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J. Biol. Chem., Vol. 276, Issue 26, 23858-23866, June 29, 2001
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,,¶
From the Department of Diabetes, University of
Newcastle upon Tyne, The Medical School, Newcastle upon Tyne NE2 4HH,
United Kingdom, and § Departament de Bioquímica i
Biologia Molecular, Universitat de Barcelona, Martí i
Franquès, 1, 08028 Barcelona, Spain
Received for publication, February 15, 2001, and in revised form, April 2, 2001
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ABSTRACT |
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 TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
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We used metabolic control analysis to determine
the flux control coefficient of phosphorylase on glycogen synthesis in
hepatocytes by titration with a specific phosphorylase inhibitor
(CP-91149) or by expression of muscle phosphorylase using recombinant
adenovirus. The muscle isoform was used because it is catalytically
active in the b-state. CP-91149 inactivated phosphorylase
with sequential activation of glycogen synthase. It increased glycogen
synthesis by 7-fold at 5 mM glucose and by 2-fold at 20 mM glucose with a decrease in the concentration of
glucose causing half-maximal rate (S0.5) from 26 to 19 mM. Muscle phosphorylase was expressed in hepatocytes
mainly in the b-state. Low levels of phosphorylase expression inhibited glycogen synthesis by 50%, with little further inhibition at higher enzyme expression, and caused inactivation of
glycogen synthase that was reversed by CP-91149. At endogenous activity, phosphorylase has a very high (greater than unity) negative control coefficient on glycogen synthesis, regardless of whether it is determined by enzyme inactivation or overexpression. This high
control is attenuated by glucokinase overexpression, indicating dependence on other enzymes with high control. The high control coefficient of phosphorylase on glycogen synthesis affirms that phosphorylase is a strong candidate target for controlling
hyperglycemia in type 2 diabetes in both the absorptive and
postabsorptive states.
The liver maintains blood glucose homeostasis by
uptake of glucose in the absorptive state, which is converted to
glycogen and triacylglycerol, and by production of glucose from
glycogenolysis and gluconeogenesis in the postabsorptive state.
Glycogen phosphorylase catalyzes the first step in glycogen degradation
(1). It is regulated by allosteric mechanisms and by
phosphorylation of Ser-14 by phosphorylase kinase. The
dephosphorylated form (phosphorylase b) is less
active than the phosphorylated form (phosphorylase a) (1).
Hormones that raise cAMP or cytoplasmic Ca2+ favor the
formation of phosphorylase a, whereas insulin and leptin have the converse effect (1, 2). Phosphorylase is a dimer and exists in
two conformations, an inactive T-state (tight) and an active R-state
(relaxed). The R-state is favored by substrate, phosphorylation, and
allosteric activators (AMP), whereas the T-state is favored by
dephosphorylation and by inhibitors (glucose, glucose
6-P,1 and caffeine) (3, 4).
The liver isoform, unlike the muscle isoform, is more tightly
controlled by phosphorylation than by allosteric regulation (1).
Two sets of evidence suggest that liver phosphorylase is a candidate
pharmacological target for controlling hyperglycemia in diabetes.
Firstly, a potent inhibitor of liver phosphorylase a that
acts synergistically with glucose lowers blood glucose in the
leptin-deficient ob/ob mouse (5). Secondly, the activity of
phosphorylase is elevated in the leptin receptor-defective db/db mouse
(6) and Zucker fa/fa rat (7), which are widely used as animal models
for human type 2 diabetes and insulin resistance.
Recent studies have applied metabolic control analysis (8) to determine
how control of glycogen synthesis is shared between diverse sites. In
skeletal muscle, a high degree of control resides at the glucose
transport/phosphorylation sites (9-11), whereas in liver, a high
degree of control is exerted by glucokinase in conjunction with its
regulatory protein (12-14). Schulz (11) developed a minimal model for
glycogen synthesis in muscle that demonstrates that as glycogenolysis
increases, the distribution of control shifts to the terminal enzymes
in the glycogen synthesis pathway. Mathematical models enable
identification of conditions that alter the distribution of control but
do not allow a quantitative estimate of the degree of control exerted
by specific sites. There has been no experimental analysis of the
control exerted by phosphorylase on glycogen synthesis. In this
study, we used a specific inhibitor of phosphorylase (5) and titrated
phosphorylase overexpression by adenovirus-mediated gene transfer
of the muscle isoform of phosphorylase (15) to determine the flux
control coefficient of phosphorylase on glycogen synthesis in
hepatocytes. Muscle phosphorylase b, unlike liver
phosphorylase b (16), is very sensitive to activation by
AMP. We took advantage of this property of muscle phosphorylase to
increase phosphorylase catalytic activity in hepatocytes independently
of the phosphorylation state of the hepatocyte.
Materials--
CP-91149 (5) was a kind gift from Pfizer
Global Research & Development, Groton Laboratories. Rabbit muscle
phosphorylase b was from Sigma. Sources of other reagents
were as described previously (2, 7).
Hepatocyte Culture and Enzyme Expression--
Hepatocytes were
isolated from male Wistar rats (B & K, Hull, United Kingdom; body
weight, 240-340 g) by collagenase perfusion of the liver (12). The
hepatocytes were suspended in minimal essential medium containing 7%
newborn calf serum and seeded in multiwell plates (12). After cell
attachment (2-3 h), they were incubated for 2 h in serum-free
minimal essential medium containing recombinant adenovirus encoding
either muscle glycogen phosphorylase (AdCMV-MGP (15)) or liver
glucokinase (AdCMV-GK (17)). The medium was then replaced by minimal
essential medium containing 10 nM dexamethasone and 5 mM glucose, and the cells were cultured for 16-18 h
(12).
