From the Programme in Cell Biology, The Hospital for Sick Children, Toronto, Ontario M5G 1X8, Canada
Received for publication, October 30, 2002, and in revised form, February 12, 2003
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ABSTRACT |
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Insulin stimulates glucose uptake in
skeletal muscle cells and fat cells by promoting the rapid
translocation of GLUT4 glucose transporters to the plasma membrane.
Recent work from our laboratory supports the concept that insulin also
stimulates the intrinsic activity of GLUT4 through a signaling pathway
that includes p38 MAPK. Here we show that regulation of GLUT4 activity
by insulin develops during maturation of skeletal muscle cells into
myotubes in concert with the ability of insulin to stimulate p38 MAPK. In L6 myotubes expressing GLUT4 that carries an exofacial
myc-epitope (L6-GLUT4myc), insulin-stimulated GLUT4myc translocation
equals in magnitude the glucose uptake response. Inhibition of p38 MAPK with SB203580 reduces insulin-stimulated glucose uptake without affecting GLUT4myc translocation. In contrast, in myoblasts, the magnitude of insulin-stimulated glucose uptake is significantly lower
than that of GLUT4myc translocation and is insensitive to SB203580.
Activation of p38 MAPK by insulin is considerably higher in myotubes
than in myoblasts, as is the activation of upstream kinases MKK3/MKK6.
In contrast, the activation of all three Akt isoforms and GLUT4
translocation are similar in myoblasts and myotubes. Furthermore,
GLUT4myc translocation and phosphorylation of regulatory sites on Akt
in L6-GLUT4myc myotubes are equally sensitive to insulin, whereas
glucose uptake and phosphorylation of regulatory sites on p38 MAPK show
lower sensitivity to the hormone. These observations draw additional
parallels between Akt and GLUT4 translocation and between p38 MAPK and
GLUT4 activation. Regulation of GLUT4 activity by insulin develops upon
muscle cell differentiation and correlates with p38 MAPK activation by insulin.
Insulin is the major regulator of blood glucose levels in the fed
state when skeletal muscle becomes the primary consumer of glucose (1).
The rate-limiting determinant of glucose utilization by muscle is its
uptake mediated by glucose transporters (2). GLUT41 is the most abundant
glucose transporter isoform in skeletal muscle and adipose tissue
(3-6) and is responsible for the majority of
insulin-dependent glucose uptake in these tissues (7, 8). It has long been recognized that insulin promotes the rapid
translocation of GLUT4 glucose transporters from intracellular membrane
compartments to the plasma membrane (3-5, 9, 10). Current views hold that GLUT4 translocation is entirely responsible for the increase in
glucose uptake in response to insulin. However, the two widely used
methods to measure GLUT4 translocation, subcellular fractionation and
photoaffinity labeling of surface GLUT4, have drawbacks that compromise
their ability to measure GLUT4 translocation accurately (see
"Discussion"). These methods have arrived at variable conclusions regarding how closely GLUT4 translocation matches the stimulation of
glucose uptake by insulin (5, 6, 11-19). Thus, the possibility exists
that the intrinsic activity of glucose transporters could be regulated
in response to the hormone, and new approaches to measure GLUT4
translocation in intact cells are required (without cellular
homogenization or protein immunoprecipitation).
To this end, we have used L6 muscle cells that stably overexpress GLUT4
encoding a myc epitope in its large exofacial loop (L6-GLUT4myc cells
(20)). GLUT4myc can be readily detected at the cell surface of intact
cells by an enzyme-linked immunosorbent-like assay (21). In these
cells, GLUT4myc shows insulin-regulated behavior consistent with that
of GLUT4 in 3T3-L1 adipocytes (22, 23). Thus, 90% of GLUT4myc is
sequestered intracellularly in the basal state, and a significant
portion translocates to the cell surface in response to insulin (24).
All GLUT4myc molecules are available for recycling to the cell surface
(25). The exocytic and endocytic rates of GLUT4myc (24) mimic those
reported for GLUT4 (26, 27). The Km of glucose
uptake is similar in L6GLUT4myc to that in the parental L6 cells (28,
29). Most significantly, the 10% of total GLUT4myc present at the cell
surface is still higher than the endogenous levels of GLUT1 or GLUT3
(by almost 100-fold (30)), and is responsible for both basal and insulin-stimulated glucose uptake. This functional preponderance was
established by the nearly complete inhibition of both basal and
insulin-stimulated glucose uptake rates by the drug indinavir (30, 31),
a rather selective inhibitor of glucose influx through GLUT4 but not
GLUTs 1, 3, and 8 (32, 33). Hence, L6-GLUT4myc cells are uniquely
suitable to make direct comparisons between glucose uptake through
GLUT4 and GLUT4 translocation. The molar expression of GLUT4myc in
L6-GLUT4myc cells is 5-10 fold higher than that of endogenous GLUT4 in
skeletal muscle (30). Using these cells we have observed that
insulin-dependent GLUT4 translocation and stimulation of
glucose uptake can be segregated in time (28), by their temperature
sensitivity (28), their susceptibility to inhibition by wortmannin
(34), and most strikingly, by specific inhibitors of p38 MAPK.
The latter (pyridinylimidazoles SB203580 and SB202190 and chemically
distinct aza-azulenes A291077 and A304000) reduced the insulin response
of glucose uptake without interfering with GLUT4 translocation in
L6-GLUT4myc myotubes and 3T3-L1 adipocytes without directly inhibiting
glucose transporters (28, 35, 36).
L6-GLUT4myc muscle cells undergo differentiation from myoblasts to
multinucleated myotubes through multiple cell fusions (37, 38). In
search for information on the mechanisms responsible for the
segregation of GLUT4 translocation and glucose uptake in myotubes, we
examined the maturation of the insulin response of GLUT4 translocation
and glucose uptake during myogenesis. We report that, in the myoblast
stage, insulin-stimulated glucose uptake is lower than in myotubes and
is not sensitive to inhibition of p38 MAPK. Yet, translocation of
GLUT4myc is similar in both stages of cellular differentiation.
Moreover, insulin stimulates p38 MAPK and its upstream activators
MKK3/6 in myotubes but slightly if at all in myoblasts. These results
further correlate the p38 MAPK pathway to GLUT4 activation, because
both events mature during L6 cell differentiation from myoblasts into myotubes.
Materials--
SB203580 was purchased from Calbiochem (La Jolla,
CA). Cytochalasin B, O-phenylenediamine dihydrochloride, and
2-deoxyglucose were obtained from Sigma (St. Louis, MO).
