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(Hypertension. 2004;44:662.)
© 2004 American Heart Association, Inc.

Scientific Contributions

Mechanisms for Increased Glycolysis in the Hypertrophied Rat Heart

Luigino Nascimben; Joanne S. Ingwall; Beverly H. Lorell; Ilka Pinz; Vera Schultz; Keith Tornheim; Rong Tian

From the NMR Laboratory for Physiological Chemistry (L.N., J.S.I., I.P., R.T.), Cardiovascular Division, Department of Medicine, Brigham and Women’s Hospital and Harvard Medical School; the Cardiovascular Division (B.H.L.), Department of Medicine, Beth Israel Deaconess Medical Center and Harvard Medical School; and the Department of Biochemistry and the Diabetes and Metabolism Unit (K.T.), Boston University School of Medicine, Boston, Mass.

Correspondence to Luigino Nascimben, MD, PhD, NMR Laboratory for Physiological Chemistry, Brigham and Women’s Hospital, 221 Longwood Ave, Room 252, Boston, MA 02115. E-mail lnascimben{at}partners.org.bwh.harvard.edu

	Abstract

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Glycolysis increases in hypertrophied hearts but the mechanisms are unknown. We studied the regulation of glycolysis in hearts with pressure-overload LV hypertrophy (LVH), a model that showed marked increases in the rates of glycolysis (by 2-fold) and insulin-independent glucose uptake (by 3-fold). Although the V_max of the key glycolytic enzymes was unchanged in this model, concentrations of free ADP, free AMP, inorganic phosphate (P_i), and fructose-2,6-bisphosphate (F-2,6-P₂), all activators of the rate-limiting enzyme phosphofructokinase (PFK), were increased (up to 10-fold). Concentrations of the inhibitors of PFK, ATP, citrate, and H⁺ were unaltered in LVH. Thus, our findings show that increased glucose entry and activation of the rate-limiting enzyme PFK both contribute to increased flux through the glycolytic pathway in hypertrophied hearts. Moreover, our results also suggest that these changes can be explained by increased intracellular free [ADP] and [AMP], due to decreased energy reserve in LVH, activating the AMP-activated protein kinase cascade. This, in turn, results in enhanced synthesis of F-2,6-P₂ and increased sarcolemma localization of glucose transporters, leading to coordinated increases in glucose transport and activation of PFK.

Key Words: cardiac function • hypertrophy • protein kinases • cardiac metabolism • cyclic AMP

	Introduction

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Glucose utilization is increased in hypertrophied and failing hearts,^1–4 but the underlying mechanisms are poorly understood. Increased glycolytic flux in the hypertrophied myocardium is important because ATP synthesis via glucose utilization may compensate for decreased capacity for ATP synthesis via other pathways.^5,6 In hearts with chronic pressure overload hypertrophy, it was recently reported that chronic depletion of the energy reserve compound PCr coupled with large changes in the ratio of PCr to free creatine led to activation of AMP-activated protein kinase (AMPK) by elevated AMP concentrations.⁷ AMPK acts as a low-on-fuel sensor and, when the cytosolic AMP concentration increases, AMPK activates enzymes in pathways that synthesize ATP and inhibits enzymes in pathways that use ATP.^8,9 Among the many consequences of activated AMPK is increased localization of glucose transporters in the sarcolemma and hence increased glucose uptake by an insulin-independent mechanism.^10,11 In addition, in a study of acute myocardial ischemia, AMPK was found to phosphorylate and thereby activate heart phosphofructokinase-2 (PFK-2), leading to increased production of fructose-2,6-bisphosphate (F-2,6-P₂), a potent activator of the rate-limiting glycolytic enzyme phosphofructokinase (PFK).¹² In the present study, we tested the hypothesis that increased glycolysis in hypertrophied hearts occurs as a consequence of chronic decreases in the energy reserve and activation of AMPK.

Using a model of pressure overload left ventricular hypertrophy (LVH) of the rat heart, in which reduced energy reserve, increased AMPK activity, and increased insulin-independent glucose uptake have all been documented,⁷ we studied the in vivo activation of PFK by determining the cytosolic concentrations of its known activators and inhibitors as well as the glycolytic flux. We also measured the V_max of the major glycolytic enzymes to assess whether the glycolytic capacity increased. Our results suggest that decreased energy reserve in hypertrophied hearts signals an increase in glycolytic flux via 2 coordinated mechanisms: (1) activating the rate limiting enzyme PFK by increasing concentrations of its allosteric activators; and (2) increasing carbon substrate for the glycolytic pathway by increasing glucose transport.

