| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
|
|
|
|||||||
|
|||||||||
Division of Cardiology, Internal Medicine, University of Texas-Houston Medical School, Houston, Texas 77030
 |
ABSTRACT |
|---|
 TOP
 ABSTRACT
 INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
|
|---|
We postulate that metabolic conditions that develop
systemically during exercise (high blood lactate and high nonesterified fatty acids) are favorable for energy homeostasis of the heart during
contractile stimulation. We used working rat hearts perfused at
physiological workload and levels of the major energy substrates and
compared the metabolic and contractile responses to an acute low-to-high work transition under resting versus exercising systemic metabolic conditions (low vs. high lactate and nonesterified fatty acids in the perfusate). Glycogen preservation, resulting from better
maintenance of high-energy phosphates, was a consequence of improved
energy homeostasis with high fat and lactate. We explained the result
by tighter coupling between workload and total 
-oxidation. Total
fatty acid oxidation with high fat and lactate reflected increased
availability of exogenous and endogenous fats for respiration, as
evidenced by increased long-chain fatty acyl-CoA esters (LCFA-CoAs) and
by an increased contribution of triglycerides to total -oxidation. Triglyceride turnover (synthesis and degradation) also appeared to
increase. Elevated LCFA-CoAs caused high total -oxidation despite
increased malonyl-CoA. The resulting bottleneck at mitochondrial uptake
of LCFA-CoAs stimulated triglyceride synthesis. Our results suggest the
following. First, both malonyl-CoA and LCFA-CoAs determine total fatty
acid oxidation in heart. Second, concomitant stimulation of peripheral
glycolysis and lipolysis should improve cardiac energy homeostasis
during exercise. We speculate that high lactate contributes to the
salutary effect by bypassing the glycolytic block imposed by fatty
acids, acting as an anaplerotic substrate necessary for high
tricarbocylic acid cycle flux from fatty acid-derived acetyl-CoA.
fatty acids; carnitine palmitoyltransferase I; malonyl-coenzyme A; long-chain fatty acyl-coenzyme A esters
| |
INTRODUCTION |
|---|
|
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
|
|---|
THE PATTERN OF SUBSTRATE USE by the heart in vivo is the combined result of preference, competition between the major energy substrates (fatty acids, lactate, and glucose), and substrate provision by the host and therefore varies according to workload and hormonal status. The heart prefers fatty acids (29, 30), which supply the majority of ATP synthesis at rest and exercise. The exercising metabolic state is characterized by high blood lactate (from skeletal muscle glycolysis; see Refs. 12 and 20) and high nonesterified fatty acids (from adrenergic stimulation of lipolysis in adipose tissue; see Refs. 12, 20, and 26), which compete with one another (4, 8, 33) and with glucose at the heart. Glycolysis and subsequent pyruvate oxidation are inhibited by fatty acids (reviewed in Ref. 28), but the effect on carbohydrate oxidation is partially offset when heart work is increased, because pyruvate dehydrogenase becomes activated by dephosphorylation (18). The end result is that fatty acids and lactate (which bypasses the glycolytic block) become the major respiratory substrates for the heart during exercise (12).
We postulate that the heart is adapted to take advantage of changes in
substrate availability that occur under systemic conditions that
accompany exercise, reflecting increased reliance on either fat or
lactate for respiration. Adaptation should manifest as differences in
the ability to maintain high-energy phosphates and other endogenous
substrates (i.e., glycogen) and/or differences in mechanical function,
depending on whether heart work is stimulated under resting compared
with exercising systemic metabolic conditions. Alternatively, the
availability of preferred substrates under resting conditions (low
systemic fat and lactate) may be inadequate to maintain energy
homeostasis of the heart at high workload. Increased reliance on fat or
lactate should result in tighter coupling between workload on the one
hand and oxidation of one or both of these substrates on the other.
Long-chain fatty acyl-CoA esters (LCFA-CoAs), the proximal substrate
for the rate-limiting step of total -oxidation, serve as an index of
a real increase in the availability of lipids for respiration,
irrespective of the source (exogenous, endogenous, or both).
We included an important improvement over existing studies of
regulation of heart fatty acid oxidation. We found that triglyceride oxidation is increased under systemic conditions that develop during
exercise, becoming a nontrivial component of total fatty acid oxidation
once contractile activity is stimulated. Oxidation of intracellular
triglycerides is regulated, in the first instance, by lipolysis
(17, 26, 32). This process
releases fatty acids into a common pool bound to fatty acid-binding
protein (reviewed in Ref. 37). We postulate that carnitine
palmitoyltransferase I (CPT I), which is the rate-limiting step for
-oxidation, subsequently regulates flux for mitochondrial uptake of
CoA esters derived from the common pool. Therefore, to better evaluate
the regulation of CPT I, we measured total fatty acid oxidation of
exogenous plus endogenous lipids, in addition to a conventional measure for exogenous fatty acid oxidation (3H2O
production from [9,10-3H]oleate). Total -oxidation of
exogenous plus endogenous lipids is probably more meaningful in terms
of regulation of CPT I by malonyl-CoA and/or by LCFA-CoAs.
| |
METHODS |
|---|
|
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
|
|---|
Heart perfusions and perfusion protocol.
Procedures and sources of materials were given previously
(15). Hearts from chow-fed male Sprague-Dawley rats
(300-350 g) were perfused under working conditions and were
subjected to the pulse-chase protocol depicted in Fig.
1. In the working-heart preparation, we cannulated the aorta and left atrium. The heart ejects
perfusate against a fixed afterload determined by the height of the
aortic overflow. The filling pressure for the left atrium is also fixed
by the height above the heart of a fluid reservoir in the oxygenator. A
portion of the venous effluent from the heart, taken directly from the
pulmonary artery, was pumped through a chamber for O2
determination and was then collected. The remaining venous effluent
drips from the apex of the heart and was also collected. The combined
venous effluent was subjected to analysis for radioactive metabolites.