Metabolic Studies--
For determination of glycogen synthesis,
hepatocytes were incubated in minimal essential medium containing
[U-14C]glucose (2 µCi/ml) and the glucose
concentrations indicated without or with insulin (10 nM)
for 3 h. Parallel incubations without radiolabel were performed
for enzyme activity or metabolite determination. Incorporation of
14C label into glycogen was determined by ethanol
precipitation (18), and rates of glycogen synthesis are expressed as
nmol glucose incorporated/3 h per mg cell protein. The glucose
concentration that causes a half-maximal rate was determined from Hill
plots, using Fig.P-Biosoft Software. Glycogen was determined
analytically using amyloglucosidase (18) and is expressed as nmol
glucosyl residues/mg protein. Glucose 6-P, ATP, and protein were
determined as described previously (19). Glycolysis was determined from detritiation of [3-3H]glucose (19) and incorporation of
[U-14C]glucose into triacylglycerol as described in
Ref. 20.
Enzyme Activity Determination--
For determination of glycogen
synthase and phosphorylase, hepatocyte monolayers were extracted as
described previously (7). Active glycogen synthase and total glycogen
synthase were determined in the absence or presence of glucose 6-P,
respectively (21). Active synthase is expressed as m-units/mg cell
protein or as the activity ratio ( Determination of Control Coefficients--
The control
coefficient (C
All results are expressed as the means ± S.E. for the number of
cell preparations indicated. Statistical analysis was performed using
Student's paired t test.
CP-91149 Causes Inactivation of Phosphorylase and Sequential
Activation of Synthase and Increases the Affinity of Glycogen Synthesis
for Glucose--
We used a potent inhibitor of liver phosphorylase
a,
[R-(R,S)-5-chloro-N-[3-(dimethylamino)-2-hydroxy-3-oxo-1-(phenylmethyl)propyl]-1H-indole-2-carboxamide (CP-91149) (5), to determine the relation between phosphorylase activity and glycogen synthesis. Incubation of hepatocytes with 2.5 µM CP-91149 caused inactivation of phosphorylase
a (Fig. 1A) and
activation of glycogen synthase in the absence of glucose 6-P (Fig.
1B). The former (but not the latter) was significant after 5 min, indicating that activation of synthase is delayed relative to
inactivation of phosphorylase. There was no change in the total
activity of glycogen synthase assayed in the presence of glucose 6-P;
accordingly, changes in the active form of synthase (Fig.
1B) were associated with similar changes in the activity ratio (data not shown). When the cells were preincubated with 2.5 µM CP-91149 for 1 h and then incubated in medium
without inhibitor for 3 h, the recovery of phosphorylase activity
relative to that in untreated controls was 20% (data not shown),
indicating that reactivation of phosphorylase is slow after removal of
inhibitor.
Fig. 2 shows the activity of
phosphorylase and the rate of glycogen synthesis in relation to varying
glucose concentration in the absence or presence of 2.5 µM CP-91149. Glucose caused a
concentration-dependent inactivation of phosphorylase, and
the inactivation by CP-91149 was additive with the effect of glucose (Fig. 2A). The rate of glycogen synthesis was a sigmoidal
function with respect to glucose concentration (Fig. 2B),
with a half-maximal rate obtained at a concentration of glucose
(S0.5) of 25.6 ± 1.1 mM and a Hill
coefficient (h) of 2.8 in the absence of inhibitor (Fig.
2B). CP-91149 (2.5 µM) caused a 7-fold
increase in glycogen synthesis at 5 mM glucose and a 2-fold
increase at 20-25 mM glucose with a leftward shift of the
glucose saturation curve and a decrease (p < 0.03) in
S0.5 for glucose to 18.9 ± 2.1 mM
(h 2.2).
The Flux Control Coefficient of Phosphorylase on Glycogen Synthesis
Determined by Inhibitor Titration--
The flux control coefficient
(C Muscle Phosphorylase Is Expressed in Hepatocytes in the b Form but
Is Activated by Glucagon--
To determine the control coefficient of
phosphorylase by enzyme overexpression, we expressed muscle
phosphorylase in hepatocytes using recombinant adenovirus. We used the
muscle isoform because it is active at physiological concentrations of
AMP and ATP, unlike the liver isoform. The activity of phosphorylase
a in control hepatocytes was 10% of total activity and was
increased 2.6-fold by glucagon (Table I).
Hepatocytes treated with a titer of AdCMV-MGP (50 µl/ml) that
resulted in a 70% increase in total phosphorylase had a similar
activity of phosphorylase a as untreated hepatocytes (Table
I). However, after incubation with glucagon, phosphorylase a
activity was higher in AdCMV-MGP-treated cells than in untreated cells
(29.3 versus 20.9 m-units/mg). This difference in activity is 17% of total muscle phosphorylase (50 m-units/mg) and is similar to
the increase in phosphorylase a caused by glucagon in
untreated hepatocytes, which is also 17% of total liver phosphorylase.
This indicates that muscle phosphorylase in AdCMV-MGP-treated
hepatocytes is expressed mainly in the b form, but it
is activated by glucagon to the same extent as endogenous liver
phosphorylase.
To account for the difference in sensitivity to AMP between liver and
muscle isoforms, two additional assays were used designated the active
phosphorylase assay and the AMP assay. The former has physiological
concentrations of AMP (0.2 mM), ATP (2 mM), and glucose (10 mM) to simulate the concentrations of these
allosteric effectors in hepatocytes, and the latter has a saturating (5 mM) concentration of AMP (Table I). In untreated
hepatocytes (which express only liver phosphorylase), the activity
assayed with 5 mM AMP was 18% of total activity (Table I).
The difference in activity between untreated and AdCMV-MGP-treated
cells in the presence of 5 mM AMP (13-66 m-units/mg) was
equal to the difference in total phosphorylase (75-125 m-units/mg).