	Materials and Methods

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Animal Model of LVH
Weanling male Wistar rats weighing 50 to 75 g were obtained (Charles River Breeding Laboratories, Wilmington, Del). A titanium clip was placed on the ascending aorta via a thoracic incision as previously described.^13,14 As the animal increases in size, this restriction causes aortic stenosis and pressure overload-induced LV hypertrophy (LVH). Sham-operated animals (control) underwent the same procedure, except that the aortic clip was omitted. Animals were studied 17 to 25 weeks after the surgery. To ensure that all LVH hearts were studied at the stage of similar degree of cardiac hypertrophy and dysfunction, progression of cardiac hypertrophy was monitored using 2-D echocardiography as previously described.^13,14 At the time of the experiments, the body weight was similar (693±33 versus 670±34 g, P=0.64) and the ratio of heart weight to body weight was increased by 50% in LVH compared with control animals (4.3±0.1 versus 2.9±0.2 mg/g, respectively). All experimental procedures were performed according to the guidelines of American Physiological Society and were approved by the institutional animal care and use committee.

Preparation and Characterization of the Isolated Perfused Hearts
Isolated rat hearts were perfused in the Langandorff mode with modified Krebs Henseleit buffer containing 118 mmol/L NaCl, 4.7 mmol/L KCl, 1.75 mmol/L CaCl₂, 1.2 mmol/L MgSO₄, 0.5 mmol/L Na₄EDTA, 25 mmol/L NaHCO₃, 1.2 mmol/L KH₂PO₄, 5 mmol/L pyruvate, and 5 mmol/L glucose. The perfusion buffer was maintained at 37°C, pH 7.4, and was saturated using a gas mixture of 95% O₂ and 5% CO₂. The perfusion flow was titrated to achieve the mean perfusion pressures of 80 and 110 mm Hg for control and LVH, respectively. These pressures were chosen based on previous observations that the coronary perfusion pressure in vivo is higher for LVH than for the control animals. Prior experience showed that this approach would achieve comparable myocardial flow rates per gram of LV weight for the 2 groups.¹⁴ Coronary flow per gram of LV, measured by timed collections of coronary venous effluent, was 13.1±0.7 versus 12.3±0.3 mL/min per gram for LVH and control hearts, respectively (P=n.s.). This strategy was chosen to avoid under-perfusion ischemia as a confounding factor for increased glycolysis in LVH hearts.

LV isovolumic pressure was continuously recorded via a water-filled latex balloon which was inserted in the LV chamber through an incision in the left atria and was connected to a pressure transducer (Viggo-Spectramed P23XL). Hearts were paced at 4.5 Hz using an electric stimulator (Grass Medical Inst). The LV end diastolic pressure was set to 10 mm Hg at the beginning of the experiment by adjusting the volume of the intraventricular balloon. The LV volume, estimated by the volume of the intraventricular balloon, was similar in the 2 groups (0.32±0.03 mL for LVH versus 0.36±0.04 mL for control hearts). Thus, for this stage of LVH and at this low/normal left ventricular end diastolic pressure, we were able to achieve similar compliance and loading conditions. LV function, estimated by the product of heart rate and LV developed pressure (RPP, mm Hg/min), was 29.4±1.1 versus 41.2±2.4 x10³ mm Hg/min for control and LVH hearts, respectively (P<0.05). Because control and LVH hearts have similar LV chamber volumes but different LV masses, we calculated the LV meridional stress using the Laplace formula and assumed that all hearts have a spherical shape.¹³ The estimated systolic wall stress for the isovolumic heart was not different in the 2 groups (88±12 versus 102±9 kdynesxcm^–2, P=n.s.). Based on this result, we concluded that the biochemical measurements in isolated perfused hearts were obtained under conditions of comparable mechanical load and energy demand.

