 to inhibit oxidative phosphorylation to freshly isolated cells incubated without glucose (see MATERIALS AND METHODS) at 6°C. A: the rate of lactate production was slowed by low pH. B: the rate of loss of ATP was not slowed by low pH. C: the loss of rod-shaped cells was slowed by low pH. D: the calculated rate of ATP utilization by cells was slowed by low pH; n = 3 experiments.

Since neither anaerobically nor aerobically stored cells had well-preserved metabolic reserves under the conditions of Fig. 1, the loss of these reserves could have contributed to the loss of function that is seen in the preserved cells compared with the fresh cells (Fig. 5A). Because we found that low pH plus the substrates glucose and pyruvate or lactate were found to conserve metabolic reserves during aerobic storage of cells (Figs. 6 and 7), we sought to determine whether cells with well-preserved metabolic reserves also suffered a functional decline.

Effect of preservation at low pH with additives on myocyte function. Cells were stored under either aerobic or anaerobic conditions in HES-free UW at pH 6.2 containing glucose, pyruvate, adenine, and ribose. After 48 h, cells were rewarmed with delayed Ca addition, as described in MATERIALS AND METHODS. Cells so treated maintained their configuration as well as freshly isolated cells, whether the cells were stored anaerobically or aerobically (Fig 9A). Cells stored aerobically, however, had far more ATP and glycogen at the end of preservation than cells stored anaerobically (Fig. 9, B and C), as found earlier (Figs. 6 and 7). After rewarming and Ca restoration, these differences were again reduced, because cells stored aerobically lost glycogen and ATP, whereas cells stored anaerobically gained both. Even so, at the end of rewarming, cells stored aerobically had significantly higher levels of ATP (P < 0.01), but glycogen of the aerobic cells was depleted during rewarming to a level similar to anaerobic cells.


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Fig. 9.   Comparison of cell morphology and metabolic reserves of cells preserved aerobically or anaerobically for 48 h in HES-free UW solution, pH 6.2, with additives. HES-free UW solution additionally contained 11 mM glucose, 5 mM lactate, 5 mM adenine, and 5 mM ribose. Cells were returned to physiological conditions in KH-HEPES using the delayed Ca protocol. Changes in cell morphology (A), ATP (B), and glycogen (C) were measured as described in MATERIALS AND METHODS. All stored cells showed good morphology but only aerobically stored cells had good metabolic reserves; n = 3 experiments.

Cells stored either way had almost normal contractile function in response to electric field stimulation, but both had a reduced response to isoproterenol (Fig. 10A). However, the %shortening of cells with isoproterenol was significantly greater for cells stored aerobically than for cells stored anaerobically (Fig. 10A). Figure 10B shows the effect of isoproterenol and its effect on the time to 50% rise. Isoproterenol significantly reduced the time to 50% rise of all cells, but more so in the freshly isolated and aerobically stored cells than in cells stored anaerobically (Fig. 10B). Isoproterenol also tended to reduce the time to 50% relengthening, although barely at a level of significance for any condition, and a 48-h storage had no detrimental effect on cell relengthening whether aerobic or anaerobic (Fig. 10C).


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Fig. 10.   Comparison of contractile parameters of cells preserved aerobically or anaerobically for 48 h in HES-free UW solution, pH 6.2, with additives. HES-free UW solution additionally contained 11 mM glucose, 5 mM lactate, 5 mM adenine, and 5 mM ribose. Cells were returned to physiological conditions using the delayed Ca protocol (see MATERIALS AND METHODS). %Shortening (A), time to 50% rise (B), and time to 50% relengthening (C) were calculated from data collected. %Shortening and time to 50% rise were significantly better in aerobically stored cells stimulated with isoproterenol than anaerobically stored cells. Thus cells stored by either protocol showed some depression of function, but aerobically stored cells had significantly better preservation of the isoproterenol response; n = 3 experiments.