2-Deoxy-D-[3H]glucose was purchased
from ICN (Irvine, CA). Monoclonal anti-myc (9E10) antibody and
antibodies to p38 L6-GLUT4myc Cell Line and Cell Culture--
For selective
experiments, wild-type L6 myoblasts grown and differentiated into
myotubes as previously reported (39) were used where indicated.
Otherwise, L6 myoblasts stably expressing GLUT4myc, created (20) and
characterized (40, 41) as described previously, were used throughout
the study. L6-GLUT4myc cells were maintained in minimal essential
medium- 2-Deoxyglucose Uptake--
Following serum depletion, cells were
treated with 10 µM SB203580 for 20 min before the
addition of insulin at the indicated concentrations and times as
described in the figure legends. Following treatments, cells were
rinsed and immediately used for measurement of 2-deoxyglucose uptake in
the absence of inhibitors as described previously (28). For the
insulin-stimulated time-course analysis, cells were grown in six-well
plates and treated as indicated, and then 2-deoxyglucose uptake was
measured for 30 s. Nonspecific uptake was determined in the
presence of 10 µM cytochalasin B, and this value was
subtracted from all other values. Cell-associated radioactivity was
determined by lysing the cells with 0.05 N NaOH, followed
by liquid scintillation counting. Total cellular protein was determined
by the Bradford method.
GLUT4myc Translocation--
GLUT4myc levels at the cell surface
of intact myoblasts or myotubes were measured by an antibody-coupled
colorimetric assay as described (24) using the anti-myc monoclonal 9E10
as the primary antibody and a donkey anti-mouse IgG conjugated to
horseradish peroxidase as the secondary antibody.
Immunoblotting and Phosphorylation of p38 MAPK,
MKK3/6, and Akt--
Briefly, cells in six-well plates were
incubated as indicated, lysed on ice with 300 µl of 2× Laemmli
sample buffer per well supplemented with 7.5% Immunoprecipitation and Assay of p38 MAPK and Akt
Activities--
Immunoprecipitation of p38 MAPK
Immunoprecipitation of Akt1, Akt2, or Akt3 from 200 µg (total
protein) of TX-100 detergent cell lysates containing phosphatase and
protease inhibitors overnight at 4 °C was performed as described (40). Akt immunocomplexes were incubated in 30 µl of kinase buffer
supplemented with Crosstide (150 µg/condition), 5 µM
ATP, and 2 µCi of [ Statistical Analysis--
Statistical analysis was performed
using either unpaired Student's t test or analysis of
variance test (Fischer, multiple comparisons) as indicated in the
figure legends.
Differential Gains in Insulin-stimulated Glucose Uptake and GLUT4
Translocation during Myogenesis--
L6 myoblasts and myotubes were
treated in parallel with 100 nM insulin for 2-15 min at
37 °C, followed by determination of 2-deoxyglucose uptake or
GLUT4myc translocation. In these experiments, insulin continued to be
present during the 30-s uptake of [3H]2-deoxyglucose (see
"Experimental Procedures"). In this way, the time courses of
insulin-stimulated GLUT4myc translocation and glucose uptake could be
effectively compared at each time of insulin incubation. In myoblasts,
the maximal -fold stimulation of glucose uptake by insulin (15-min time
point) was lower than the -fold stimulation in GLUT4myc translocation
(1.6 ± 0.1-fold compared with 2.4 ± 0.1, respectively, Fig.
1A). On the other hand, in
myotubes insulin-stimulated both responses more than 2-fold by 15 min
(2.2 ± 0.1-fold for glucose uptake compared with 2.4 ± 0.1, for translocation, Fig. 1B). Interestingly, the magnitude of
the GLUT4myc translocation response was comparable in myoblasts and
myotubes, but the full response of insulin-stimulated glucose uptake
was not realized in the myoblasts.
In myoblasts, there was no difference in the rate of stimulation of
glucose uptake by insulin and GLUT4myc translocation at the early times
of insulin addition. The estimated t1/2 of glucose
uptake was 3.3 min and that of the arrival of GLUT4myc at the plasma
membrane also was 3.3 min. In myotubes, GLUT4myc rapidly translocated
to the plasma membrane with a t1/2 of 2.0 min. In
contrast, the stimulation of glucose uptake in L6 myotubes was delayed
by a lag of about 2 min before a significant response could be
measured. Thus, maximal stimulation of glucose uptake is reached only
between 10 and 15 min in the myotubes and is significantly delayed
(t1/2 of 5 min) with respect to the response in myoblasts.
SB203580 Inhibits 2-Deoxyglucose Uptake in Myotubes but Not in
Myoblasts--
L6-GLUT4myc myoblasts or myotubes were preincubated
with the pyridinylimidazole SB203580 (10 µM) or
Me2SO vehicle only for 20 min. This was followed by
treatment with insulin (100 nM) for an additional 20 min in
the presence of SB203580. Uptake of 2-deoxyglucose was then measured in
the absence of the inhibitor. Insulin caused a significant increase in
glucose uptake in myoblasts (1.7 ± 0.1-fold above basal,
p < 0.001; Fig.
2A) and myotubes (2.2 ± 0.2-fold above basal, p < 0.001, Fig. 2B).
Preincubation with SB203580 did not have a significant effect on either
basal or insulin-stimulated glucose uptake in myoblasts (Fig.
2A). Similarly, preincubation with SB203580 did not
affect the basal rate of glucose uptake in myotubes. However, it
reduced the stimulation of glucose uptake by 65% (insulin: 2.2 ± 0.2-fold, insulin plus SB203580: 1.4 ± 0.1-fold,
p < 0.001, Fig. 2B). It is unlikely that
pyridinylimidazoles act by directly binding and inhibiting GLUT4myc,
because both basal and insulin-stimulated rates of glucose uptake are
mediated by GLUT4myc in these cells (30, 31), but only the stimulated uptake was reduced by the drug. Moreover, SB203580 was not present during the transport assay. Furthermore, SB203580 did not inhibit glucose uptake when added only to the glucose transport solution in
glucose uptake assays lasting up to 30 min (28, 35), confirming that
the inhibition is due to an event different from direct inhibition of
GLUT4. Nonetheless, to ascertain that the myc epitope is not related to
the sensitivity to SB203580, the selective inhibition of
insulin-stimulated glucose uptake by SB203580 in myotubes vis à vis was confirmed in wild-type L6 muscle cells (Table
I). This result conforms to our previous
observation that SB203580 reduces the insulin-dependent
portion of glucose uptake in wild-type L6 myotubes (35) and further
establishes that maturation into myotubes is required for this
susceptibility to SB203580 to manifest. Hence, glucose uptake in
L6-GLUT4myc cells obeys similar regulation as in parental,
untransfected cells.