The perfusate during the chase was Krebs-Henseleit buffer containing 5 mM glucose, 40 µU/ml regular insulin, 0.5 or 5.0 mM
sodium-L-lactate, and 0.4 or 1.2 mM sodium oleate prebound
to 3% bovine serum albumin (fatty acid free) and was equilibrated with
95% O2-5% CO2. The free Ca2+
concentration (~1.4 mM) was adjusted by dialyzing against 10 vol of
albumin-free buffer containing 1.5 mM CaCl2 and 0.1 mM EDTA. The left atrial filling pressure was 15 cmH2O, and
the afterload was initially set at 100 cmH2O and raised to
140 cm at the time of adrenergic stimulation. The aortic flow, which is
not acted on metabolically by the heart, was recirculated, and the
coronary flow was not recirculated during the chase period. We studied two perfusion conditions (n = 25 perfusions each)
corresponding to resting and exercising systemic metabolic states. The
resting state was 0.5 mM lactate in the perfusate, starting at 20 min of the protocol, and 0.4 mM oleate, starting at 45 min, with other conditions as described above. These perfusions were exactly as described in a previous study (15). The exercising
metabolic state was 5.0 mM lactate in the perfusate, starting at 20 min of the protocol, and 1.2 mM oleate, starting at 45 min, with other conditions as described above (see Fig. 1).
|
O2) to O2
consumption resulting from oxidation of each substrate by use of the
following formula (all units are µmol · min
1 · g dry wt1):
MO2 = 25.5 (total -oxidation,
acyl units) + 6 (glucose oxidation) + 6 (glycogen
oxidation) + 3 (lactate oxidation) + 0.5 (pyruvate release).
Pyruvate release (measured enzymatically) contributes slightly to
O2 consumption, because pyruvate is more oxidized than its
precursors (glucose, glycogen, or lactate). We assumed 25.5 mol
O2/mol for the stoichiometry of oxidation of the average acyl group (i.e., like oleate). This is valid because 80-90% of total -oxidation is of exogenous oleate (Table 6), and oleate is the
most abundant fatty acyl moiety in heart triglycerides (27). Glycerol is not well oxidized by heart
(21), and the contribution (if any) of
triglyceride-derived glycerol to O2 consumption was
ignored. We verified the underlying assumption of uniform isotopic
enrichment of glycogen and found that the above relation accounts for
O2 consumption quantitatively by substituting exogenous oleate oxidation plus net triglyceride degradation for total
-oxidation (15). In the present study, we report
individual values for triglyceride degradation, oleate oxidation, and
total -oxidation.
Contractile performance and O2 consumption.
Aortic pressure was measured with a Hewlett-Packard dome transducer on
the side arm of the aortic cannula interfaced to a Gould physiological
record (model 2400S; Cleveland, OH). Flow rates were measured from the
filling time for a graduated chamber (aortic flow) or by timed
collection of coronary flow into preweighed vials. Hydraulic power (in
W) is the product of cardiac output (coronary plus aortic flow, in
m3/s) times the afterload (in Pa). To measure
O2 consumption (MO2), a
portion of the coronary flow from the pulmonary artery (10 ml/min) was
pumped through a stirred, thermostatted 3-ml chamber fitted with a
Clark electrode (Yellow Springs Instruments) and then returned to the
heart chamber. A second electrode was fitted to the bottom of the
oxygenator, and the two electrode currents were displayed continuously.
MO2 was calculated from the
arteriovenous concentration difference for O2 times
the coronary flow, with the use of 1.06 mM for the concentration of
dissolved O2 in a solution equilibrated with an atmosphere
of 100% O2 (34). Electrodes were calibrated with air-saturated water (19.6% O2 saturation after
correction for water vapor, 47 mmHg at 37°C). O2
saturation was maintained between 70 and 85% throughout the protocol
(the theoretical maximum is 89%).
Description of the protocol.
Several features of the perfusion protocol (Fig. 1) are in need of
clarification. The first 45 min were designed so that glycogen would be
degraded and then resynthesized, as described previously (15). The first 20 min of perfusion without carbon
substrate (in the presence of O2) cause about one-half of
the preexisting glycogen to be consumed (15) and were
included so that preexisting glycogen would not dilute newly
synthesized glycogen. Hearts were then switched to conditions that
would promote glycogen resynthesis. In one group of hearts, in addition
to nonradioactive glucose, we included [U-14C]glucose so
that glycogen would become radiolabeled (pulse portion of the
protocol). The rationale for including
D--hydroxybutyrate and L-lactate during the
resynthesis phase was to divert exogenous [U-14C]glucose
from glycolysis into glycogen synthesis. Lactate and -hydroxybutyrate are good energy substrates for the heart and replace glucose as the respiratory substrate. Oxidation of
-hydroxybutyrate by the heart is similar to that of short-chain
fatty acids. We used -hydroxybutyrate instead of a more
physiological long-chain fatty acid during the pulse to obviate
technical difficulties associated with adding a long-chain fatty
acid-albumin complex to the perfusate. Starting at 45 min of the
protocol, we replaced -hydroxybutyrate with a long-chain fatty
acid-albumin complex to more accurately model a physiological state,
and we began measuring the metabolic activity of the heart. For the set
of perfusions in which glycogen was labeled
([14C]glycogen group), [U-14C]glucose was
replaced with nonradioactive glucose starting at 45 min of the protocol
("chase" portion of the protocol).
Analytic procedures. 14CO2 and 3H2O were measured in fresh perfusate as described previously (14). The sum for [14C]lactate plus [14C]pyruvate was determined in deproteinized perfusate by paper chromatography (14). Pyruvate release (total, nonradioactive) was measured enzymatically in deproteinized perfusate (3).
Frozen hearts, stored at
70°C, were weighed and ground to a fine
powder under liquid N2, and a portion was taken for
dry-weight determination. Heart dry weights were corrected for trapped
albumin (~15% of the total dry wt) with the use of 0.46 for the
fractional extracellular fluid space volume (previously determined with
[U-14C]sucrose) and 3.6% dry weight of the perfusate.
Procedures for glycogen content and radiochemical enrichment of
glycogen were given previously (15). Other metabolites
were measured in freshly prepared 6% perchloric acid extracts of
heart, adjusted to pH 5 with buffered KOH. Adenine nucleotides,
phosphocreatine, and glucose were measured by use of established
enzymatic methods (3). The calculation for intracellular
glucose, with the use of [U-14C]sucrose as the
extracellular fluid space marker, was given previously (Table 3 in Ref.