This confirms that 5 mM AMP fully activates the
b isoform of muscle but not the b isoform of
liver. The active phosphorylase assay shows a similar activity as the
phosphorylase a assay for the endogenous phosphorylase in
untreated hepatocytes but shows a 2-fold higher activity in hepatocytes
expressing muscle phosphorylase (Table I). This confirms that muscle
phosphorylase b but not liver phosphorylase b is
catalytically active at physiological concentrations of AMP and ATP.
Fig. 4 shows the effects of varying
concentrations of AMP in the absence or presence of 2 mM
ATP on endogenous phosphorylase activity in untreated hepatocytes and
on the muscle isoform (as determined by subtraction of endogenous
activity from AdCMV-MGP-treated cells). It is noteworthy that at
saturating concentrations of AMP (5 mM), the activity of
the muscle isoform is equal to total muscle activity, whereas the
activity of the endogenous enzyme is 18% of total liver phosphorylase.
This is due to the low affinity of liver phosphorylase b for
its substrate, inorganic phosphate, at saturating AMP concentrations
(16).
Table II shows the effects of the four
titers of AdCMV-MGP used in the rest of this study on the activity of
phosphorylase as determined by the different assays. With titrated
expression of the muscle isoform, there was negligible change in
phosphorylase a. However, there was a progressive increase
in activity by all the assays containing AMP. Control coefficients were
determined using either the active phosphorylase assay or the AMP
assay.
Expression of AdCMV-MGP Inhibits Glycogen Synthesis and Inactivates
Glycogen Synthase--
Expression of muscle phosphorylase in
hepatocytes using the titers of AdCMV-MGP shown in Table II caused a
sharp inhibition of glycogen synthesis at the lowest viral titer (5 µl/ml), with little further inhibition at higher titers (Fig.
5A). However, there was a
progressive decrease in the active form of glycogen synthase with
increasing phosphorylase overexpression (Fig. 5B). The
activity ratio of synthase (
The inactivation of synthase by treatment with AdCMV-MGP was not
associated with changes in glucose 6-P (means ± S.E. (in nmol/mg
protein): untreated cells, 0.92 ± 0.07; cells treated with 5 µl/ml AdCMV-MGP, 0.97 ± 0.08; cells treated with 10 µl/ml AdCMV-MGP, 0.95 ± 0.09; cells treated with 25 µl/ml AdCMV-MGP, 0.94 ± 0.02; and cells treated with 50 µl/ml AdCMV-MGP,
0.93 ± 0.04; n = 3) or changes in cell ATP
content (data not shown) or the rate of glycolysis as determined from
detritiation of [3-3H]glucose at 25 mM
glucose (in nmol glucose/h per mg: untreated cells, 357 ± 29;
cells treated with 5 µl/ml AdCMV-MGP, 345 ± 32; cells treated
with 10 µl/ml AdCMV-MGP, 346 ± 33; cells treated with 25 µl/ml AdCMV-MGP, 337 ± 39; and cells treated with 50 µl/ml AdCMV-MGP, 361 ± 38; n = 3). Incorporation of
[U-14C]glucose into triacylglycerol during incubation
with 25 mM glucose was not affected by expression of muscle
phosphorylase (in nmol/6 h/mg: control, 16.1 ± 0.6; cells treated
with insulin, 24.9 ± 2.1; cells treated with 10 µl/ml
AdCMV-MGP, 16.1 ± 1.4; and cells treated with 10 µl/ml
AdCMV-MGP + insulin, 22.8 ± 1.6; n = 3), whereas
incorporation of [U-14C]glucose into glycogen in the same
experiments was inhibited (in nmol/6 h/mg: control, 130 ± 15;
cells treated with insulin, 183 ± 21; cells treated with 10 µl/ml AdCMV-MGP, 76 ± 6*; and cells treated with 10 µl/ml
AdCMV-MGP + insulin, 117 ± 11* (n = 4; *,
p < 0.02 compared with no AdCMV-MGP)). Glycogen
deposition determined analytically (nmol/6 h/mg: control, 207 ± 22; cells treated with insulin, 335 ± 25; cells treated with 10 µl/ml AdCMV-MGP, 100 ± 8**; cells treated with 10 µl/ml
AdCMV-MGP + insulin, 192 ± 11*; n = 4 (* and
**, p < 0.02 and p < 0.01, respectively, as compared with no AdCMV-MGP)) was inhibited by
treatment with AdCMV-MGP.
Control Coefficients of Phosphorylase Determined by
Expression of Muscle Phosphorylase--
Control coefficients of
phosphorylase on glycogen synthesis (or on synthase activity),
determined from the initial slope of double logarithmic plots of
glycogen synthesis (or synthase activity) against phosphorylase
activity of the experiments in Fig. 5 are summarized in Table
III. Flux control coefficients of
phosphorylase on glycogen synthesis were about Effects of Glucokinase Overexpression on the Control Coefficient of
Phosphorylase--
Glucokinase has a very high flux control
coefficient on glycogen synthesis (12), which is counterbalanced in
part by the negative control exerted by its regulatory protein (14). To determine whether the control coefficient of phosphorylase on glycogen
synthesis is dependent on glucokinase activity, we overexpressed glucokinase by 60% above endogenous activity (27 ± 4 versus 17 ± 3 m-units/mg protein). Glucokinase
overexpression stimulated glycogen synthesis (Fig.
6A) and activated glycogen
synthase (Fig. 6B). Expression of muscle phosphorylase
inhibited glycogen synthesis by 50% in controls expressing endogenous
glucokinase and by 30% in cells overexpressing glucokinase (Fig.
6A) and caused parallel inactivation of glycogen synthase
(Fig. 6B). Total glucokinase activity and distribution
between free and bound states were not affected by phosphorylase
overexpression (data not shown). Fig. 6C shows the relation
between glycogen synthesis and active glycogen synthase. In the control
incubations with 25 mM glucose, there was a nonlinear
relation as observed with 10 mM glucose (Fig. 5C). In cells overexpressing glucokinase, the response was
shifted toward higher synthesis and synthase activity. The flux control coefficient of phosphorylase on glycogen synthesis (as determined from
the initial slope of the double logarithmic plot of the data in Fig.