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

The conditions used here to emulate aerobic and anaerobic storage of the whole organ during preservation are only an approximation to the conditions that myocytes in the whole organ would experience, in which the whole organ is either continuously perfused or simply cold stored. In the whole organ, other cell types are present, potentially releasing agents that could impact myocyte survival. The extent of hypoxia may be greater in the whole organ than in a pellet of cells that are simply allowed to settle in a tube. Even so, the isolated cells stored in a pellet at 6°C demonstrate some of the characteristics of ischemia that we have previously characterized with this model at 37°C (19), namely a decline in ATP and glycogen levels, but on a time scale reflecting the lower metabolic rate of hypothermia. Likewise, diluted cells allowed to settle in a thin layer as in our aerobic conditions may be slightly hypoxic, and hence less aerobic than cells in a perfused whole heart. Even so, these cells were clearly more aerobic than cells in the anaerobic condition, and, in preliminary experiments, survived better than cells that were agitated to ensure complete oxygenation. The beneficial effect of added substrates on ATP and glycogen levels in cells under the aerobic condition further supports the view that cells under the aerobic condition had free access to the HES-free UW and were not ischemic.

The complete loss of glycogen and ATP from cells stored anaerobically for 24 h (Fig. 1) was expected since glycogen was the only source of ATP under these conditions, but we were surprised that cells stored aerobically in the absence of additional substrates also lost almost all of their glycogen. If all of the glycogen stores (~140 nmol of glucose equivalents per milligram; Fig. 6C) were metabolized aerobically, 5,460 nmol of ATP would be generated, enough for almost 2 days at a rate of 2 nmol of ATP per minute at 6°C (Fig. 8D). However, even in hearts under normal aerobic conditions, most glucose equivalents passing through glycolysis are not oxidized but rather are converted to lactate (41), so most of the glycogen lost in our experiments was presumably converted to lactate. Although HES-free UW solution contains no substrates, cells do have endogenous stores of triglycerides of ~21 nmol/mg, which, on hydrolysis to fatty acids (63 nmol/mg), have the potential to generate 8,852 nmol of ATP by oxidative phosphorylation using a yield of 140.5 nmol of ATP per nanomole of fatty acid (35). A store of this size would potentially last for 3 days under aerobic conditions if it was the sole source of ATP. Thus glycogen was lost even though sufficient substrates were probably available to maintain ATP levels by oxidative metabolism. The heart normally metabolizes both fatty acids and glucose simultaneously. Although the proportion of each substrate made available to the heart does affect the proportion of each utilized, the conditions used here favor glycogen depletion because glycogen was the only source of glucose units for glycolysis.

The view that glycogen was depleted because it was the only source of sugar substrate is supported by the beneficial impact of additional substrates like pyruvate and lactate, which directly compete with pyruvate generated from glycogen for oxidation and which also potentially serve as substrates for glycogen synthesis. Glucose alone, however, was surprisingly ineffective at preventing glycogen depletion. A strong potentiation of glycogen synthesis from glucose by the substrates lactate and pyruvate was observed in dog heart by Laughlin et al. (29). The effect of lactate plus glucose on glycogen content under aerobic conditions can be understood as a diversion of glucose from glycolysis to glycogen synthesis (11). These authors found that lactate caused an inhibition of glycolysis at phosphofructokinase via an increase in citrate and a drop in fructose 2,6-bisphosphate, and lactate also caused an increase in the substrate UDP glucose and the stimulator of glycogen synthase, glucose 6-phosphate. It should be appreciated, however, that from our measurements, we could not distinguish an effect on glycogen synthesis from an effect on reducing the rate of glycogen breakdown, since the observation was a slowing of the rate of loss of total glycogen.

Some insight into the mechanism by which low pH delayed the loss of glycogen was gained from experiments in which the pH dependence of the rate of ATP utilization by the cells was measured (Fig. 8D). These showed a 45% reduction in that rate at 6°C when pH was reduced from 7.4 to 6.2 (Fig. 8D). Note that this reduced rate of ATP utilization cannot be accounted for in terms of the well-known inhibition of glycolysis by low pH (38), since at both pH values, ATP levels were equally well maintained over the time period when the rate of ATP utilization was measured (0-60 min). The reduced rate of lactate production thus reflects a reduced rate of ATP utilization rather than a reduced ability to synthesize ATP at the rate required. Similar considerations allow us to conclude that the rate of lactate efflux from the cells was not rate limiting for glycolysis in the measurements of Fig. 8, even at the low pH. Lactate efflux is expected to be pH sensitive because the lactate transporter carries lactate in the protonated (uncharged acidic) form (43). If intracellular lactate buildup had inhibited the rate of glycolysis, then cell ATP levels would have fallen more than in the pH 7.4 controls, but this was not observed (Fig. 8). A stimulation of lactate influx by low pH, on the other hand, may have enhanced the ability of lactate to inhibit glycogenolysis (Fig. 7C).