15). Pi was measured with the acid molybdate reaction
(9). Perchloric acid-insoluble material was used for the
assay of LCFA-CoAs on the basis of hydrolyzable CoA-SH
(1), measured enzymatically with 
-ketoglutarate
dehydrogenase (2). Malonyl-CoA was measured
radiochemically with purified rat liver fatty acid synthase (1.1 U/mg
protein) and [acetyl-3H]acetyl-CoA. We included a
malonyl-CoA internal standard for each determination as described by
McGarry et al. (24). Triglycerides were analyzed on lipid
extracts of powdered heart (5) after isolation of the
triglyceride fraction by TLC, as described previously (15). Isolated triglycerides were subjected to alkaline
hydrolysis, and the neutralized hydrolysate was analyzed for glycerol
with the use of glycerol kinase (3). Values were referred
to acyl content (glycerol/3). To measure de novo triglyceride
synthesis, the neutralized hydrolysate was taken for scintillation
counting and referred to new acyl group incorporation on the basis of
the specific activity of extracellular [9,10-3H]oleate.
Enzyme assays.
Glycogen synthase was measured as described by Skurat et al.
(31) with the use of the filter-binding assay of Thomas et al. (36). The glycogen phosphorylase assay was based on
the method of Gilboe et al. (13). Phosphorylase
a was measured in the presence of 0.5 mM caffeine to prevent
activation by endogenous AMP (19), and total phosphorylase
was measured in the presence of 3 mM AMP. Pyruvate dehydrogenase was
measured by 14CO2 production from 2 mM
[1-14C]pyruvate, as described by Harris et al.
(16). The active form was measured directly, and the total
activity was measured after incubation for 30 min at 30°C with 1 mM
CaCl2 plus 5 mM MgCl2 to allow
dephosphorylation by endogenous phosphatases. This procedure produced
stable, maximal values. The total activity of pyruvate dehydrogenase
was not different between any of the groups, averaging 23 ± 1 µmol · min1 · g dry wt1
(n = 50).
| |
RESULTS |
|---|
|
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
|
|---|
Contractile performance and O2 consumption.
Figure 2 shows performance of the hearts
for the two metabolic states. Resting systemic metabolic conditions
were defined as hearts perfused with 0.4 mM oleate and 0.5 mM lactate
(low fat and low lactate). Exercising systemic conditions were defined as hearts perfused with 1.2 mM oleate and 5.0 mM lactate (high fat and
high lactate). Figure 2A shows contractile performance (hydraulic power, which is the rate of pressure-volume work), and Fig.
2B shows O2 consumption
(MO2). There was no difference in the
values between the two metabolic states. Values depicted are for hearts
subjected to the full-length protocol ("prolonged stimulation").
For each of the two metabolic states, there were two other treatment
groups (unstimulated and acute stimulation) that were freeze-clamped at
earlier times in the protocol for tissue analysis. Values for
contractile performance and MO2 for the
latter groups overlapped those presented in Fig. 2 and thus were
omitted for clarity. The individual isotope treatment groups
([14C]glucose, [14C]glycogen, and
[14C]lactate) were well matched for contractile
performance and MO2 throughout the
protocol (data not presented).
|
Substrate oxidation.
Figure 3 shows the time course of
substrate oxidation during the chase period in the presence of high fat
and lactate. The time course with low fat and lactate was reported
previously (15). Table 1
gives O2 consumption resulting from oxidation of
each substrate for both conditions. Rates were determined by the Fick principle (arteriovenous concentration difference times
coronary flow, but the "arterial" side was fresh perfusate or
endogenous substrate) on the basis of 14CO2
production for carbohydrate oxidation or
3H2O production from exogenous
[9,10-3H]oleate. There were three prominent effects of
high concentrations of fat and lactate on rates of substrate oxidation.
First, exogenous oleate oxidation, which was not significantly
increased under control conditions, was increased with the work jump in
the presence of high fat and lactate. Second, the increases in
oxidation of glucose and glycogen with the work jump were strongly
inhibited. Third, glucose and glycogen were replaced by lactate as the
respiratory substrate. This replacement, however, was less than
stoichiometric in terms of ATP synthesis (equivalent to O2
consumption) during acute stimulation, because the increase in the
contribution of carbohydrate oxidation to O2 consumption
was less with high fat and lactate [28.7 ± 3.5 vs. 16.2 ± 2.7 (P < 0.05) for acute stimulation and 27.0 ± 2.7 vs. 20.2 ± 2.64 µmol · min1 · g
dry wt1 (not significant) for prolonged stimulation for
low vs. high fat and lactate, respectively; Table 1]. This resulted
because the increase in total -oxidation was more pronounced with
high fat and lactate (see Table 6 and Fatty acid oxidation).
|
|
Energy homeostasis based on conservation of high-energy phosphates.
Because contractile activity and MO2
during adrenergic stimulation (particularly the peak values) did not
differ between the two systemic metabolic states, ATP turnover was not
compromised. We therefore examined more sensitive indexes of energy
homeostasis. The static content of ATP was slightly decreased with
prolonged adrenergic stimulation. The small decreases during acute
stimulation were not significant (Table
2). ATP content is not a sensitive index
of energy homeostasis, nor does the value predict ATP turnover unless
adenine nucleotides are severely depleted. However, small decreases in
ATP are amplified (by adenylate kinase) into larger (percentage wise)
changes in AMP, making AMP an important effector for the signaling of
energetics (i.e., at phosphorylase, phosphofructokinase, and the
AMP-kinase cascade) and a more sensitive index of energy homeostasis.
Phosphocreatine and Pi also seem to be sensitive indexes of
energy homeostasis. The values are shown in Table 2. With low fat and
lactate, phosphocreatine was decreased, and AMP was increased during
acute contractile stimulation. The changes in Pi and
phosphocreatine were persistent (see Prolonged stimulation in Table 2).