Measurement of Glycolytic Rate and Myocardial Oxygen Consumption
After stabilization, hearts were switched to a recirculating perfusion system containing a total of 100 mL of buffer. The protocol was started by adding [5-³H]glucose (400 000 disintegration/min) to the buffer. An aliquot of buffer (0.5 mL) was sampled every 5 minutes.³ H₂O was separated from [5-³H]glucose by filtering the buffer sample through a column containing an anion exchange resin (Dowex 1x4–400) pretreated with 0.4-mol/L potassium tetraborate.² Glycolytic rate was calculated from the amount of [³H₂O] accumulating in the recirculating buffer with time, and normalized by the dry weight of the heart. In the same hearts, myocardial oxygen consumption (MVO₂) was calculated from the difference in O₂ content in perfusion buffer and effluent from the coronary sinus measured by a Clarke-type O₂ electrode (Orion Research Inc) and normalized to heart weight and multiplied by coronary flow.

³¹P NMR Measurements
The isolated perfused heart was placed in a 20-mm glass tube and was positioned in the center of a 9.4 Tesla superconducting magnet (Oxford Instruments) as previously described.¹⁵ Baseline spectra were obtained over 20 minutes by signal averaging 520 free induction decays, which were used for quantifying ATP, PCr, and P_i contents and intracellular pH.¹⁵

After collecting baseline spectra, the heart was switched to a buffer containing 5-mmol/L 2-deoxyglucose (2DG) in place of glucose for the measurement of glucose uptake as previously reported.⁷ ATP production was maintained in these hearts by supplying pyruvate (5-mmol/L) in the perfusate throughout the experiment.¹⁶ During 2DG perfusion, 10 ³¹P NMR spectra, each obtained by signal averaging 52 free induction decays (total acquisition time of 2 minutes for each spectrum), were consecutively collected and the rate of time-dependent accumulation of 2-deoxyglucose phosphate (2DGP) was determined.

Biochemical Assays
Hearts used for the ³¹P NMR protocols were freeze-clamped and stored at -80°C. Enzyme activities and metabolite concentrations were measured in specimens of these hearts. Total creatine (the sum of Cr and PCr),¹⁷ F-2,6-P₂,¹⁸ and citrate¹⁹ concentrations were determined as previously described. Enzyme assays for the activities of the glycolytic enzymes PFK,²⁰ glyceraldehyde-3-phosphate dehydrogenase (GAPDH),¹⁹ lactate dehydrogenase (LDH),²¹ and the mitochondrial enzyme citrate synthase (CS)²² were performed as previously described.

Data Analysis and Statistics
The myocardial ATP content determined by high-performance liquid chromatography (HPLC) was 26.1±1.4 and 23.8±1.2 nmol/mg protein for control (n=4) and LVH hearts (n=5). These values were used to calibrate [ß-P]ATP peak area of the baseline spectra and to calculate [ATP] assuming an intracellular water content of 0.48 µL per mg blotted tissue²³ and a protein content of 0.18 mg protein per mg blotted wet tissue. [PCr], [P_i], and [2DGP] for both control and LVH hearts were calculated by comparing their peak areas to that of the ATP in the same heart. Intracellular pH was calculated by comparing the chemical shift between the P_i and PCr peaks in each spectrum to values from a standard curve.

Cytosolic free [ADP] was calculated using the creatine kinase reaction equilibrium expression:

[ATP], [PCr], and pH were measured by ³¹P NMR spectroscopy. Free Cr was calculated from the difference between [total creatine] measured in heart homogenates and [PCr] determined using ³¹P NMR spectroscopy. The equilibrium constant calculated for a free Mg concentration of 0.86 mmol/L was taken from reference.²⁴

Cytosolic free [AMP] was calculated using the adenylate kinase (AK) reaction equilibrium expression:²⁵

Data are presented as mean±SE. Two-factor repeated measures ANOVA was used to compare the slopes for the time-dependent accumulation of 2DGP in the 2 groups. Unpaired 2-tailed Student t test was used to compare the rest of the measurements between the 2 groups. A probability value <0.05 was considered significant.