The reduced rate of ATP utilization at low pH observed here contrasts with our earlier observation that, in cells in a KH bicarbonate medium with Ca and glucose at 37°C, there is no reduction in the rate of ATP utilization when the pH is reduced from 7.4 to 6.2 (21). Part of the difference between observations could arise from mitochondrial Ca cycling in that study (21), which would not be present to the same degree in the current study, because UW solution has no added Ca, and it even binds Ca to a significant degree (7). The mitochondrial ATPase significantly contributes to the rate of ATP utilization in ischemic rat heart cells (19). The pH effect seen here could arise from an inhibition of mitochondrial ATPase activity by low pH, since this ATPase is inhibited by low pH (37).

Can the rate of ATP utilization measured here be fairly applied to cells stored under aerobic conditions? To measure the rate of ATP utilization, the use of rotenone is required. We have no reason to think that this will result in a false measure of the pH dependence of the rate of ATP utilization. This principle is, therefore, also likely to apply to aerobic cells such as those in Fig. 6, and will, in part, account for the improved preservation of glycogen reserves by low pH seen under these conditions (Fig. 6C).

Under anaerobic conditions, a 45% reduction in the rate of ATP utilization would have a proportional effect on the rate of glycogen utilization. It would also contribute, however, to the reduced glycogen utilization observed under aerobic conditions at low pH (Fig. 6C), even though under these conditions most ATP is provided by oxidative phosphorylation. A second mechanism by which lowered pH may have resulted in a lowered rate of glycogen depletion in aerobic cells is via an inhibition of glycogenolysis and an improvement in the fraction of pyruvate that is oxidized in the mitochondria rather than reduced to lactate. Recent studies in skeletal muscle suggest that glycogenolysis is inhibited by acidosis, first because of reduced phosphorylase activation resulting from reduced production of the phosphorylase activator AMP, and second because of increased pyruvate oxidation relative to lactate production (24).

The relationship between ATP content and cell configuration and metabolism in heart cells is complex. In ischemic cells at 37°C without glucose, the ATP decline is biphasic: an initial decline with no change in cell configuration is associated with upregulation of the glycolytic metabolism of glycogen to lactate, which is needed to supply the demand for ATP (19). This phase is followed, on exhaustion of glycogen, by ATP plummeting to zero, causing cell rigor contracture and termination of glycolysis and high energy phosphate metabolism (5, 19). Cells reoxygenated after such contracture can immediately resynthesize ATP to the extent that AMP and IMP have not yet been degraded, but the cell configurational change is irreversible at this temperature (23). Indeed, reoxygenation further degrades structural integrity as cells change from a square to a round configuration, a change that may be attributed to disruption of intracellular Ca metabolism (23). In the anaerobically stored cells studied here, on the other hand, ATP depletion during cold storage did not result in cell contracture (Fig. 3A), presumably because contracture is inhibited by low pH (Fig. 6A) and by hypothermia. Challenge of these cells with physiological medium that contained Ca caused a configurational change that began even in the cold and became complete when the cells were warmed (Fig. 3A). This configurational change was Ca dependent; it could have been prevented by removal of Ca from the medium (Fig 3A). The increase in ATP seen on reoxygenation, on the other hand, presumably reflected restored oxidative phosphorylation of the remaining nucleotide pool, and this process was independent of Ca (Fig. 3B) or the configurational change (Fig 3A). The relationship between ATP and configuration in Fig. 8 can also be understood in these terms. The initial drop in ATP, which occurs with little change in cell configuration, is analogous to the first phase of ATP decline seen under ischemic conditions at 37°C, where glycolysis upregulates in response to the ATP demand. But what accounts for the progressive loss of rod-shaped cells seen at pH 7.4, which occurs even though ATP levels are comparable with those at pH 6.2? The same question could be asked regarding the data in Fig. 6 for aerobically stored cells. The beneficial effect of low pH on cell configuration is very marked (Fig 6A). The loss of rod-shaped configuration was not accompanied by an ATP decline (Fig 6B). Contracture under these conditions could thus reflect a Ca-dependent contracture rather than a rigor contracture. Cell Ca stores may be released to the cytosol and have difficulty leaving the cell because of the hypothermia. The pH dependence could then reflect the well-known competition between protons and Ca for binding at the myofilaments (44). In summary, hypothermia appears to protect cells from rigor (low ATP) contracture but may leave them susceptible to Ca-dependent contracture; the latter, however, can be prevented by low pH.