In contrast, in hearts stimulated in the presence of high fat and
lactate, the corresponding changes were either attenuated or, as in the
case of phosphocreatine, absent. This indicates that there was improved
homeostasis of high-energy phosphates under exercising systemic
metabolic conditions, which is consistent with our hypothesis that the
heart is better suited to perform high work under these conditions.
|
Energy homeostasis based on preservation of glycogen.
Table 3 gives the glycogen concentration
of hearts in both states. Values for net glycogen degradation (decrease
in value after the work jump) are in parentheses. The glycogen
concentration before stimulation was the same in the two groups. With
low fat and lactate, one-half of glycogen became degraded after
prolonged (20 min) stimulation. The effect of high fat and lactate was
to attenuate net glycogenolysis (57% attenuation acutely and 45% attenuation after prolonged stimulation). We suggest that the difference in glycogen preservation is an important consequence of
improved homeostasis of high-energy phosphates resulting from contractile stimulation in the presence of high fat and lactate. Bear
in mind that net glycogen balance reflects both synthesis and
degradation. De novo glycogen synthesis occurred during net glycogenolysis in this study (see below). Therefore, it is reasonable to ask whether net glycogen sparing resulted from reduced
glycogenolysis, increased glycogen synthesis, or both.
|
Fluxes for glycogenolysis and glycogen synthesis.
Figure 4 shows phosphorylase flux (total
glycolytic [14C]carbon flux from
[14C]glycogen to 14CO2 + [14C]lactate + [14C]pyruvate) in the
two systemic states during the chase period. Total glycolytic flux from
glucose plus glycogen is also shown. Cumulative phosphorylase flux
after the work jump was 54 ± 4 µmol/g dry wt with low fat and
lactate and 38 ± 4 µmol/g dry wt with high fat and lactate
(P < 0.05 vs. control). Oxidation of glycogen (Fig. 3)
was inhibited to a greater extent by high fat and lactate than was
total phosphorylase flux (Fig. 4), the difference being that portion of
glycogen that was metabolized to lactate plus pyruvate. The difference
in total phosphorylase flux between the two systemic metabolic states
(30%) accounted for most, but not all, of net glycogen sparing
attributed to stimulating contractile activity in the presence of
increased fat and lactate. The remaining difference resulted from
increased de novo glycogen synthesis. De novo synthesis, measured by
the incorporation of exogenous [U-14C]glucose into
glycogen during the 20-min period of adrenergic stimulation
([U-14C]glucose isotope treatment group) was stimulated
(75%) by high fat and lactate [4.0 ± 0.7 (n = 5) in controls vs. 7.0 ± 0.7 µmol/g dry wt (n = 5) with high fat and lactate; P < 0.05 vs. control].
|
Regulation of phosphorylase and glycogen synthase.
Potentially important or known features of the regulation of
phosphorylase are shown in Tables 2 and
4. Table 2 shows AMP, which stimulates
phosphorylase allosterically, and Pi, which is a substrate
for phosphorylase. Table 4 shows intracellular glucose, which inhibits
phosphorylase a allosterically, and the activity state of
phosphorylase (percentage of total enzyme that is in the
a-form, or phosphorylated form, resulting from the action of
phosphorylase kinase). The end result (phosphorylase flux) is shown in
Fig. 4.
|
|
Fatty acid oxidation.
Table 6 lists exogenous oleate oxidation
measured by 3H2O production from
[9,10-3H]oleate and total -oxidation. Total
-oxidation was calculated by an independent method with the use of
the values for MO2 and O2
consumption resulting from oxidation of each carbohydrate given in
Table 1 (see METHODS). With low levels of fat and lactate in the perfusate, the increase in exogenous oleate oxidation by the
work jump (19 and 21% during acute and prolonged stimulation, respectively) was not significant, but total -oxidation was
significantly increased (33 and 40% during acute and prolonged
stimulation, respectively). By comparison, exogenous oleate oxidation
and total -oxidation were increased to a greater extent in the
presence of high fat and lactate, especially during acute stimulation
(59 and 79% increases, respectively, compared with 19 and 33%
increases with low fat and lactate). In other words, there was tighter
coupling between workload and -oxidation, both exogenous and total,
in the presence of high fat and lactate. Increased total -oxidation after prolonged adrenergic stimulation was accompanied by a decrease in
malonyl-CoA (Table 6).
|
Triglyceride turnover.
The disparity between oleate oxidation and total -oxidation (little
or no difference with low fat and lactate and a bigger difference with
high fat and lactate) suggested an increase in the contribution of
triglycerides to total -oxidation in the presence of high fat and
lactate. This was born out by direct measurement. Table
7 shows total triglyceride content and de novo synthesis based on incorporation of [9,10-3H]oleate
into the triglyceride fraction. We also calculated fluxes for
triglyceride synthesis and degradation by use of the relationship whereby the change in pool size equals synthesis minus degradation. With low fat and lactate, triglyceride turnover was small (roughly 0.2%/min), and the pool size was unchanged during the chase. Turnover appeared to increase in the presence of high fat and lactate to ~2%/min (it was not possible to calculate degradation before
stimulation from the available data). With high fat and lactate, there
was a fourfold higher de novo synthesis of triglycerides before
and during stimulation of heart work. Degradation of triglycerides also
appeared to increase (10-fold). However, there is considerable error in
the estimates of degradation, because we took a small difference
between two larger values of triglyceride content as part of the
calculation. The change in triglyceride degradation (P < 0.1) did not reach statistical significance. Because glycogen oxidation was largely suppressed (Fig. 3 and Table 1), provision of
high fat and lactate produced a switch from glycogen to triglycerides as the primary endogenous substrate in the high-workload state. Because
of increased triglyceride lipolysis in the presence of high fat and
lactate, the contribution of endogenous lipids to total -oxidation
was increased during stimulation of heart work from 11% with low fat
and lactate to 20% with high fat and lactate.
|
Malonyl-CoA and LCFA-CoA levels.