	Results

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Glycolytic Flux
To confirm that glycolysis was increased in this model of LVH, we measured the glycolytic rate. The insulin-independent glycolytic rate was 0.82±0.07-µmol glucose/min per gram dry weight in LVH and 0.41±0.05-µmol glucose/min per gram dry weight in control hearts (P<0.05). MVO₂ was 29.8±2.4 versus 28.8±1.2-µmol O₂/min per gram dry weight in LVH and control hearts, respectively (P=n.s.). Thus, the glycolytic rate was 2-fold higher in LVH hearts compared with control hearts despite similar rates of ATP synthesis estimated from MVO₂.

Glucose Entry
We assessed glucose uptake by measuring the rate of 2DG uptake in isolated perfused hearts using ³¹P NMR spectroscopy. Figure 1A shows 2 sets of ³¹P NMR spectra, one obtained from a control heart (left panel) and the other from a LVH heart (right panel) during 20 minutes of perfusion with 2DG. The accumulation of 2DGP appears as a new resonance peak at 7.05 ppm downfield from the PCr resonance. The 2DGP peak appears earlier and accumulates faster in the LVH heart compared with the control heart. Figure 1B shows the linear fits for the time-dependent accumulation of 2DGP obtained from 5 control and 5 LVH hearts. The equation is y=0.34x–0.49 (r=0.95) for control and y=1.31x–0.66 (r=0.99) for LVH hearts where x is time (min) and y is 2DGP concentration (mmol/L). The slope of the fitted line, which estimates the rate of 2DG uptake, is 3.8 times higher in LVH compared with control hearts (P<0.05).

View larger version (28K): [in this window] [in a new window]   Figure 1. A, Representative stacked plots of 31P NMR spectra of a control heart (left panel) and a LVH heart (right panel) perfused with 2-deoxyglucose (2DG). The major resonances are, labeled from left to right, 2-deoxyglucose phosphate (2DGP), phosphocreatine (PCr), and the -, -, and ß-phosphates of ATP. Each stack consists of 10 spectra, each signal averaged for 2 minutes, obtained from the beginning of 2DG perfusion. See Methods for details. B, Plot of the time-dependent accumulation of 2DGP for control () and LVH hearts (•) perfused with 2-deoxyglucose. Values are means±SE of 2DGP concentration (mmol/L) collected consecutively every 2 minutes from the beginning of 2DG perfusion. Data points were fitted using linear regression analysis. Control: Y(mmol/L)=0.34x(min)–0.49 (r=0.95); LVH: Y(mmol/L)=1.31x(min)–0.66 (r=0.99).

Activities of the Key Enzymes
Activities (V_max) of 3 major glycolytic enzymes PFK, GAPDH, and LDH, as well as the mitochondrial enzyme CS, and AMPK measured in tissue homogenates of LVH and control hearts are presented in Table 1. In spite of higher glycolytic flux observed for LVH hearts, the V_max of the glycolytic enzymes in LVH were not different compared with control hearts, suggesting that increased glycolysis in this model of LVH was not because of increased capacity of the glycolytic pathway. CS, a measure of mitochondrial mass, was also not different (0.86±0.03 versus 0.80±0.04 IU/mg protein for control and LVH hearts respectively, P=n.s), in accord with unchanged MVO₂.

View this table: [in this window] [in a new window]   TABLE 1. Glycolytic Rate and Enzyme Activities

Concentrations of Activators and Inhibitors of PFK
Figure 2 shows typical baseline ³¹P NMR spectra for a LVH and a control heart. Spectra such as these were used to measure the intracellular concentrations of ATP, PCr, P_i, and intracellular pH and to calculate the free concentrations of ADP and AMP. Because the peak areas are proportionate to the amounts but not the concentrations of ATP, PCr, and P_i in the heart, greater ATP peaks in the LVH heart reflect increased myocardial mass. [ATP], however, was not different in LVH compared with the controls (Table 2). [PCr] was markedly decreased (14.5±0.9 versus 24.1±1.6 mmol/L, P<0.001), whereas the total Cr content was reduced only slightly (23.4±0.9 versus 27.8±1.0 mmol/L, P<0.002) in LVH hearts compared with controls. These changes led to a 4-fold increase in the ratio of free Cr to PCr. Intracellular free [ADP], [AMP], and [P_i], all activators of PFK activity, were substantially increased (Table 2).