Insulin increased GLUT4myc at the cell surface by 2.3 ± 0.2-fold
in myoblasts and 2.5 ± 0.2-fold in myotubes (Fig. 2, C
and D). In contrast to the stimulation of glucose uptake,
SB203580 did not affect GLUT4myc density at the cell surface of basal
or insulin-stimulated myotubes (Fig. 2D). This suggests that
pretreatment with SB203580 reduces the glucose transport activity of
GLUT4myc molecules that had been fully inserted into the plasma
membrane, consistent with previous observations (28, 35).
As shown in Fig. 1 above, the time course and maximal response to
insulin of glucose uptake in myotubes differs from that in myoblasts.
The myotubes responded with an initial delay but then achieved a higher
stimulation of glucose uptake than the myoblasts. Given that SB203580
lowered the maximal glucose uptake response in myotubes (see Fig. 2),
we examined how inhibition of p38 MAPK may affect the time course of
insulin-stimulated glucose uptake in myotubes. Preincubation of
L6-GLUT4myc myotubes with SB203580 prior to determination of the
insulin-stimulated time course of glucose uptake reduced the maximal
stimulation of glucose uptake to the same levels as in
insulin-stimulated myoblasts (Fig. 3).
This observation is in keeping with the lack of inhibition of insulin
action by SB203580 in myoblasts. However, the lag in stimulation was
not prevented (Fig. 3). These tantalizing results raise the possibility
that an event downstream of p38 MAPK is responsible for the maximal
stimulation of glucose uptake and that other factors upstream or
parallel to p38 MAPK contribute to the delay in stimulation of glucose
uptake observed in myotubes relative to myoblasts (see
"Discussion").
Insulin-induced p38 MAPK Phosphorylation Increases during
Myogenesis--
Activation of p38 MAPK by diverse stimuli leads to its
phosphorylation by upstream kinases on tyrosine and threonine residues in the TGY motif of its regulatory domain (42). The phosphorylation of
these two sites is widely used as an indication of heightened p38 MAPK
activity and can be detected with anti-phospho p38 MAPK antibody that
recognizes the dual phosphorylated form of p38 MAPK isoforms Insulin-stimulated p38 MAPK Basal p38 MAPK Activity Decreases during
Myogenesis--
Interestingly, the basal p38 MAPK phosphorylation
status decreased with differentiation of L6 cells. Phospho-p38 MAPK
levels in myotubes were only 45% of that seen in myoblasts (Table
II). The basal activity for the Insulin-induced MKK3/6 Phosphorylation Is Regulated
during Myogenesis--
The dual-specificity tyrosine and serine
protein kinases MKK3 and MKK6 are upstream kinases of p38 MAPK family
members activated by extracellular stresses and cytokines (43). MKK3
activates the p38 MAPK Activation of Each Akt Isoform by Insulin Is Comparable in
Myoblasts and Myotubes--
A number of studies support a role for Akt
in GLUT4 translocation (40, 44-47). In L6 myoblasts, overexpression of
a dominant-negative mutant of Akt1 blocked insulin-stimulated GLUT4
translocation (40). This mutant also prevented the insulin-induced
acceleration of the transit of GLUT4 through the recycling endosome
(25). As shown in Fig. 1, the rates by which insulin increased GLUT4 translocation in myoblasts and myotubes were strikingly similar. This
observation suggests that the translocation response of the GLUT4
system is fully developed at both stages of the myogenic process. To
document this possibility we measured the time course of activation by
insulin for Akt isoforms 1, 2, and 3 (protein kinase B GLUT4 Translocation Has Greater Insulin Sensitivity as Compared
with Glucose Uptake--
The time delay observed in the glucose uptake
response of the myotubes in the first 2 min after the addition of
insulin (Fig. 1) raised the possibility that glucose uptake may be less
sensitive to the hormone compared with GLUT4 translocation. Indeed,
focusing on the range of concentrations of 10 nM and below
(where insulin action is deemed to occur primarily through insulin
receptors (48)) we found that glucose uptake was markedly less
sensitive to insulin compared with GLUT4myc translocation (Fig.
7A). At higher insulin
concentrations, both curves followed relatively similar behavior (Fig.
7A, inset).
The striking difference in insulin sensitivity of GLUT4 translocation
and glucose uptake prompted an examination of the insulin sensitivities
of enzyme activities thought to lead to each of these phenomena,
respectively, Akt and p38 MAPK. The results in Fig. 7B
illustrate that Akt phosphorylation is more sensitive to insulin
compared with p38 MAPK phosphorylation. These results qualitatively
link Akt with GLUT4 translocation and p38 MAPK with glucose uptake at
low insulin doses.
The development of the L6 cell line that ectopically expresses
high levels of GLUT4myc has allowed us to differentiate GLUT4 translocation from GLUT4 transport activity under diverse conditions. An important feature of this cell line is that glucose uptake in the
basal and insulin-stimulated states in both myoblasts and myotubes is
mediated by GLUT4myc (30, 31). We found that insulin-stimulated GLUT4myc translocation precedes the stimulation of glucose uptake in
time, suggesting that GLUT4myc may undergo an activation step following
its insertion into the plasma membrane (28). Furthermore, we identified
p38 MAPK as a signal that may be involved in this activation of GLUT4,
because inhibitors of p38 MAPK reduce the insulin response of glucose
uptake without affecting GLUT4 translocation (28, 35). This link
between p38 MAPK and glucose uptake has been extended by the inhibition
of insulin-stimulated glucose uptake by two unrelated families of p38
MAPK inhibitors and to a dominant-negative p38 MAPK mutant expressed in
3T3-L1 adipocytes (36).