These levels are given in Table 6. Both were increased in the presence
of high fat and lactate. In general, the increase in total
-oxidation after the work jump accompanied a decrease in the content
of malonyl-CoA. The set point between malonyl-CoA and -oxidation
appeared to be increased in the presence of high fat and lactate.
| |
DISCUSSION |
|---|
|
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
|
|---|
Our objective is to explain the changes in substrate utilization by the heart on stimulation of heart contractile activity, taking into account the increases in systemic lactate and nonesterified fatty acids that accompany exercise. Keep in mind that the usual situation in vivo is an increase in heart work coincident with or preceding changes in substrate milieu induced by exercise. It would be difficult to model the dynamic blood levels of substrates that occur in vivo at the onset of exercise with the use of an isolated heart preparation. Therefore, we decided to perfuse hearts under one of two metabolic states (high fat/high lactate for the exercising state and low fat/low lactate for a resting state) so that the metabolic and contractile response of the heart to an abrupt increase in workload could be compared between these two states.
The levels of lactate and nonesterified fatty acids used in the present study to model the exercising state are quite high and occur only under extreme conditions of exercise in untrained individuals. For example, in the case of untrained (but not trained) running humans, exercise can produce a lactate level of 5 mM by 30 min (20), which is the concentration used in this study. The associated increase in nonesterified fatty acids is slower to develop (20). For untrained (but not trained) rats, running a treadmill produces an increase in systemic lactate to 3.3 mM by 10 min (2). Obviously, we cannot make a direct quantitative comparison of our results to either rats or humans in vivo. The results should, however, predict trends in substrate utilization.
We have to consider the possibility that our protocol (Fig. 1) somehow altered the pattern of contractile or metabolic response of the hearts, because there are no analogous physiological situations that require the heart to work without exogenous carbon substrate but with O2. Energy starvation might be expected to increase production of adenosine (from AMP) or lactate. We have not investigated this issue systematically. However, it seems likely that adenosine, lactate, and any other diffusible metabolites, if accumulated initially, will have washed away by the time hearts were stimulated. This follows from the observation that hearts continued to consume O2 at a high rate during the 20 min of perfusion without exogenous substrate (Fig. 2B). Therefore, hearts did not experience energy deprivation to the same extent as it occurs, for example, during total ischemia. Continued O2 consumption indicates that exogenous substrates were replaced, in large part, by oxidation of endogenous substrates. In addition, hearts were perfused under nonrecirculating conditions with fresh perfusate during the 30 min that we measured metabolic and contractile activity, including the 10 min before adrenergic stimulation. This procedure would have removed accumulated adenosine, lactate, and any other diffusible metabolites by the time hearts were stimulated.
We accounted for four features of the exercising state that seem most relevant to the metabolic and contractile activity of the heart: increased afterload, increased adrenergic stimulation of the heart, high systemic lactate (normally derived from skeletal muscle contraction in vivo), and increased nonesterified fats, which, in vivo, result from adrenergic stimulation of lipolysis in adipose tissue and in the heart itself.
The presence of exercising systemic metabolic conditions produced a
switch from glycogen to triglycerides as the predominant endogenous
substrate, a switch from glucose to lactate as the predominant
exogenous carbohydrate, and tighter coupling between workload and
-oxidation of fatty acids on stimulation of heart work. Increased
heart work in the absence of changes in substrate milieu that
characterize the exercising state (i.e., with low fat and lactate)
produced a transient loss of energy homeostasis. There were small
differences in high-energy phosphates between the two systemic
metabolic states. However, differences in high-energy phosphates
available to support contraction, based on differences in adenine
nucleotides and phosphocreatine, were trivial compared with ATP
turnover or with the ATP equivalent of differences in glycogen content
between the two metabolic states. Of greater importance, a compensatory
response to reduced energy charge is stimulation of phosphorylase flux
by AMP (from ATP hydrolysis), which stimulates phosphorylase
b allosterically, and Pi (from hydrolysis of
phosphocreatine and ATP), which is a substrate for phosphorylase. The
energetic significance of the resulting glycogen depletion could
manifest during repetitive contractile challenge or in terms of the
time for recovery. We suggest that the resulting glycogen depletion is
potentially more important in terms of the tolerance of the heart to a
repetitive challenge than are immediate effects resulting from small,
physiological changes in high-energy phosphates, because contractile
performance was not compromised in the short term. Also, the requisite
glycogen repletion prolongs the time for recovery from exercise. The
loss of homeostasis of high-energy phosphates and resultant glycogen
depletion were obviated when we stimulated hearts in the presence of
high lactate and nonesterified fatty acids. Therefore, the rationale
for an exercise warm-up is improved cardiovascular tolerance resulting
from energetically more-favorable metabolic conditions, because the
warm-up will tend to increase systemic levels of (especially) lactate
and of nonesterified fatty acids to a lesser extent.
One should be cautious in the interpretation of total tissue AMP. The tissue levels of AMP reported in Table 2, measured in perchloric acid extracts of freeze-clamped heart, predict concentrations within the cell (roughly 0.5 mM) that are much too high to regulate phosphorylase. Phosphorylase is activated by submicromolar concentrations of AMP. Cytosolic concentrations of free AMP are, therefore, probably also submicromolar. Bünger and Soboll (6) calculated free cytosolic concentrations of AMP (roughly 100 nM) from mass-action ratios for creatine kinase and myokinase. They suggested that most of cellular AMP is sequestered, possibly within mitochondria (11). A similar problem arises in the interpretation of the whole tissue levels of malonyl-CoA given in Table 6, because the predicted free concentrations are much too large to allow appreciable flux through heart CPT I. Presumably, most of malonyl-CoA, like AMP, is sequestered, either by compartmentalization or by protein binding. We suggest that total tissue levels of the two ligands (AMP and malonyl-CoA) reflect the effective concentration at their respective target enzymes. This statement is true under the condition that the relevant process, such as protein binding, is reversible. Therefore, we suggest that changes in flux for phosphorylase or CPT I can be interpreted on the basis of changes in the whole tissue content of their respective regulatory ligands. The actual free concentration of the ligand at its receptor, however, is not readily calculated from whole tissue levels of AMP or malonyl-CoA.