View larger version (16K): [in this window] [in a new window]   Figure 2. 31P NMR spectra of a control heart (left) and a LVH heart (right) during baseline perfusion with Krebs Henseleit buffer. The major resonances are assigned as (from left to right) extracellular inorganic phosphate (Po), intracellular inorganic phosphate (Pi), phosphocreatine (PCr), and -, -, and ß- phosphates of ATP. The spectrum for the LVH heart shows lower PCr/ATP ratio and increased contents of Pi and phosphomonoester compounds than in the control heart.

View this table: [in this window] [in a new window]   TABLE 2. Concentrations of Activators and Inhibitors of PFK

In a previous study, we found that a similar increase in [AMP] in LVH resulted in activation of AMPK.⁷ Because AMPK has been shown to phosphorylate and activate PFK-2, leading to increased synthesis of F-2,6-P₂ in ischemic hearts, we tested whether [F-2,6-P₂] would be increased in LVH hearts. Using the same LVH hearts in which increased AMPK activity was observed, we found that F-2,6-P₂ content in the LV tissue was increased by 6-fold (Table 2). Furthermore, changes in [F-2,6-P₂] closely correlated to both 1- and 2-AMPK activities in these hearts (R²=0.6 and R²=0.71, respectively). Thus, decreased energy reserve in LVH resulted in marked increases (up to 10-fold) in the concentrations of the known PFK activators ADP, AMP, P_i, and F-2,6-P₂, whereas the concentrations of the inhibitors of PFK, namely ATP, H⁺, and citrate, were all similar in LVH and control hearts.

	Discussion

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The major findings of this study are that, in the presence of unchanged capacity of the glycolytic pathway, increases in glucose entry and activation of the rate-limiting enzyme PFK contribute to increased flux through the glycolytic pathway in hypertrophied hearts. These findings are unlikely a result of differences in workload conditions used in this study for 2 reasons. First, we have matched LV wall stress and MVO₂ in the 2 groups. Furthermore, previous studies have shown that increases of glycolytic rates in hypertrophied hearts are relative insensitive to workload, ie, glycolytic rate is higher at low workload and does not increase further at higher workload.^2–3 In fact, compared with controls, hypertrophied hearts have lower lipid oxidation rates and have increased rates of glycolysis at baseline workload. Increasing workload requires an increase in the rate of ATP production sustained by increasing lipid oxidation in control hearts and by increasing glucose oxidation without further increases in glycolysis in hypertrophied hearts.^2–3 Instead, our results suggest that these changes can be explained by increased intracellular free [ADP] and [AMP], due to decreased energy reserve of the hypertrophied heart, that in turn triggers the AMP-activated protein kinase (AMPK) cascade resulting in coordinate increases in glucose transport and activation of PFK. It has been well documented that compensated cardiac hypertrophy is associated with decreased content of the energy reserve compound PCr; [ATP] falls only in failing heart.^7,16,26 This is illustrated in the ³¹P NMR spectrum as decreased PCr/ATP in the LVH heart (Figure 2). We have previously found an inverse relationship between PCr/ATP and the rate of glucose uptake in hypertrophied hearts, suggesting a role of myocardial energy status in the regulation of substrate metabolism.²⁷ Results of the present study suggest that the signal of decreased energy reserve can be transmitted by altered intracellular free concentrations of ADP and AMP, leading to changes in the metabolic pathways for substrate utilization.

Figure 3 illustrates the proposed mechanisms by which myocardial energetics regulate glucose utilization in LVH. First, a decreased energy reserve can directly activate PFK. Decreased [PCr] with only minimal fall in total [creatine] indicates that there are significant increases in intracellular free [ADP] and [AMP], calculated via the creatine kinase and adenylate kinase equilibrium expressions. Because [ADP] and [AMP] are in µmol/L range, whereas [ATP] is in mmol/L range, large increases in [ADP] and [AMP] can occur when [PCr] decreases with little or no change in [ATP]. This is the case in LVH hearts studied here in which [ADP] increased 4-fold and [AMP] more than 10-fold. It has been shown in in vitro studies that ADP and AMP allosterically activate PFK, but at higher concentrations of ADP and AMP than calculated here for the intact heart.^28,29 Importantly, a study modeling PFK regulation in PCr-containing tissues found that perturbations of [ATP], [ADP], and [P_i] in the physiological range can be regulatory.³⁰ Concomitant increases in glycolytic flux and elevations in free [ADP], [AMP], and [Pi] in LVH hearts raises the possibility that changes in free [ADP], [AMP], and [Pi] found in this study are sufficient to cause allosteric activation of PFK in vivo.