6A) was significantly lower in cells overexpressing
glucokinase (Table III), whereas the control coefficient of
phosphorylase on glycogen synthase (Fig. 6B) was unchanged
(Table III).
CP-91149 Partially Counteracts the Effects of AdCMV-MGP--
To
test whether the inhibition of glycogen synthesis and inactivation of
glycogen synthase by treatment with AdCMV-MGP can be explained by the
catalytic activity of muscle phosphorylase b as opposed to
other mechanisms, we determined whether CP-91149 counteracts the
effects of phosphorylase overexpression. CP-91149 is a potent inhibitor
of human liver phosphorylase a (IC50 = 0.13 µM (5)) and human muscle phosphorylase a
(IC50 = 200 nM (24)). Fig.
7 shows that it is also a potent
inhibitor of muscle phosphorylase b (IC50 ~0.3
µM) assayed in the presence of 10 mM glucose,
0.2 mM AMP, and 2 mM ATP. Table
IV shows the combined effects of the inhibitor and phosphorylase overexpression. The inhibitor (10 µM) caused a 2-fold increase in glycogen synthesis
(p < 0.03) and cell glycogen content
(p < 0.04) and 40% activation of glycogen synthase
(p < 0.02) in the untreated controls expressing only endogenous phosphorylase, and it counteracted the suppression of
glycogen synthesis and inactivation of glycogen synthase in cells
treated with low titers of AdCMV-MGP. However, the inhibitor did not
fully counteract the effects of high titers of AdCMV-MGP (Table IV).
Because the control coefficients in Table III were determined from the
initial slope using the lowest titer of AdCMV-MGP, these values are
most likely due to the catalytic activity of muscle phosphorylase.
Metabolic control analysis is a powerful analytical approach to
describe how control of flux through a metabolic pathway is distributed
among enzymes that have direct or indirect effects on pathway flux (8).
The flux control coefficient of an enzyme is a measure of the
sensitivity of metabolic flux to small changes in enzyme activity or
concentration. Heinrich and Rapoport (25) defined the control strength
in terms of the fractional change in flux that results from a
fractional change in enzyme activity, whereas Kacser and Burns (26)
defined the sensitivity coefficient in terms of the fractional change
in flux that results from a fractional change in enzyme concentration.
If the enzyme rate is proportional to the enzyme concentration, then
the two definitions are equivalent. However, when an enzyme is
regulated by covalent modification, as in the case of liver
phosphorylase (1), then the relevant coefficient for determining the
degree of control on pathway flux is expressed in terms of the
activity of the enzyme. The control coefficient usually has a value
between zero (minimum control) and unity (high control) and is either
positive or negative, depending on whether the enzyme stimulates or
inhibits pathway flux. Enzymes with control coefficients greater than
unity are considered to be rare (23). In muscle, a high degree
of control of glucose utilization lies at or before glucose
phosphorylation (9-11), and in liver high degree of control lies at
glucose phosphorylation (12-14). In hepatocytes, glucokinase has a
high control coefficient on glycogen synthesis, which is
glucose-dependent. This is explained by the unique
compartmentation of glucokinase that involves
glucose-dependent partitioning of the enzyme between a free
active state and an inactive state bound to its regulatory protein (12,
14). The stimulation of glycogenic flux by an increase in glucokinase
activity by either translocation (19) or enzyme overexpression (27) is
at least in part explained by the increase in glucose 6-P concentration (11), the product of the glucokinase-catalyzed reaction, which is a
potent activator of glycogen synthase (28). Glucose 6-phosphatase, which lowers the concentration of glucose 6-P in hepatocytes, has a
negative control coefficient on glycogen synthesis (13). However, in
contrast with glucokinase, the control coefficient of glucose
6-phosphatase is much lower than unity and is glucose-independent, confirming that the high control coefficient of glucokinase and its
glucose dependence are best explained by the subcellular
compartmentation of glucokinase and its association with its regulatory
protein (14). The recent findings that stimulation of glycogen
synthesis by leptin is associated with inactivation of phosphorylase
(2) and that impaired glycogen synthesis in hepatocytes from fa/fa rats
is associated with elevated phosphorylase activity (7) raised the
question of the degree of control of glycogen synthesis by
phosphorylase activity. In this study, we demonstrate that phosphorylase has a very high negative flux control coefficient on
glycogen synthesis, based on two independent approaches (titration with
a specific phosphorylase inhibitor and expression of the muscle isoform
of phosphorylase).
Three key findings emerged from the studies with phosphorylase
inhibitor CP-91149. First, it caused time-dependent
inactivation of phosphorylase a and sequential activation
of glycogen synthase. This is analogous to the mechanism proposed by
Stalmans et al. (22) for the glucose-induced inactivation of
phosphorylase and sequential activation of glycogen synthase. Binding
of glucose to phosphorylase a causes a conformational change
(R-state to T-state) that renders the enzyme a better substrate for
dephosphorylation by protein phosphatase-1. Glucose thus favors the
conversion of phosphorylase a to phosphorylase b.
Because phosphorylase a is a potent inhibitor of glycogen
synthase phosphatase by binding to the C terminus of the liver-specific
glycogen-targeting subunit (GL) of protein phosphatase-1
(29, 30), the decrease in phosphorylase a alleviates the
inhibition of synthase phosphatase. This results in a delayed
activation of glycogen synthase relative to the inactivation of
phosphorylase, which has been described as the "sequential activation
of synthase" (1, 22). The present results support a model whereby
CP-91149 favors the T-conformation of phosphorylase and thereby causes
the inactivation of phosphorylase and sequential activation of
synthase. Second, the phosphorylase inhibitor markedly increases the
sensitivity of glycogen synthesis to glucose (S0.5, 19 versus 26 mM) by causing a greater fold
stimulation of glycogen synthesis at 5 mM glucose than at
20 mM glucose. This is consistent with the higher activity
of phosphorylase a at low glucose and the greater fractional
inactivation by the inhibitor at low glucose. The rate of glucose
phosphorylation in hepatocytes is a sigmoidal function with respect to
[glucose] but with a higher S0.5 for glucose than can be
explained by glucokinase kinetics (20 versus 9 mM) (31). This higher S0.5 for glucose
phosphorylation in the intact cell is explained by the glucokinase
regulatory protein, which functions as a competitive inhibitor with
respect to glucose and as a nuclear receptor for the enzyme (32, 33).