Regulation of glycogenolysis and glycogen synthesis. Our results suggest that although the well-known stimulation of phosphorylase flux by AMP, and possibly Pi, operates in our heart perfusions after adrenergic stimulation, phosphorylation of phosphorylase to the active or a-form is not an important determinant of the increase in enzyme flux. On the contrary, the expected shift in the portion of phosphorylase to the a-form, reflecting activation of phosphorylase kinase, seems paradoxically to terminate glycogen breakdown. This conclusion is based on the observation that phosphorylase flux ceased despite persistent activation of phosphorylase (see Prolonged stimulation in Table 4). We explain the result by an accumulation of intracellular glucose, which inhibits phosphorylase a allosterically. This effect was more pronounced with high fat and lactate, because there was greater conversion of phosphorylase to the a-form, higher intracellular glucose, and less cumulative phosphorylase flux (Fig. 4). We conclude that phosphorylase flux during the work jump is a true index for homeostasis of high-energy phosphates (particularly AMP and possibly Pi) and not the degree of activation of phosphorylase kinase.
We observed de novo glycogen synthesis during the period of adrenergic stimulation in both groups of hearts (low and high fat and lactate groups). Despite the observation that synthase was activated (phosphorylated) to the same extent in both groups, there was greater glycogen synthesis in the presence of high fat and lactate. We explain the result by greater allosteric stimulation of synthase by glucose-6-P, which increased only in the high-fat, high-lactate group (Table 5). Therefore, both allosteric and covalent mechanisms explained the overall pattern of glycogen synthesis. First, increased phosphorylation of synthase to the active, or glucose-6-P-independent, form after adrenergic stimulation, which occurred to the same extent in the two groups (low and high fat and lactate groups), accounts for the basal rate of de novo glycogen synthesis (4 ± 0.7 µmol/g dry wt observed with low fat and lactate). Superimposed on the basal rate is additional glycogen synthesis resulting from allosteric stimulation by an increase in glucose-6-P, which occurred in the presence of high fat and lactate (to 7 ± 0.7 µmol/g wet wt, P < 0.05). Although synthase activation during adrenergic stimulation might tend to promote futile cycling of glycogen, closer examination reveals that the rationale for this phenomenon may be to hasten the transition of the heart into a recovery phase, and the potential for futile cycling (an energy-consuming process) is actually rather small. The latter statement is based on the observation that the group exhibiting greater de novo glycogen synthesis (high fat and lactate) is also the group exhibiting less phosphorylase flux (Fig. 4). In addition, most of glycogenolysis occurred early on, immediately after adrenergic stimulation (Fig. 4), and yet the activation of synthase by phosphorylation was delayed (compare acute and unstimulated values for synthase activity state in Table 5). The suggestion that synthase activation primes the heart for repletion of glycogen reserves during the subsequent recovery phase was offered previously to explain synthase activation during transient ischemia (25). Activation of glycogen synthase, therefore, may be a response to diverse signals (increased work and ischemia) that initially cause glycogen utilization. Increased fatty acids contributed to improved energy homeostasis. First, a real increase in the availability of fatty acids for respiration, irrespective of whether they were derived from intracellular or extracellular sources, was evidenced by increased content of LCFA-CoAs (the substrate for the rate-limiting enzyme of total-oxidation). Second, the contribution of triglycerides to
total -oxidation was doubled in the presence of high fat and lactate
to 20% during contractile stimulation. Third, we found tighter
coupling of -oxidation to the workload resulting from a larger
increase in total fatty acid oxidation with the work jump in the
presence of high fat and lactate. The increase for fatty acids replaced
some of the increase in carbohydrate oxidation that we observed when
work was stimulated in the presence of low concentrations of fat and
lactate; lactate replaced glucose and glycogen, but the replacement was
less than stoichiometric in terms of ATP synthesis.
We propose that increased lactate serves an important permissive role,
with the prediction that improved energy homeostasis requires increases
in both lactate and fatty acids. By itself, high lactate stimulates
triglyceride turnover in perfused heart (7) and therefore
probably contributed to increased turnover in the present study when
combined with high fatty acids. Of greater importance, lactate bypasses
the glycolytic block imposed by high fatty acid oxidation, becoming
essentially the only oxidizable carbohydrate and a major anaplerotic
substrate required to maintain high Tricarbocyclic acid cycle flux from
fatty acid-derived acetyl-CoA. In turn, high mitochondrial acetyl-CoA
from -oxidation promotes anaplerosis from lactate by stimulating
pyruvate carboxylase (36a). In vivo, systemic lactate, pyruvate, and
alanine probably all provide substrate for pyruvate carboxylase in
heart mitochondria.
As expected, increased total fatty acid oxidation with the work jump
was generally accounted for by decreased levels of malonyl-CoA, reflecting the degree of inhibition by malonyl-CoA of CPT I (reviewed in Ref. 22). Unexpectedly, the set point between malonyl-CoA and total
-oxidation was increased in the presence of high fat and lactate.
Total fatty acid oxidation was high despite increased levels of
malonyl-CoA. We explain this result by increased levels of LCFA-CoAs,
which promote their own oxidation, either by mass action or by
antagonizing the action of malonyl-CoA at CPT I (23). We
propose that the lack of a strict inverse correlation between malonyl-CoA and flux through CPT I reflects modulation of the sensitivity to inhibition of CPT I by malonyl-CoA, resulting from variation in LCFA-CoA levels. Irrespective of whether the bottleneck at
CPT I resulted from increased provision of CoA-esters or inhibition of
CPT I, increased substrate for the acyl-transferase that initiates triglyceride synthesis probably contributed to stimulated lipogenesis.
That total glycolytic flux (Fig. 4) is small compared with oxidative
phosphorylation (Table 1) is not a new observation, but it has an
important consequence with respect to the role of glycolytically
derived ATP. Glycolytic ATP is postulated to be compartmentalized for
ion homeostasis (38, 39). In the presence of
high fat and lactate, we calculate that glycolytic ATP synthesis contributed 1.3 and 5% to total ATP synthesis before and after adrenergic stimulation, respectively. These values are contrasted to
the roughly 25% requirement of basal metabolism, attributable in large
part to Ca2+- and Na+-K+-ATPases.
Therefore, if glycolytic ATP were used entirely for ion homeostasis, it
could only support a small portion of the total requirement for this purpose.