Here we further investigate the phenomenon of GLUT4 activation and find
that insulin-stimulated GLUT4 activation and p38 MAPK stimulation
develop upon differentiation of L6 muscle cells from myoblasts into
myotubes. In myoblasts, insulin causes a minor stimulation of the
kinase and produces a submaximal stimulation of glucose uptake
(1.6-fold) despite full translocation of GLUT4 to the plasma membrane
(2.4-fold). In differentiated myotubes, insulin stimulates glucose
uptake maximally and stimulates p38 MAPK robustly. Similarly, the
putative upstream activators of p38 MAPK, MKK3/6, are phosphorylated in
response to insulin in myotubes but not in myoblasts. In contrast to
the maturation of these responses, GLUT4 translocation and stimulation
of the phosphatidylinositol 3-kinase-Akt signaling axis, thought to be
involved in signaling GLUT4 translocation by insulin, are similar in
myoblasts and myotubes. In addition, the stimulation of GLUT4
translocation and Akt activation are more sensitive to insulin than the
stimulation of glucose uptake and p38 MAPK. These results segregate
GLUT4 translocation with stimulation of Akt on the one hand and GLUT4
activation and stimulation of p38 MAPK on the other.
Methodological Considerations in the Analysis of GLUT4
Activity--
Insulin stimulates rapidly the
Vmax of glucose uptake into muscle and fat cells
without altering the Km of the transporter proteins
for glucose (49, 50). An accepted explanation of this finding is a rise
in the number of glucose transporters at the cell surface (GLUT4
translocation). However, the magnitude of GLUT4 translocation
frequently reported in the literature does not account for the greater
magnitude in insulin-stimulated glucose uptake, when GLUT4 levels are
assessed in isolated membranes from muscle and adipose tissue (6,
13-17). The discrepancy remains when glucose uptake and GLUT4 content
are measured in the same vesicular membrane preparation (12, 18). These
results suggest that activation of GLUT4 may occur in response to insulin.
On the other hand, studies using the ATB-BMPA glucose transporter
photolabel have supported the theory that GLUT4 translocation is the
only mechanism by which insulin increases glucose uptake, because in
many cases the magnitude of insulin-stimulated translocation approximates the increase in glucose uptake (51-54). However, early studies proposed that the ATB-BMPA interaction with glucose
transporters reflects not only their availability at the cell surface
but also their state of activity, because the label binds to the
glucose-binding site on the transporters (55, 56). A similar argument
has been made in cells where ATB-BMPA binding to GLUT4 or GLUT1
increases without net gain in surface transporters assessed by an
alternate method of biotinylation of the transporters at the cell
surface (56-58). Moreover, changes in the Ki of
inhibition of glucose uptake by ATB-BMPA were noted upon treatment of
brown adipocytes with norepinephrine (16). Lastly, early on during the
characterization of the photolabel it was reported that cytochalasin B,
a potent non-competitive inhibitor of glucose uptake that binds to the
inward-facing glucose-binding site of glucose transporters, precludes
exofacial binding of ATB-BMPA to the glucose transporter (55). The
ensuing explanation was that the outward-facing and inward-facing
glucose-binding sites are mutually exclusive. Potentially, a cytosolic
protein or a post-translational modification on the glucose transporter
could lock the glucose-binding site of the glucose transporter in an
inward-facing position, preventing it from bringing glucose into the
cell. We surmise that an insulin signal could conceivably relieve this
constraint. Based on the time course of insulin-dependent
GLUT4 translocation and stimulation of glucose uptake, this would occur
subsequent to GLUT4 insertion into the plasma membrane and would
require input from insulin-stimulated p38 MAPK activity.
Correlation between p38 MAPK and Glucose Uptake--
The
hypothesis arises that the intrinsic activity of glucose transporters
can be regulated by the cell in response to insulin, and documenting it
requires methodological approaches that can compare glucose uptake and
gain in surface GLUT4 in equivalent cellular preparations. In the
present study, we find that inhibition of p38 MAPK with SB203580 lowers
the insulin response of glucose uptake in myotubes to match the rate
observed in myoblasts. This is consistent with our previous
demonstration that SB203580 reduces the Vmax of
insulin-stimulated glucose uptake but not the Km in
L6 myotubes (28). However, the time lag between GLUT4 translocation and
glucose uptake in particular to myotubes is unaffected by inhibitors of
p38 MAPK (Fig. 3), suggesting that additional mechanisms define the
response to the hormone in mature cells. It is possible that this lag
results from a delay in either p38 MAPK activation and/or in the
removal of an inhibitor or the recruitment of an activating protein
shortly after GLUT4 is inserted into the plasma membrane.
Supporting the concept that activation of GLUT4 occurs in addition to
its translocation in response to insulin, these two processes also
differ in their sensitivity to insulin. GLUT4 translocation occurs at
slightly lower concentrations of insulin than are required to stimulate
glucose uptake (Fig. 7). Both responses occur within the range of
signaling through the insulin receptor. Similarly, activation of
Akt/protein kinase B is more insulin-sensitive compared with p38 MAPK.
We hypothesize that at some point along the insulin signaling pathway,
the signals leading to the activation of Akt and GLUT4 translocation
segregate from those leading to p38 MAPK activation and stimulation of
glucose uptake. Indeed, activation of p38 MAPK by insulin is not
down-stream of phosphatidylinositol 3-kinase (34). Conceivably, steps
downstream of Akt and p38 MAPK or the GLUT4 molecules itself could be
the site of integration of these separate signals, resulting in
insulin-stimulated glucose uptake. The points of divergence and
convergence are the subject of separate investigation.
Maturation of the p38 MAPK and Glucose Uptake Responses during
Myogenesis--
The lower stimulation of glucose uptake by insulin
despite full insulin-stimulated GLUT4 translocation in myoblasts
suggests that GLUT4 activation does not occur at this stage of
myogenesis. Moreover, as stated above, SB203580 treatment of L6
myotubes brings down the glucose uptake response of L6 myotubes to the
level of the response in myoblasts. This difference between myoblasts
and myotubes correlates with the ability of insulin to phosphorylate MKK3/6 and activate p38 MAPK and with the acquisition of susceptibility to inhibitors of p38 MAPK given prior to insulin action. Similarly, p38
MAPK was activated in 3T3-L1 adipocytes (36) but not in undifferentiated 3T3-L1
fibroblasts.2 One possible
reason for the increased regulation of p38 MAPK by insulin could be the
increase in insulin receptor density as L6 and 3T3-L1 cells
differentiate (59, 60). However, this is unlikely because activation of
Akt isoforms in myoblasts is comparable to that in myotubes.
Alternatively, a differentiation-dependent change in the
expression of a putative p38 MAPK-regulatory protein could be the
explanation. Finally, the proximity of p38 MAPK activators and p38 MAPK
may be favored in myotubes. Future studies will be required to examine
this possibility.