View larger version (43K): [in this window] [in a new window]   Figure 3. Proposed mechanism(s) for increased glycolysis in LVH. Decreased energy reserve in hypertrophied hearts signals an increase in glycolytic flux via 2 coordinated mechanisms in which AMPK plays a central role: (1) increasing carbon substrate for the glycolytic pathway by increasing glucose transport; and (2) activating the rate limiting enzyme PFK by increasing concentrations of its allosteric activators. See text for further details.

Second, by stimulating AMPK, decreased energy reserve also leads to increased glucose entry with an increase of G-6-P and hence F-6-P, the substrate of PFK, and increased [F-2,6-P₂], which allosterically activates PFK. AMPK is a sensor of cellular energy status because it is highly sensitive to [AMP]. Increased [AMP] leads to activation of AMPK via both allosteric and phosphorylation mechanisms.⁹ Once activated, AMPK functions to restore energy homeostasis by modulating multiple metabolic pathways.⁹ During acute metabolic stresses such as ischemia when substantial loss of ATP occurs, increases in AMPK activity stimulate glucose uptake and glycolysis by increasing translocation of GLUT-4 and increasing the synthesis of F-2,6-P₂, a potent stimulant of PFK, in cardiac muscle.¹² We recently reported that AMPK activity was increased in LVH hearts with chronic depletion of energy reserve.⁷ In this model, increased AMPK activity was associated with increased sarcolemma localization of glucose transporters under nonischemic conditions.⁷ Here, we report a marked elevation of [F-2,6-P₂] in the same LVH hearts and a close relationship between [F-2,6-P₂] and the AMPK activity. These results suggest that the signal of chronic depletion of energy storage, namely increased cytosolic [AMP], is transduced via the AMPK system, resulting in alterations of substrate (glucose) utilization.

Previous investigators studying hypertrophied and failing hearts have suggested that the increased glycolytic flux may be in part because of an increased V_max of several glycolytic enzymes.^4,31 In contrast to those observations, we found that PFK, GAPDH, and LDH activities were unchanged despite an increase in glycolytic flux. These differences are likely due to differences in models and stages of hypertrophy studied. For example, distinct from many previous studies, LVH induced by ascending aortic constriction, as used in this study, is not associated with hypertension. Moreover, it is important to recognize that V_max represents the capacity of the enzyme, not the actual reaction velocity. For most biochemical reactions, in vivo velocity is much less than the enzyme capacity or V_max. In the present study, the assayed V_max for PFK was roughly 200 times the measured glycolytic rate. Despite a 2-fold increase in glycolytic rate in LVH, flux through the glycolytic pathway was 1% of the V_max for the key enzymes, suggesting that an increase in the capacity of the enzymes was not necessary for increased glucose utilization in hearts with pressure overload hypertrophy.

Perspectives
Our results suggest that the primary mechanisms underlying increased glycolytic rate in pressure overload hypertrophied hearts are increased carbon substrate due to an increased glucose entry and activation of the rate-limiting enzyme PFK in the pathway. PFK is activated by increased [ADP], [AMP], [P_i], and [F-2,6-P₂] with unchanged concentrations of the inhibitors [ATP], [citrate], or [H⁺]; there are no changes in enzyme capacity. Mechanisms proposed here for the regulation of glucose utilization in hypertrophied hearts reflect a novel link between myocardial energy status and substrate metabolism in a chronic disease model. These observations support the roles for AMP as a molecular signal and for AMPK as a transducer coupling chronic energy depletion and alterations in substrate utilization.

	Acknowledgments

 
This work was supported by National Institutes of Health grants HL67970, HL59246, and AG000837 (R.T.); HL52864 (B.H.L.); HL52320 and HL63985 (J.S.I.); and Juvenile Diabetes Research Foundation grants 1–2002 to 372 and center grants 4–2002 to 456 (K.T.). L.N. was supported by a fellowship award from Merck Sharp & Dohme Italy Spa.

Received April 29, 2004; first decision May 11, 2004; accepted July 29, 2004.

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