Glycogen synthesis has a higher S0.5 for glucose than
glucose phosphorylation in intact cells (33). The inhibitor studies
show that phosphorylase is a major component of the mechanism that
accounts for the difference in glucose saturation curves of glycogen
synthesis and glucose phosphorylation. Third, the titrations with
increasing concentration of inhibitor show a large fractional increase
in glycogen synthesis for a corresponding inactivation of
phosphorylase, with a control coefficient greater than unity.
We show in this study that muscle phosphorylase expressed using
recombinant adenovirus is a powerful tool to alter the catalytic activity of phosphorylase in hepatocytes independently of the cAMP
status and/or the phosphorylation state of the cell. In the absence of
glucagon, muscle phosphorylase is expressed mainly in the
unphosphorylated (b) form. However, it is partially
catalytically active at physiological concentrations of AMP and ATP,
unlike liver phosphorylase b (16), and this enables
determination of the flux control coefficient of phosphorylase in
defined substrate and hormone conditions. By using low titers of
adenovirus that result in small fractional changes in phosphorylase
activity and a short culture time after treatment with the adenovirus
(<20 h), secondary changes in gene expression are minimized. Two
metabolic effects of muscle phosphorylase expression were noted,
inactivation of glycogen synthase, which was progressive with enzyme
expression, and a decrease in glycogen synthesis, which reached a
plateau at low levels of phosphorylase expression. The lack of effect of AdCMV-MGP treatment on ATP, glucose 6-P, glucokinase activity, glycolysis, or conversion of glucose to triacylglycerol indicates that
at the viral titers and incubation times used, the effects of muscle
phosphorylase expression are confined to glycogen metabolism. The
inactivation of synthase and lack of effect on triacylglycerol metabolism contrast with findings on phosphorylase overexpression in
muscle cultures (34). This may represent a tissue difference, or it may
be related to the lower levels of phosphorylase overexpression used in
the present study.
The inhibition of glycogen synthase by muscle phosphorylase
overexpression is of interest because it was not associated with either
a change in glucose 6-P, an activator of synthase phosphatase (28), or
an increase in phosphorylase a, a potent allosteric inhibitor of synthase phosphatase (1). Two types of mechanism can be
considered, involving either catalytic activity of phosphorylase or an
effect of the protein independent of catalytic activity. Catalytic
activity of muscle phosphorylase may cause dissociation of glycogen
synthase from glycogen, an allosteric effector of the enzyme (35), or
from glycogenin, which is also a substrate for phosphorylase (36), or
it may cause dissociation of a glycogen-targeting subunit of protein
phosphatase-1 such as GL or PTG (37) from glycogen. Muscle
phosphorylase b may bind to a glycogen-targeting unit and
cause inactivation of synthase phosphatase activity either through
an allosteric effect (29, 30) or by competitive binding with glycogen
synthase (38). GL has a high-affinity site for phosphorylase a as well as a substrate site, whereas PTG has
a single binding site for glycogen synthase and phosphorylase (38). Inhibition of synthase phosphatase by phosphorylase b has
been demonstrated, but with much lower affinity than that for
phosphorylase a (39). The experiments with CP-91149, which
counteracted the inhibitory effects of low levels of muscle
phosphorylase overexpression on glycogen synthesis and glycogen
synthase, suggest that the catalytic activity of phosphorylase accounts
for the inhibition of synthase by low levels of phosphorylase
expression. However, an additional protein effect independent of
catalytic activity at higher levels of muscle phosphorylase expression
cannot be ruled out.
Two points are of interest with regard to the inhibition of glycogen
synthesis by phosphorylase overexpression. First, the high control
coefficient is observed at both 10 and 25 mM glucose and is
also observed in the presence of insulin. This contrasts with the
strong glucose dependence of the control coefficient of glucokinase
(12, 13). Second, unlike the control coefficients of glucokinase or its
regulatory protein (positive and negative, respectively), which are
sustained over a wide range of protein overexpression (2-3-fold above
endogenous activity (13)), inhibition of glycogen synthesis by
muscle phosphorylase reaches a plateau (50% inhibition) at low levels
of phosphorylase overexpression (<30% above endogenous
activity), with no further inhibition at higher protein
expression. If the inhibition of [14C]glucose
incorporation into glycogen were due to increased degradation of
glycogen rather than inhibition of synthesis, then a progressive or a
linear response as a function of phosphorylase activity would be
expected, as is observed for inactivation of synthase. A more plausible
explanation for the sharp inhibition of glycogen synthesis that reaches
a plateau at fairly low activities of phosphorylase is that it
represents inhibition of synthesis and that there are two compartments
of glycogen synthesis, only one of which is sensitive to inhibition by phosphorylase.