In summary, our results suggest that systemic metabolic conditions that
develop during exercise (increased lactate and nonesterified fatty
acids) are favorable for energy homeostasis of the heart during
contractile stimulation. High fat and lactate suppress glycogen
oxidation and, to a lesser extent, glycogenolysis, which are
compensatory responses to the decrease in energy charge that occurs
when hearts are stimulated in the presence of low levels of fat and
lactate. Improved energy homeostasis resulted from increased
availability of nonesterified fatty acids of both exogenous and
endogenous origin, the latter from increased triglyceride turnover
stimulated by high lactate. We suggest that increased lactate, acting
as the primary anaplerotic substrate, and another oxidizable substrate
are important components of the overall response, because
lactate bypasses the glycolytic block imposed by high fatty acid
oxidation. Increased LCFA-CoAs signal the availability of fatty acids
of exogenous plus endogenous origin for respiration in the presence of
high fat and lactate and promote their own oxidation, either by mass
action or by modulation of the sensitivity of CPT I to inhibition by
malonyl-CoA. Therefore, total -oxidation is regulated, not by
malonyl-CoA alone but by the combined effects of malonyl-CoA and
LCFA-CoAs.
| |
ACKNOWLEDGEMENTS |
|---|
We thank Patrick H. McNulty for providing valuable critique of the manuscript.
| |
FOOTNOTES |
|---|
This work was supported by National Heart, Lung, and Blood Institute Grant RO1-43133 and a Grant-in-Aid from the American Heart Association, Texas Affiliate (97G-329 and 9960099Y).
Address for reprint requests and other correspondence: H. Taegtmeyer, Univ. of Texas-Houston Medical School, 6431 Fannin, Houston, TX 77030 (E-mail: Taegtmeyer{at}uth.tmc.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 19 March 1999; accepted in final form 10 March 2000.
| |
REFERENCES |
|---|
|
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
|
|---|
1.
Awan, MM,
and
Saggerson ED.
Malonyl-CoA metabolism in cardiac myocytes and its relevance to the control of fatty acid oxidation.
Biochem J
295:
61-66,
1993.
2.
Azevedo, JL, Jr,
Linderman JK,
Lehman SL,
and
Brooks GA.
Training decreases muscle glycogen turnover during exercise.
Eur J Appl Physiol Occup Physiol
78:
479-486,
1998[ISI][Medline].
3.
Bergmeyer, HU.
Methods of Enzymatic Analysis. New York: Academic, 1974.
4.
Bielefeld, DR,
Vary TC,
and
Neely JR.
Inhibition of carnitine palmitoyl-CoA transferase activity and fatty acid oxidation by lactate and oxfenicine in cardiac muscle.
J Mol Cell Cardiol
17:
619-625,
1985[ISI][Medline].
5.
Bligh, EG,
and
Dyer WJ.
A rapid method of total lipid extraction and purification.
Can J Biochem Physiol
37:
911-917,
1959.
6.
Bünger, R,
and
Soboll S.
Cytosolic adenylates and adenosine release in perfused working heart.
Eur J Biochem
159:
203-213,
1986[ISI][Medline].
7.
De Groot, MJM,
Willemsen PHM,
Coumans WA,
Van Bilsen M,
and
Van der Vusse GJ.
Lactate induced stimulation of myocardial triacylglycerol turnover.
Biochim Biophys Acta
1006:
111-115,
1989[Medline].
8.
Drake, AJ,
Haines JR,
and
Noble MIM
Preferential uptake of lactate by the normal myocardium in dogs.
Cardiovasc Res
14:
65-72,
1980[ISI][Medline].
9.
Fiske, CH,
and
Subbarow Y.
The colorimetric determination of phosphorus.
J Biol Chem
66:
375-400,
1925
10.
Flaim, SF,
Minteer WJ,
Clark DP,
and
Zelis R.
Cardiovascular response to acute aquatic and treadmill exercise in the untrained rat.
J Appl Physiol
46:
302-308,
1979
11.
Geisbuhler, T,
Altschuld RA,
Trewyn RW,
Ansel AZ,
Lamka K,
and
Brierley GP.
Adenine nucleotide metabolism and compartmentalization in isolated adult rat heart cells.
Circ Res
54:
536-546,
1984
12.
Gertz, EW,
Wisneski JA,
Stanley WC,
and
Neese RA.
Myocardial substrate utilization during exercise in humans.
J Clin Invest
82:
2017-2025,
1988.
13.
Gilboe, DP,
Larson KL,
and
Nuttall FQ.
Radioactive method for the assay of glycogen phosphorylase.
Anal Biochem
47:
20-27,
1972[ISI][Medline].
14.
Goodwin, GW,
Ahmad F,
Doenst T,
and
Taegtmeyer H.
Energy provision from glycogen, glucose, and fatty acids on adrenergic stimulation of heart work.
Am J Physiol Heart Circ Physiol
274:
H1239-H1247,
1998
15.
Goodwin, GW,
Taylor CS,
and
Taegtmeyer H.
Regulation of energy metabolism of the heart during acute increase in heart work.
J Biol Chem
273:
29530-29539,
1998
16.
Harris, RA,
Yamanouchi K,
Roach PJ,
Yen TT,
Dominianni SJ,
and
Stephens TW.
Stabilization of glycogen stores and stimulation of glycogen synthesis in hepatocytes by phenacyl imidazolium compounds.
J Biol Chem
264:
14674-14680,
1989
17.
Heathers, GP,
Al-Muhtaseb N,
and
Brunt RV.
The effect of adrenergic agents on the activities of glycerol 3-phosphate acyltransferase and triglyceride lipase in the isolated perfused rat heart.
J Mol Cell Cardiol
17:
785-796,
1985[ISI][Medline].
18.
Hiraoka, T,
DeBuysere M,
and
Olson MS.
Effect of -agonists on the regulation of pyruvate dehydrogenase in the perfused rat heart.
J Biol Chem
255:
7604-7609,
1980
19.
Hue, L,
Bontemps F,
and
Hers HG.
The effect of glucose and potassium ions on interconversion of the two forms of glycogen phosphorylase and of glycogen synthase in isolated rat liver preparations.
Biochem J
152:
105-114,
1975[ISI][Medline].
20.