The potentiation of intracellular Ca actions at higher pH values could have other detrimental consequences. A major source of cardiac dysfunction resulting from a brief period of ischemia ("stunning") is thought to arise from troponin I degradation that results from the activation of calpain by the ischemia-induced intracellular Ca overload (33). It is possible that the morphology of cells stored at the higher pH in intact tissue may be less damaged by any such Ca than are isolated cells, because cells in the intact tissue are anchored to each other, so they may be less likely to undergo hypercontracture. Even so, the processes described here are likely to happen in the intact organ, even if their consequences are less dramatic. Thus preservation of rod-shaped morphology by isolated cells can only be a positive indicator of preservation quality.

A major finding of the present work is that cells that have been severely depleted of ATP, when protected from rigor contracture by hypothermia, can, under the appropriate rewarming conditions, be restored to a level of function surprisingly close to normal. This was the case for cells stored anaerobically. There may, of course, be long-term consequences of a period of severe ATP depletion that our study does not address. Cells stored aerobically, on the other hand, never underwent glycogen depletion and did not go through such extreme changes in ATP level. In our study, the immediate functional benefits were only subtle, but there may also be long-term benefits. Regardless, the aerobic protocol avoids this major stress and must be preferred for that reason.

These studies show the importance of the protocol used for restoration of the preserved myocardial tissue to physiological conditions, because the protocol used can strongly influence the structural integrity of the cells. This was particularly evident in cells preserved by anaerobic cold storage. Rewarming anaerobically cold-stored cells in the presence of Ca, which most closely emulates clinical practice with donor hearts, resulted in hypercontracture of essentially all cells (Fig. 3A). A great benefit observed was the allowing of these cells to rewarm before Ca was restored (Fig. 4A). This benefit could result from allowing sarcolemmal pH and Na gradients to recover, thus avoiding Ca overload by Na/H and Na/Ca exchange (1, 32), a process that also would require time for ATP resynthesis to drive the Na pump. The susceptibility of the cold-stored cells to Ca could thus be a consequence of the acidosis that develops under the cold storage conditions and the severe deprivation of the cell ATP content. The cells stored aerobically, by contrast, were much more robust in their tolerance of Ca during the rewarming phase (Fig. 4A). If an objective for optimization of preservation conditions is to have cells that tolerate Ca on warm reperfusion, then aerobic preservation conditions are to be preferred over anaerobic.

Even though metabolic reserves were severely depleted during both 24-h anaerobic and aerobic preservation conditions without added substrates (Figs. 3 and 4), the degree of functional recovery of these cells was surprisingly good. We began these experiments with the expectation that the preservation strategies that best preserved metabolic reserves in cells would be the strategies that resulted in the best preservation in cellular function. This expectation has been supported by these studies but not to the extent we expected. Cells stored for 48 h under aerobic conditions that gave the best preservation of morphology, glycogen, and ATP (Fig. 9) showed significantly better contractile function (Fig. 10) than cold-stored cells that lost essentially all of their glycogen and ATP during the cold storage period (Fig. 9). The difference was, however, only evident in cells inotropically stimulated with isoproterenol. What is perhaps more remarkable about these results is how well the function of the anaerobically cold-stored cells was preserved, given the severity of the metabolic deprivation they underwent. These results, rather, challenge the notion that loss of ATP is what must be prevented if function is to be well restored.