Inhibitors of p38 MAPK (the pyridinylimidazoles SB203580 and SB202190
and the aza-azulenes A291077 and A304000) reduce insulin-stimulated glucose uptake without affecting GLUT4 translocation in L6 myotubes (28, 36). It is important to reiterate that the p38 MAPK inhibitors cannot directly inhibit GLUT4 glucose transporters, because they have
no effect on glucose uptake when added only to the transport assay (28,
35, 36). In addition, basal glucose uptake, which is mediated entirely
by GLUT4 in these cells (30, 31), is not inhibited by preincubation
with the p38 MAPK inhibitors (28, 35, 36). The connection between p38
MAPK and GLUT4 activation is strengthened by the similar
IC50 value of pyridinylimidazoles to reduce
insulin-stimulated glucose uptake and inhibit p38 MAPK activity (28).
Moreover, inducible expression of a dominant-negative mutant of p38
MAPK resulted in lower insulin-stimulated glucose uptake in 3T3-L1
adipocytes (36). Thus, p38 MAPK appears to contribute to
insulin-stimulated glucose uptake not only in myotubes but also in
3T3-L1 adipocytes and in isolated white and brown adipocytes and
skeletal muscles (28, 35, 61, 62). It is plausible that a lower GLUT4
activation contributes to the insulin resistance of glucose uptake in
peripheral tissues that precedes or accompanies type 2 diabetes. Hence,
a full understanding of the mechanism by which insulin regulates GLUT4
activation could give rise to the possibility of new therapeutic
interventions to improve glucose utilization.
In conclusion, the regulation of the activity of glucose uptake by p38
MAPK in L6-GLUT4myc cells develops with differentiation of the cell
from myoblasts into myotubes. Inhibiting p38 MAPK reduces the
stimulation of glucose uptake in myotubes to the level observed in
myoblasts. In contrast, GLUT4 translocation is equally developed in
myoblasts as in myotubes. In myotubes, GLUT4 translocation and Akt
stimulation had higher insulin sensitivities than stimulation of
glucose uptake and p38 MAPK phosphorylation. These observations draw
important parallels between Akt and GLUT4 translocation and between p38
MAPK and GLUT4 activation.
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
(C-20), p38
MAPK (K-16), Akt1, and Akt2 were
from Santa Cruz Biotechnology (Santa Cruz, CA). Antibodies to Akt3 and
Crosstide (Akt peptide substrate) were purchased from Upstate
Biotechnology (Lake Placid, NY). ATF2 recombinant protein, specific
antibodies to phosphorylated p38 MAPK (Thr-180 and Tyr-182),
phosphorylated MKK3/6 (Ser189/207) and phospho-Akt (S473), and anti-pan
p38 MAPK antibodies were purchased from Cell Signaling (Beverly, MA).
All other reagents were purchased at reagent grade quality.
supplemented with 10% FBS in a humidified atmosphere of air
and 5% CO2 at 37 °C. For experiments with myoblasts
only, L6 cells were seeded in medium containing 10% FBS and used at
confluence, 2 days after seeding. L6 cells were differentiated in
medium supplemented with 2% FBS into myotubes within 7 days after
seeding. Cells were serum-depleted for 3-4.5 h prior to all
experimental manipulations. Inhibitors were administered in
Me2SO, and the maximum concentration of the vehicle did not
exceed 0.05% (v/v). This concentration of vehicle was without effect
on any of the parameters measured.
-mercaptoethanol
(v/v), protease, and phosphatase inhibitors (28). Lysates were passed 5 times through a 27-gauge syringe and heated for 15 min at 65 °C.
50-µg aliquots of total protein were resolved by 10% SDS-PAGE to
detect phosphorylation of p38 MAPK, MKK3/6, or Akt by immunoblotting
using the corresponding phospho-specific antibodies at 1:500 dilutions.
Anti-p38
and anti-p38
MAPK antibodies were used at 1:1000 and
1:200 dilutions, respectively. Goat anti-rabbit IgG conjugated to
horseradish peroxidase was used as secondary antibody at a 1:7000
dilution. Proteins were detected by the Enhanced Chemiluminescence
method according to the manufacturer's instructions (PerkinElmer Life
Sciences, Boston, MA). Immunoblots were exposed to x-ray film to
produce bands within the linear range, then quantified using National Institutes of Health (NIH) Image software.
or
from 500 µg (total protein) of TX-100 detergent cell lysates containing
phosphatase and protease inhibitors was performed overnight at 4 °C
was performed as described (28). Protein concentration of the lysates
was determined by the bicinchoninic acid method. p38 MAPK
immunocomplexes were incubated for 30 min at 30 °C in 50 µl of
kinase buffer supplemented with 2 µg of ATF2 recombinant protein and
200 µM ATP per condition. Reactions were continuously
mixed on a platform shaker and were stopped by the addition of 25 µl
of 2× Laemmli sample buffer and heating for 30 min at 65 °C.
Samples were sedimented (12,000× g), and then 50 µl of
the supernatant was resolved by 10% SDS-PAGE and electrotransferred
onto polyvinylidene difluoride membranes to detect phosphorylation of
ATF2, using phospho-specific ATF-2 antibody.
-32P]ATP per condition.
Reactions were sedimented by centrifugation for 30 s. 25 µl of
the supernatants from each reaction were transferred to a Whatman P81
phosphocellulose filters and washed four times for 5 min each with 1%
phosphoric acid (v/v) and once with double-distilled water. Filters
were allowed to air dry before liquid scintillation counting (40).
RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
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Fig. 1.
Time course of insulin-stimulated
2-deoxyglucose uptake and GLUT4 translocation in L6-GLUT4myc myoblasts
and myotubes. L6-GLUT4myc myoblasts (A) and myotubes
(B) were treated for the indicated times with 100 nM insulin at 37 °C. Cell surface density of GLUT4myc
(closed squares) or 2-deoxyglucose uptake (open
circles) were measured in parallel culture plates. To avoid
skewing the glucose uptake curve to the right, the time of
insulin treatment shown does not include the 30 s required for the
uptake assay. Data points are the mean ± S.E. of five to eight
experiments performed in triplicate. Insulin-stimulated glucose uptake
and GLUT4myc translocation are expressed relative to the respective
basal values to allow for a clearer comparison between the assays
(because GLUT4myc translocation is measured in optical density units
that are normalized to the control untreated value within each
experiment). The basal rate of 2-deoxyglucose uptake in these
experiments was 20.2 ± 4.1 pmol/min/mg of protein in myoblasts
and 7.9 ± 0.2 pmol/min/mg of protein in myotubes.