Johnson, RH,
Walter JL,
Krebs HA,
and
Williamson DH.
Metabolic fuels during and after exercise in athletes and non-athletes.
Lancet
2:
452-455,
1969[ISI][Medline].
21.
Kreisberg, RR.
Effect of diabetes and starvation on myocardial triglyceride and free fatty acid utilization.
Am J Physiol
210:
379-384,
1966.
22.
McGarry, JD,
and
Brown NF.
The mitochondrial carnitine palmitoyltransferase system. From concept to molecular analysis.
Eur J Biochem
244:
1-14,
1997[ISI][Medline].
23.
McGarry, JD,
Leatherman GF,
and
Foster DW.
Carnitine palmitoyltransferase I. The site of inhibition of hepatic fatty acid oxidation by malonyl-CoA.
J Biol Chem
253:
4128-4136,
1978
24.
McGarry, JD,
Stark MJ,
and
Foster DW.
Hepatic malonyl-CoA levels of fed, fasted and diabetic rats as measured using a simple radioisotopic assay.
J Biol Chem
253:
8291-8293,
1978
25.
McNulty, PH,
and
Luba MC.
Transient ischemia induces regional myocardial glycogen synthase activation and glycogen synthesis in vivo.
Am J Physiol Heart Circ Physiol
268:
H364-H370,
1995
26.
Nilsson, NÖ,
Strålfors P,
Fredrikson G,
and
Belfrage P.
Regulation of adipose tissue lipolysis: effects of noradrenaline and insulin on phosphorylation of hormone-sensitive lipase and on lipolysis in intact rat adipocytes.
FEBS Lett
111:
125-130,
1980[ISI][Medline].
27.
Olsen, RE,
and
Hoeschen RJ.
Utilization of endogenous lipid by the isolated perfused rat heart.
Biochem J
103:
796-801,
1967[ISI][Medline].
28.
Randle, PJ,
Priestman DA,
Mistry SC,
and
Halsall A.
Glucose fatty acid interactions and the regulation of glucose disposal.
J Cell Biochem
55S:
1-11,
1994.
29.
Rothlin, ME,
and
Bing RJ.
Extraction and release of individual free fatty acids by the heart and fat depots.
J Clin Invest
40:
1380-1386,
1961.
30.
Shipp, JC,
Opie LH,
and
Challoner D.
Fatty acid and glucose metabolism in the perfused heart.
Nature
189:
1018-1019,
1961.
31.
Skurat, AV,
Lim S-S,
and
Roach PJ.
Glycogen biogenesis in rat 1 fibroblasts expressing rabbit muscle glycogenin.
Eur J Biochem
245:
147-155,
1997[ISI][Medline].
32.
Small, CA,
Garton AJ,
and
Yeaman SJ.
The presence and role of hormone-sensitive lipase in heart muscle.
Biochem J
258:
67-72,
1989[ISI][Medline].
33.
Spitzer, JJ.
Effect of lactate infusion on canine myocardial free fatty acid metabolism in vivo.
Am J Physiol
226:
213-217,
1974.
34.
Starnes, JW,
Wilson DF,
and
Ereci
ska M.
Substrate dependence of metabolic state and coronary flow in perfused rat heart.
Am J Physiol Heart Circ Physiol
249:
H799-H806,
1985
35.
Staten, MA,
Mathews DE,
Cryer PE,
and
Bier DM.
Physiological increments in epinephrine stimulate metabolic rate in humans.
Am J Physiol Endocrinol Metab
253:
E322-E330,
1987
36.
Thomas, JA,
Schlender KK,
and
Larner J.
A rapid filter paper assay for UDPglucose-glycogen glucosyltransferase, including an improved biosynthesis of UDP-14C-glucose.
Anal Biochem
25:
486-499,
1968[ISI][Medline].
36a.
Utter, MF,
Keech DB,
and
Scrutton MC.
A possible role of acetyl-CoA in the control of gluconeogenesis.
Adv Enzyme Regul
2:
49-68,
1964[ISI][Medline].
37.
Van der Vusse, GJ,
Glatz JFC,
Stam HCG,
and
Reneman RS.
Fatty acid homeostasis in the normoxic and ischemic heart.
Physiol Rev
72:
881-940,
1992
38.
Weiss, J,
and
Hiltbrand B.
Functional compartmentation of glycolytic versus oxidative metabolism in isolated rabbit heart.
J Clin Invest
75:
436-447,
1985.
39.
Weiss, JN,
and
Lamp ST.
Glycolysis preferentially inhibits ATP-sensitive K+ channels in isolated guinea pig cardiac myocytes.
Science
238:
67-69,
1987
This article has been cited by other articles:
|
|
 |
|
  C. J. Zuurbier, F. J. Hoek, J. van Dijk, N. G. Abeling, J. C. M. Meijers, J. H. M. Levels, E. de Jonge, B. A. de Mol, and H. B. Van Wezel Perioperative hyperinsulinaemic normoglycaemic clamp causes hypolipidaemia after coronary artery surgery Br. J. Anaesth., April 1, 2008; 100(4): 442 - 450. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 M. S. Bray, C. A. Shaw, M. W. S. Moore, R. A. P. Garcia, M. M. Zanquetta, D. J. Durgan, W. J. Jeong, J.-Y. Tsai, H. Bugger, D. Zhang, et al. Disruption of the circadian clock within the cardiomyocyte influences myocardial contractile function, metabolism, and gene expression Am J Physiol Heart Circ Physiol, February 1, 2008; 294(2): H1036 - H1047. [Abstract] [Full Text] [PDF]
|
), hearts were chased
with resting levels of lactate (0.5 mM) and oleate (0.4 mM/3%
albumin). For exercising metabolic state (
), hearts
were chased with exercising levels of lactate (5.0 mM) and oleate (1.2 mM/3% albumin). Values are means ± SE for 15 perfusions for each
metabolic state.
,
14CO2 from exogenous
[U-14C]lactate; 
,
14CO2 from [14C]glycogen. Values
are means ± SE for 5 perfusions in each treatment group.
), hearts were chased with resting levels
of lactate (0.5 mM) and oleate (0.4 mM/3% albumin). For exercising
metabolic state (