The metabolic reserves of cells stored aerobically for 48 h were quite well maintained throughout (Fig. 9): rewarming resulted in significant glycogen loss in these cells, as in fresh cells (Fig. 9C), but these cells had sufficient glycogen reserves after 48 h of aerobic storage to handle this metabolic cost. ATP levels were no different from fresh cells exposed to the same rewarming protocol (Fig. 9B). Even so, these cells still showed depressed function compared with unstored cells (Fig. 10). The decreased contractility of the cells preserved aerobically could result from cell damage independent of metabolic or energetic considerations.

The relatively small functional impact of well-preserved metabolism seen here could be an underestimation of the impact that would be seen in the working heart, because the isolated unloaded myocytes do little work when they shorten. We have previously measured the metabolic cost of beating for isolated rat heart cells, and it is approximately equivalent to that of nonworking whole hearts (20). Nonworking measures of whole heart function can indicate good myocardial preservation when working measures would indicate a significant loss of function (30). It thus could be that the good preservation of metabolic reserves seen under our best aerobic conditions would result in a much greater preservation of function under working conditions than is evident by measures of unloaded shortening. This may indeed be why the functional difference was only detected in cells treated with isoproterenol (Fig. 10A), a condition that can be considered one measure of inotropic reserve.

The loss of response to isoproterenol could be related to alterations in the beta -adrenergic signaling pathway induced by ischemia as well as to a loss of metabolic reserves: after warm ischemia plus reperfusion of guinea pig hearts, a decrease in sensitivity to isoproterenol has been observed that is largely prevented by pyruvate, an effect attributed to the antioxidant effect of pyruvate on free radical-induced injury to adenylate cyclase, as well as to its effect on cytosolic energetics (45).

Another surprising result shown in Fig. 9 is that the cold-stored cells do not appear to exhibit a pH paradox. Their rod-shaped morphology is as well preserved as that of freshly isolated cells, in spite of 48 h of storage under conditions of low pH, followed by restoration of normal pH. The pH paradox is manifested in neonatal rat cardiac myocytes as cell blebbing and death on restoration of normal pH after a period of normothermic metabolic deprivation at low pH (3). This phenomenon was not thought to be caused by excessive influx of extracellular Ca because it also occurred in the absence of Ca in the pH 7.4 recovery medium (3). Critical differences could be that in the studies with the neonatal myocytes, the cells were normothermic during the period of metabolic deprivation and both Na and Ca were present, whereas in our studies with adult cells, the ischemic period was hypothermic and the HES-free UW solution was a low-Na medium without Ca. Thus the pH paradox could be dependent on Ca, which is accumulated during metabolic deprivation, rather than it being a property of neonatal but not adult myocytes.

The aerobic conditions that best preserve myocyte metabolic reserves and contractile function are not necessarily the conditions that best preserve the function of vascular smooth muscle and endothelial cells. Thus insofar as whole heart function is limited by the preservation of vascular function, these conditions may not be the best for the whole heart. This remains to be determined.

In conclusion, these studies show that the morphology, metabolic reserves, and function of adult heart cells can be quite well preserved for at least 48 h during hypothermia if the cells are maintained aerobically in HES-free UW solution at low pH in the presence of the substrates glucose and pyruvate or lactate. Anaerobic cold storage in HES-free UW solution, on the other hand, which emulates the condition currently used for heart preservation, results in the complete loss of cell ATP and glycogen within 24 h. Although these cells show normal baseline function, their inotropic reserves may well be compromised.


    ACKNOWLEDGEMENTS

This work was supported by a grant from the University of Wisconsin Foundation.


    FOOTNOTES

Address for reprint requests and other correspondence: R. A. Haworth, Dept. of Surgery, Univ. of Wisconsin, Clinical Science Center, 600 Highland Ave., Madison WI 53792 (E-mail: haworth{at}surgery.wisc.edu).

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.

Received 17 August 2000; accepted in final form 23 April 2001.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
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Am J Physiol Cell Physiol 281(3):C758-C772
0363-6143/01 $5.00 Copyright © 2001 the American Physiological Society




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