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Fig. 2.
SB203580 inhibits insulin stimulation of
glucose uptake in L6-GLUT4myc myotubes but not in L6- GLUT4myc
myoblasts. L6-GLUT4myc myoblasts (A and C)
and myotubes (B and D) were pretreated with 10 µM SB203580 or vehicle alone for 20 min. Thereafter, they
were incubated for an additional 20 min with or without 100 nM insulin at 37 °C in the continued presence of
inhibitor or vehicle, as indicated. Cells were rinsed free of
inhibitors, and 2-deoxyglucose uptake (A and B)
and cell surface density of GLUT4myc proteins (C and
D) were determined as described under "Experimental
Procedures." The results are the mean ± S.E. of 4-7
experiments performed in triplicate and represent the fold change of
the response to insulin (filled columns) compared with basal
(open columns) for each culture stage. The basal rate of
2-deoxyglucose uptake in these experiments was 29.2 ± 1.9 pmol/min/mg of protein in myoblasts and 9.8 ± 1.8 in myotubes.
The asterisk in B represents a significant
difference with insulin-treated cells in the absence of SB203580,
p < 0.01.
Differential effect of SB203580 on insulin-stimulated glucose uptake in
wild-type L6 myoblasts and myotubes
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Fig. 3.
SB203580 brings the response of glucose
uptake by L6-GLUT4myc myotubes to levels seen in myoblasts.
L6-GLUT4myc myotubes were pretreated with 10 µM SB203580
(open circles) or vehicle alone (closed circles)
for 20 min followed by incubation with 100 nM insulin at
37 °C for the indicated times in the continued presence of SB203580
or vehicle alone. For comparison, L6-GLUT4myc myoblasts were incubated
with 100 nM insulin for the indicated times
(triangles). 2-Deoxyglucose uptake was measured in the
absence of inhibitors, as described in Fig. 1. The time of insulin
treatment shown does not include the time of the uptake assay (30 s).
Data points are the mean ± S.E. of four experiments performed in
triplicate. Insulin-stimulated glucose uptake is expressed relative to
the respective basal values in each culture stage.
and
(42). We monitored the initial time course of p38 phosphorylation
upon insulin treatment (100 nM) of myoblasts and myotubes,
by immunoblotting cell lysates with anti-phospho-p38 MAPK antibody.
Representative immunoblots are shown in Fig.
4A for myoblasts and Fig.
4B for myotubes. The p38 MAPK protein levels in each sample
(shown below) were detected upon subsequent immunoblotting with an
anti-pan p38 MAPK antibody to ensure equal sample loading (Fig. 4,
A and B). Quantification of five similar
experiments was performed to determine a phospho-p38:p38 protein ratio.
The results indicated that p38 MAPK phosphorylation was more responsive to insulin in myotubes (phospho-p38 MAPK was increased 2.0 ± 0.1-fold at 5 min and 2.5 ± 0.3-fold at 10 min relative to the
myotube basal levels, Fig. 4B) than myoblasts (phospho-p38
MAPK increased to a maximum of 1.4 ± 0.1-fold at 5 min,
p < 0.01, Fig. 4A). These observations are
consistent with the ineffectiveness of SB203580 on insulin-stimulated
glucose uptake in L6 myoblasts.
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Fig. 4.
Insulin-stimulated p38 MAPK phosphorylation
and activity are more robust in myotubes than in myoblasts.
L6-GLUT4myc myoblasts (A) and myotubes (B) were
treated with or without 100 nM insulin for the indicated
time periods at 37 °C. Cells were lysed in Laemmli sample buffer in
preparation for SDS-PAGE. 50-µg aliquots of total protein from each
sample were immunoblotted with either a phospho-specific p38 MAPK
antibody or an antibody that recognizes all four isoforms of p38 MAPK
as described under "Experimental Procedures." A representative
immunoblot is shown for either myoblasts or myotubes. The amount of
phosphorylated p38 MAPK or p38 MAPK protein was quantified by
densitometric scanning and analysis with NIH Image software. All values
are expressed as the ratio of phospho-p38 to p38 protein with a value
of 1 ascribed to unstimulated cells. Results are the mean ± S.E.
of four to five experiments. The asterisk and double
asterisk in parts A and B represent a
significant difference from the measurement at time zero,
p < 0.01 and p < 0.001, respectively.
L6-GLUT4myc myoblasts (C) and myotubes (D) were
treated with or without 100 nM insulin for the indicated
time periods at 37 °C. Cells were lysed with 1% TX-100 in buffer
containing phosphatase and protease inhibitors in preparation for
immunoprecipitation of p38 or p38
MAPK from 500 µg of total
protein. Enzyme activities were determined by an in vitro
kinase assay using GST-ATF-2 as substrate as described under
"Experimental Procedures." Kinase reactions were prepared for
SDS-PAGE and immunoblotted with anti-phospho-ATF-2 antibody. Shown are
representative immunoblots from the in vitro kinase
reactions. All immunoblots were quantified using densitometric scanning
and NIH Image. Results represent the mean ± S.E. of three to four
experiments performed in duplicate.
and
Activity Is Regulated
during Myogenesis--
To further examine the effect of insulin on p38
MAPK activity during myogenesis, we complemented the above results of
p38 MAPK phosphorylation with measurements of the kinase activity of
the two isoforms of the enzyme that are susceptible to inhibition by
SB203580. p38 MAPK
and
were each immunoprecipitated from basal
and insulin-treated myoblasts and myotubes, and their activity was
measured in vitro toward recombinant ATF2.
Immunoprecipitates were subsequently processed by SDS-PAGE and
immunoblotted for phospho-ATF2. Several experiments were quantified by
densitometric scanning of immunoblots like that shown in Fig. 4
(C and D). In myoblasts, the insulin-stimulated
p38 MAPK
and
activities were moderately elevated 3.3 ± 0.6-fold and 3.3 ± 0.7-fold above basal, respectively (Fig.
4C). In myotubes, insulin increased p38 MAPK
and
activities markedly by 11.1 ± 0.5-fold and 7.8 ± 2.2-fold
above basal, respectively (Fig. 4D). The activities measured
in vitro show higher changes than those observed with the
anti-phospho-p38 antibody, possibly because the anti-phospho p38 MAPK
antibody also detects other isoforms of the enzyme. Although insulin
was effective in stimulating p38 MAPK activity in both myoblasts and
myotubes, p38 MAPK responded to insulin more robustly in myotubes.