The lack of correlation between the rate of glycogen synthesis and the
activity of glycogen synthase at high phosphorylase overexpression
could be explained by compartmentation of glycogen synthase (40). The
rate of glycogen synthesis in hepatocytes may depend on the fraction of
glycogen synthase that is associated with glycogen or the protein
primer, glycogenin. An increase in phosphorylase activity may cause
dissociation of synthase from glycogenin, which is a substrate for
phosphorylase (36). By analogy with the high flux control coefficient
of glucokinase on glycogen synthesis, which is explained by the
subcellular compartmentation of glucokinase (12), the high flux control
coefficient of phosphorylase on glycogen synthesis could be due in part
to an effect of phosphorylase catalytic activity on the subcellular
compartmentation of glycogen synthase. This hypothesis is consistent
with both the higher flux control coefficient of phosphorylase on
glycogen synthesis relative to the control coefficient on glycogen
synthase and the effect of glucokinase overexpression, which lowers the
flux control coefficient of phosphorylase on glycogen synthesis but not
on synthase. Overexpression of glucokinase increases the hepatocyte
glucose 6-P content (27), and glucose 6-P affects the translocation of
glycogen synthase (41). A difference in subcellular compartmentation of
glycogen synthase in cells overexpressing glucokinase could therefore
explain the lower control coefficient and could also explain the lower fractional inhibition of glycogen synthesis by phosphorylase
overexpression in cells with elevated glucokinase activity.
One of the advantages of metabolic control analysis is that by
providing a quantitative estimate for the degree of control exerted by
an enzyme, it enables the study of how this control changes in
different physiological or pathological states (8). This study shows
that glucokinase overexpression by 60% above endogenous activity
lowers the control coefficient of phosphorylase on glycogen synthesis
by 50%. This implies that for a small activation of phosphorylase
(below that causing saturation of the response), a smaller inhibition
of glycogen synthesis would occur in cells with a higher ratio of
glucokinase to its regulatory protein. This is of interest from a
physiological perspective because the ratio of glucokinase to its
regulatory protein decreases during fasting and increases on refeeding
(42). Therefore, it can be inferred that the flux control coefficient
of phosphorylase on glycogen synthesis would be greatest at low ratios
of glucokinase to regulatory protein, such as those that occur in the
fasted to fed transition.
Several inhibitors of glycogen phosphorylase described recently
(5, 24, 43-46) could be of therapeutic benefit for inhibiting hepatic
glycogenolysis in type 2 diabetes. The high control coefficient of
phosphorylase on glycogen synthesis suggests that phosphorylase inhibitors would also be highly effective in promoting hepatic glycogen
synthesis in the absorptive state and that the overall effect of an
inhibitor that causes inactivation of phosphorylase and sequential
activation of synthase is to increase the affinity of glycogen
synthesis for glucose.
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
glucose 6-P/+ glucose P).
Phosphorylase was assayed spectrometrically in the glycogenolytic
direction from the phosphate-induced hydrolysis of glycogen coupled to
phosphoglucomutase and glucose 6-P dehydrogenase (2, 7). Phosphorylase
a was assayed as described in Ref. 2. Total liver or total
muscle phosphorylase was assayed as described in Ref. 7 by converting phosphorylase b to phosphorylase a with
phosphorylase kinase (22). Total muscle phosphorylase was also assayed
in the presence of 5 mM AMP under assay conditions that
were otherwise similar to those used for assay of phosphorylase
a. To determine the catalytic activity of the muscle isoform
under physiological conditions, an assay designated "active
phosphorylase assay" was used. This assay was based on the
phosphorylase a assay, except that 10 mM glucose, 0.2 mM AMP, and 2 mM ATP were
substituted for caffeine. Other assays were modifications of the
active phosphorylase assay, using different concentrations of AMP and
ATP. Enzyme activities are expressed as m-units/mg cell protein, where
1 m-unit converts 1 nmol substrate/min. Glucokinase activity (free and
bound fractions) was determined as described in Ref. 12.
J/J) resulting from
a fractional change in enzyme activity (e/e) (8). It was
determined from the initial slope of double logarithmic plots of flux
(J) against enzyme activity (e) for titrations of
CP-91149 or enzyme expression with Ad-CMV-MGP (23).
The control coefficient of phosphorylase on glycogen synthase
activity was determined from the slope of ln active glycogen synthase
against ln phosphorylase activity.
(Eq. 1)
RESULTS
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ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
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Fig. 1.
Inactivation of phosphorylase and sequential
activation of synthase by CP-91149. Hepatocytes were incubated in
medium with either 5 ( 
) or 15 mM (
) glucose for
3 h and with 2.5 µM CP-91149 for the time intervals
indicated. Control incubations (0.1% Me2SO) showed no
change in activity. The activities of phosphorylase a
(A) and active glycogen synthase (B) are
expressed as m-units/mg cell protein. Values represent the means ± S.E. for four cultures; p < 0.05 compared with
untreated cells.
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Fig. 2.
CP-91149 increases the sensitivity of
glycogen synthesis for glucose. Hepatocytes were incubated for
3 h with the concentrations of glucose indicated and either 0.1%
Me2SO alone ( ) or 2.5 µM CP-91149 ().
Parallel incubations were performed without or with
[U-14C]glucose for determination of phosphorylase
a (expressed as m-units/mg; A) and glycogen
synthesis (expressed as nmol/3 h per mg; B). The
concentration of glucose that causes a half-maximal rate
(S0.5) and the Hill coefficient (h) were
determined from Hill plots. Values represent the means ± S.E. for
four cultures.
1.6 ± 0.16 (means ± S.E.;
n = 4).
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Fig. 3.
Determination of the control coefficient of
phosphorylase on glycogen synthesis by titration with CP-91149.
A, hepatocytes were incubated in medium containing 15 mM glucose and varying concentrations of CP-91149 for
determination of phosphorylase a (expressed as m-units/mg)
and glycogen synthesis (expressed as nmol/3 h per mg). B,
glycogen synthesis versus phosphorylase a. The
control coefficient of phosphorylase a on glycogen synthesis
was determined from the initial slope of the double logarithmic plot
(B, inset). Values are the means ± S.E. for four
experiments.
Phosphorylase a and total phosphorylase in hepatocytes treated with
AdCMV-MGP
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Fig. 4.
Effects of AMP and ATP on endogenous and
expressed muscle phosphorylase activity. Phosphorylase activity
was assayed at the AMP concentrations indicated in either the absence
( and 
) or presence ( and 
) of 2 mM ATP in
extracts of hepatocytes that were either untreated (endogenous
activity, and ) or treated with AdCMV-MGP (50 µl/ml) ( and
). MGP activity in AdCMV-MGP-treated cells was determined by
subtraction of endogenous activity. Activities are expressed as
m-units/mg cell protein. Values in parentheses show the
activity at 5 mM AMP as a percentage of total phosphorylase
assayed by the phosphorylase kinase assay.
Effects of varying titers of AdCMV-MGP on phosphorylase activity in
different assay conditions
glucose 6-P/+ glucose P) decreased by 50% at the highest viral titers (control, 0.26 ± 0.03; 50 µl of AdCMV-MGP, 0.13 ± 0.02; insulin, 0.35 ± 0.02; 50 µl of AdCMV-MGP + insulin, 0.17 ± 0.02 (means ± S.E.);
n = 4). When the rate of glycogen synthesis was plotted
against the active glycogen synthase, the relation was nonlinear (Fig.
5C), indicating that progressive inactivation of glycogen
synthase is not associated with further inhibition of glycogen
synthesis. Insulin stimulated glycogen synthesis and activated glycogen
synthase in both untreated and AdCMV-MGP-treated cells, indicating that
phosphorylase expression does not override the effects of insulin.
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Fig. 5.
Muscle phosphorylase inhibits glycogen
synthesis and inactivates synthase. Hepatocytes were either
untreated ( and ) or treated ( and ) with the four titers
of AdCMV-MGP shown in Table II. Glycogen synthesis (expressed as nmol/3
h per mg) was determined from 3-h incubations with 10 mM
[U-14C]glucose without ( and ) or with ( and
) 10 nM insulin. Active glycogen synthase and
phosphorylase activity (determined by the active phosphorylase assay)
were determined from parallel incubations without radiolabel and are
expressed as m-units/mg. Values represent the means ± S.E. for
four cultures.
2 and were similar
regardless of whether they were determined by the active phosphorylase
assay or by the AMP assay (Table III). Similar coefficients were
obtained from the initial slopes by the other AMP-containing assays
shown in Table II (data not shown). The control coefficients of
phosphorylase on synthase activity were lower than those on
glycogen synthesis by about 50% (Table III).
Control coefficients of phosphorylase on glycogen synthesis and
glycogen synthase
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Fig. 6.
Combined effects of overexpression of
glucokinase and muscle phosphorylase on glycogen synthesis and glycogen
synthase activity. Hepatocytes were either untreated ( 
and )
or treated (
and ) with the four titers of AdCMV-MGP shown in
Table II without ( and ) or with ( and ) AdCMV-GKL.
Glycogen synthesis (expressed as nmol/3 h per mg) was determined from
3-h incubations with 25 mM [U-14C]glucose.
Active glycogen synthase and phosphorylase activity (determined by the
AMP assay) were determined from parallel incubations without radiolabel
and are expressed as m-units/mg. Values represent the means ± S.E. for four cultures.
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Fig. 7.
Inhibition of muscle phosphorylase
b by CP-91149. Purified muscle phosphorylase
b was assayed by the active phosphorylase assay containing
10 mM glucose, 0.2 mM AMP, and 2 mM
ATP at the concentrations of CP-91149 indicated. Activity in the
presence of inhibitor is expressed as a percentage of control activity
without inhibitor. Values represent the means ± S.E. for four
determinations.
Combined effects of AdCMV-MGP treatment and CP-91149
DISCUSSION
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ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
| |
ACKNOWLEDGEMENTS |
|---|
We thank Dr. Judith Treadway for help and advice with the CP-91149 studies, Dr. Gérald van de Werve for helpful discussions.
| |
FOOTNOTES |
|---|
* This work was supported by the Medical Research Council and by the Royal Society through an award under the European Science Exchange Programme.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.
¶ To whom correspondence should be addressed. Tel.: 44-191-2227033; Fax: 44-191-2220723; E-mail: Loranne.Agius@ncl.ac.uk.
Published, JBC Papers in Press, April 17, 2001, DOI 10.1074/jbc.M101454200
| |
ABBREVIATIONS |
|---|
The abbreviation used is: glucose 6-P, glucose 6-phosphate.
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ABSTRACT
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EXPERIMENTAL PROCEDURES
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DISCUSSION
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| 1. | Bollen, M., Keppens, S., and Stalmans, W. (1998) Biochem. J. 336, 19-31 |
| 2. | Aiston, S., and Agius, L. (1999) Diabetes 48, 15-20 |
| 3. | Browner, M. F., and Fletterick, R. J. (1992) Trends Biochem. Sci. 17, 66-71 |
| 4. | Johnson, L. N. (1992) FASEB J. 6, 2274-2282 |
| 5. | Martin, W. H., Hoover, D. J., Armento, S. J., Stock, I. A., McPherson, R. R., Danley, D. E., Stevenson, R. W., Barrett, E. J., and Treadway, J. L. (1998) Proc. Natl. Acad. Sci. U. S. A. 95, 1776-1781 |
| 6. | Board, M., Hadwen, M., and Johnson, L. N. (1995) Eur. J. Biochem. 228, 753-761 |
| 7. | Aiston, S., Peak, M., and Agius, L. (2000) Diabetologia 43, 589-597 |
| 8. | Kacser, H., and Burns, J. A. (1979) Biochem. Soc. Trans. 7, 1149-1160 |
| 9. | Shulman, R. G., Bloch, G., and Rothman, D. L. (1995) Proc. Natl. Acad. Sci. U. S. A. 92, 8535-8542 |
| 10. | Jucker, B. M., Barucci, N., and Shulman, G. I. (1999) Am. J. Physiol. 277, E505-E512 |
| 11. | Schulz, A. R. (1998) Arch. Biochem. Biophys. 353, 172-180 |
| 12. | Agius, L., Peak, M., Newgard, C. B., Gomez-Foix, A. M., and Guinovart, J. J. (1996) J. Biol. Chem. 271, 30479-30486 |
| 13. | Aiston, S., Trinh, K., Lange, A. J., Newgard, C. B., and Agius, L. (1999) J. Biol. Chem. 274, 24559-24566 |
| 14. | De la Iglesia, N., Mukhtar, M., Seoane, J., Guinovart, J. J., and Agius, L. (2000) J. Biol. Chem. 275, 10597-10603 |
| 15. | Gomez-Foix, A. M., Coats, W. S., Baque, S., Alam, T., Gerard, R. D., and Newgard, C. B. (1992) J. Biol. Chem. 267, 25129-25134 |
| 16. | Stalmans, W., and Gevers, G. (1981) Biochem. J. 200, 327-336 |
| 17. | Becker, T. C., Noel, R. J., Johnson, J. H., Lynch, R. M., Hirose, H., Tokuyama, Y., Bell, G. I., and Newgard, C. B. (1996) J. Biol. Chem. 271, 390-394 |
| 18. | Agius, L., Peak, M., and Alberti, K. G. M. M. (1990) Biochem. J. 266, 91-10 |
| 19. | Agius, L. (1997) Biochem. J. 325, 667-673 |
| 20. | Bligh, E. G., and Dyer, W. J. (1959) Can. J. Biochem. Physiol. 37, 911-917 |
| 21. | Thomas, J. A., Schlender, K. K., and Larner, J. (1968) Anal. Biochem. 25, 486-499 |
| 22. | Stalmans, W., De Wulf, H., Hue, L., and Hers, H.-G. (1974) Eur. J. Biochem. 41, 127-134 |
| 23. | Fell, D. A. (1992) Biochem. J. 286, 313-330 |
| 24. | Hoover, D. J., Lefkowitz-Snow, S., Burgess-Henry, J. L., Martin, W. H., Armento, S. J., Stock, I. A., McPherson, R. K., Genereux, P. E., Gibbs, E. M., and Treadway, J. L. (1998) J. Med. Chem. 41, 2934-2938 |
| 25. | Heinrich, R., and Rapoport, T. A. (1974) Eur. J. Biochem. 42, 89-95 |
| 26. | Kacser, H., and Burns, J. A. (1973) Symp. Soc. Exp. Biol. 27, 65-104 |
| 27. | Seoane, J., Gomez-Foix, A. M., O'Doherty, M., Gomez-Ara, C., Newgard, C. B., and Guinovart, J. J. (1996) J. Biol. Chem. 271, 23756-23760 |
| 28. | Guinovart, J. J., Gómez-Foix, A. M., Seoane, J., Fernandez-Novell, J. M., Bellido, D., and Vilaro, S. (1997) Biochem. Soc. Trans. 25, 157-160 |
| 29. | Doherty, M. J., Moorhead, G., Morrice, N., Cohen, P., and Cohen, P. T. W. (1995) FEBS Lett. 375, 294-298 |
| 30. | Armstrong, C. G., Doherty, M. J., and Cohen, P. T. W. (1998) Biochem. J. 336, 699-704 |
| 31. | Bontemps, F., Hue, L., and Hers, H.-G. (1978) Biochem. J. 174, 603-611 |
| 32. | Van Schaftingen, E. (1989) Eur. J. Biochem. 179, 179-184 |
| 33. | Agius, L. (1998) Adv. Enzyme Regul. 38, 303-311 |
| 34. | Baqué, S., Guinovart, J. J., and Gómez-Foix, A. M. (1996) J. Biol. Chem. 271, 2594-2598 |
| 35. | Solling, H. (1979) Eur. J. Biochem. 94, 231-242 |
| 36. | Cao, Y., Skurat, A. V., DePaoli-Roach, A. A., and Roach, P. J. (1993) J. Biol. Chem. 268, 21717-21721 |
| 37. | Newgard, C. B., Brady, M. J., O'Doherty, R. M., and Saltiel, A. R. (2000) Diabetes 49, 1967-1977 |
| 38. | Fong, N. M., Jensen, T. C., Shah, A. S., Parekh, N. N., Saltiel, A. R., and Brady, M. J. (2000) J. Biol. Chem. 275, 35034-35039 |
| 39. | Alemany, S., and Cohen, P. (1986) FEBS Lett. 198, 194-202 |
| 40. | Fernandez-Novell, J. M., Bellido, D., Vilaro, S., and Guinovart, J. J. (1997) Biochem. J. 321, 227-231 |
| 41. | Fernandez-Novell, J. M., Arino, J., Vilaro, S., Bellido, D., and Guinovart, J. J. (1992) Biochem. J. 288, 497-501 |
| 42. | Vandercammen, A., and Van Schaftingen, E. (1993) Biochem. J. 294, 551-556 |
| 43. | Bergans, N., Stalmans, W., Goldmann, S., and Vanstapel, F. (2000) Diabetes 49, 1419-1426 |
| 44. | Fosgerau, K., Westergaard, N., Quistorff, B., Grunnet, N., Kristiansen, M., and Lundgren, K. (2000) Arch. Biochem. Biophys. 380, 274-284 |
| 45. | Oikonomakos, N. G., Tsitsanou, K. E., Zographos, S. E., Skamnaki, V. T., Goldman, S., and Bischoff, H. (1999) Protein Sci. 8, 1930-1945 |
| 46. | Rath, V. L., Ammirati, M., Danley, D. E., Ekstrom, J. L., Gibbs, E. M., Hynes, T. R., Mathiowetz, A. M., McPherson, R. K., Olson, T. V., Treadway, J. L., and Hoover, D. J. (2000) Chem. Biol. 7, 677-682 |
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