Notably, the stimulation of p38 MAPK
and
activities by
hyperosmolar stress was not different in myoblasts from that in
myotubes (results not shown).
and
isoforms of p38 MAPK measured by in vitro kinase assays
was also reduced during differentiation of L6 cells into myotubes to 43 and 58% of the activities seen in myoblasts, respectively. However,
these changes in activity were not due to a change in the expression
level of p38 MAPK (Table II).
Basal p38 MAPK and
activities are down-regulated during
L6-...GLUT4myc differentiation
or
were also immunoprecipitated from 500 µg of myoblast or myotube
cell lysates, and basal p38 MAPK activities were determined by an
in vitro kinase assay using GST-ATF-2 as substrate as
described under "Experimental Procedures." To assess changes in p38
MAPK expression, equal amounts of myoblast or myotube cell lysates were
also immunoblotted with an antibody that recognizes all four isoforms
of p38 MAPK, an antibody specific to p38 MAPK
, and an antibody
specific to p38 MAPK
. All immunoblots were quantified using
densitometric scanning and NIH Image software. The results were
calculated as the ratio of the level seen in myotubes versus
myoblasts, expressed as percent. Results represent the mean ± SE
of three to four experiments performed in duplicate.
,
, and
isoforms, whereas MKK6
activates the p38 MAPK
,
,
, and
isoforms (43). MKK3 and
MKK6 are themselves phosphorylated by upstream kinases, and this event
can be detected by immunoblotting cell lysates with an
anti-phospho-MKK3/6 antibody (this antibody does not distinguish
between MKK3 and MKK6, therefore the results are presented herein as
phosphorylation of MKK3/6). We analyzed the effect of insulin on MKK3/6
phosphorylation to explore if this pathway operates in response to the
hormone and if it shows preference in myotubes. Insulin was unable to
stimulate MKK3/6 in myoblasts within the first 10 min after its
addition (Fig. 5A). In
contrast, treatment of myoblasts with hyperosmolar sodium chloride
increased MKK3/6 phosphorylation by 5- to 10-fold (result not shown).
In these same experiments, insulin caused Akt phosphorylation in
myoblasts demonstrating the effectiveness of the hormone to activate
its receptor (result not shown). In contrast to the results obtained
with myoblasts, insulin rapidly induced phosphorylation of MKK3/6 in
myotubes within 1 min (1.8 ± 0.2-fold, p < 0.05), which declined back to basal level by 10 min (Fig.
5B). These results are consistent with the development of a
p38 MAPK-activating pathway during the differentiation of L6 muscle
cells (Fig. 4, B and D) and suggest the
possibility that insulin may activate p38 MAPK via MKK3/6. Whether
there is a causal relationship between MKK3/6 and specific p38 MAPK
isoforms in response to insulin is the subject of separate
examination.
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Fig. 5.
MKK3/6 is phosphorylated in response to
insulin only in myotubes. L6-GLUT4myc myoblasts (A) and
myotubes (B) were treated with or without insulin (100 nM) for the indicated times, and cells were lysed using
Laemmli sample buffer in preparation for SDS-PAGE. Fifty micrograms of
total protein from each sample was immunoblotted with an antibody that
recognizes equally the phosphorylated forms of MKK3 and MKK6 as
described under "Experimental Procedures." The amounts of
phosphorylated MKK3/6 were quantified using densitometric scanning and
analysis with NIH Image software. Results are the mean ± S.E. of
four to six experiments for each myoblasts and myotubes and are
expressed relative to the value in the corresponding unstimulated
cells. The asterisks in B represent a significant
difference from the measurement at time zero, p < 0.05.
,
, and
). The magnitude and activation rate of each independent isoform was
remarkably similar for myoblasts and myotubes (Fig.
6). By 5 min the activities of all three
isoforms of Akt activity were stimulated by at least 70% of their
maximum in either myoblasts or myotubes (Fig. 6). These observations
are consistent with our earlier study where IRS1-associated
phosphatidylinositol 3-kinase activity was equally stimulated by
insulin in myoblasts and myotubes (7.8-fold in myoblasts compared with
6.4-fold in myotubes (41)). Interestingly, Akt 1 and 3 were activated
~7-fold by insulin, whereas Akt2 was activated only about 2-fold
above basal levels. The robust response to insulin of the
phosphatidylinositol 3-kinase
Akt axis in myoblasts and myotubes,
contrasts with the developmental increase in p38 MAPK activation during
myogenesis.
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Fig. 6.
Insulin stimulates the activities of Akt1,
Akt2, and Akt3 in both myoblasts and myotubes. L6-GLUT4myc
myoblasts (A) and myotubes (B) were treated with
or without insulin (100 nM) for the indicated times prior
to preparation of cell lysates with 1% TX-100 in buffer containing
phosphatase and protease inhibitors in preparation for
immunoprecipitation. Each Akt isoform was immunoprecipitated from 500 µg of myoblast or myotube cell lysates using isoform-specific
antibodies. Akt activities were determined in vitro by
phosphorylation of Crosstide peptide substrate as described under
"Experimental Procedures." Results of Akt activities are the
mean ± S.E. of four to six experiments in each of myoblasts or
myotubes.
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Fig. 7.
GLUT4 translocation and Akt phosphorylation
are more sensitive to insulin than glucose uptake and p38 MAPK
phosphorylation. L6-GLUT4myc myotubes were treated with the
indicated doses of insulin for 20 min prior to determination of
GLUT4myc translocation (A, closed squares) or
glucose uptake (A, open circles) and
phosphorylation of Akt Ser473 (B, closed squares)
or p38 MAPK (B, open circles) as described under
"Experimental Procedures." The immunoblots were quantified using
densitometric scanning and analysis with NIH Image software. The
results in the main panels are expressed as the percentage of the
maximal response to facilitate comparisons among the different
parameters measured (the fold change in Akt phosphorylation is several
times higher than the fold change in p38 MAPK phosphorylation: maximal
stimulations of 11.0 ± 1.8-fold and 3.3 ± 0.5-fold,
respectively). For clarity, only the responses to 10 nM
insulin and below were illustrated. The inset in
A depicts the -fold increase in insulin-mediated GLUT4myc
translocation (closed squares) and 2-deoxyglucose uptake
(open circles) after 20 min of exposure to a full range of
insulin concentrations (up to 100 nM), expressed as -fold
stimulation above basal. Results are the mean ± S.E. of three to
four experiments.
DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
![]() |
FOOTNOTES |
---|
* This work was supported in part by the Canadian Institutes of Health Research (CIHR, Grant MT-12601).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.
Both authors contributed equally to this work.
§ Supported by a fellowship from the Canadian Pediatric Endocrinology Group and CIHR.
¶ Supported by a fellowship from The Hospital for Sick Children.
Supported by summer studentships from the Banting and Best
Diabetes Centre and the Department of Physiology from the University of Toronto.
** Supported by a studentship from CIHR.
Supported by a fellowship from the Canadian Diabetes Association.
§§ To whom correspondence should be addressed: Programme in Cell Biology, The Hospital for Sick Children, 555 University Ave., Toronto, Ontario M5G 1X8, Canada. Tel.: 416-813-6392; Fax: 416-813-5028; E-mail: amira@sickkids.ca.
Published, JBC Papers in Press, March 11, 2003, DOI 10.1074/jbc.M211136200
2 Z. Nawaz and A. Klip, unpublished observation.
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ABBREVIATIONS |
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The abbreviations used are: GLUT4, glucose transporter 4; MAPK, mitogen-activated protein kinase; FBS, fetal bovine serum; TX-100, Triton X-100; MKK, MAPK kinase; ATF2, activating transcription factor 2; ATB-BMPA, 2-N-4-(1-azi-2,2,2-trifluoroethyl)benzoyl-1,3-bis(D-mannose-4-xyloxy)-2 propylamine.
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REFERENCES |
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---|
1. | Baron, A. D., Brechtel, G., Wallace, P., and Edelman, S. V. (1988) Am. J. Physiol. 255, E769-E774[Medline] [Order article via Infotrieve] |
2. |
Shulman, G. I.
(2000)
J. Clin. Invest.
106,
171-176 |
3. | Kono, T., Robinson, F. W., Blevins, T. L., and Ezaki, O. (1982) J. Biol. Chem. 257, 10942-10947[Abstract] |
4. | Simpson, I. A., Yver, D., Hissin, P. J., Wardzala, L. J., Karnieli, E., Salans, L. B., and Cushman, S. W. (1983) Biochim. Biophys. Acta 763, 393-407[Medline] [Order article via Infotrieve] |
5. | Klip, A., Ramlal, T., Young, D. A., and Holloszy, J. O. (1987) FEBS Lett. 224, 224-230[CrossRef][Medline] [Order article via Infotrieve] |
6. | Gumà, A., Zierath, J. R., Wallberg-Henriksson, H., and Klip, A. (1995) Am. J. Physiol. 268, E613-E622[Medline] [Order article via Infotrieve] |
7. | Abel, E. D., Peroni, P., Kim, J. K., Kim, Y. B., Boss, O., Hadro, E., Minnemann, T., Shulman, G. I., and Kahn, B. B. (2001) Nature 409, 729-733[CrossRef][Medline] [Order article via Infotrieve] |
8. | Zisman, A., Peroni, O. D., Abel, E. D., Michael, M. D., Mauvais-Jarvis, F., Lowell, B. B., Wojtaszewski, J. F., Hirshman, M. F., Virkamaki, A., Goodyear, L. J., Kahn, C. R., and Kahn, B. B. (2000) Nat. Med. 6, 924-928[CrossRef][Medline] [Order article via Infotrieve] |
9. |
Rodnick, K. J.,
Slot, J. W.,
Studelska, D. R.,
Hanpeter, D. E.,
Robinson, L. J.,
Geuze, H. J.,
and James, D. E.
(1992)
J. Biol. Chem.
267,
6278-6285 |
10. |
Hirshman, M. F.,
Goodyear, L. J.,
Wardzala, L. J.,
Horton, E. D.,
and Horton, E. S.
(1990)
J. Biol. Chem.
265,
987-991 |
11. | Kahn, B. B., Simpson, I. A., and Cushman, S. W. (1988) J. Clin. Invest. 82, 691-699[Medline] [Order article via Infotrieve] |
12. | King, P. A., Horton, E. D., Hirshman, M. F., and Horton, E. S. (1992) J. Clin. Invest. 90, 1568-1575[Medline] [Order article via Infotrieve] |
13. | Goodyear, L. J., Hirshman, M. F., Smith, R. J., and Horton, E. S. (1991) Am. J. Physiol. 261, E556-E561[Medline] [Order article via Infotrieve] |
14. | Marette, A., Richardson, J. M., Ramlal, T., Balon, T. W., Vranic, M., Pessin, J. E., and Klip, A. (1992) Am. J. Physiol. 263, C443-C452[Medline] [Order article via Infotrieve] |
15. | Ferrara, C. M., and Cushman, S. W. (1999) Biochem. J. 343, 571-577[CrossRef][Medline] [Order article via Infotrieve] |
16. | Shimizu, Y., Satoh, S., Yano, H., Minokoshi, Y., Cushman, S. W., and Shimazu, T. (1998) Biochem. J. 330, 397-403[Medline] [Order article via Infotrieve] |
17. | Omatsu-Kanbe, M., Zarnowski, M. J., and Cushman, S. W. (1996) Biochem. J. 315, 25-31[Medline] [Order article via Infotrieve] |
18. | Rosholt, M. N., King, P. A., and Horton, E. S. (1994) Am. J. Physiol. 266, R95-R101[Medline] [Order article via Infotrieve] |
19. |
Ploug, T.,
van Deurs, B.,
Ai, H.,
Cushman, S. W.,
and Ralston, E.
(1998)
J. Cell Biol.
142,
1429-1446 |
20. | Kishi, K., Muromoto, N., Nakaya, Y., Miyata, I., Hagi, A., Hayashi, H., and Ebina, Y. (1998) Diabetes 47, 550-558[Abstract] |
21. | Wang, Q., Khayat, Z., Kishi, K., Ebina, Y., and Klip, A. (1998) FEBS Lett. 427, 193-197[CrossRef][Medline] [Order article via Infotrieve] |
22. | Clark, A. E., Holman, G. D., and Kozka, I. J. (1991) Biochem. J. 278, 235-241[Medline] [Order article via Infotrieve] |
23. |
Yang, J.,
and Holman, G. D.
(1993)
J. Biol. Chem.
268,
4600-4603 |
24. |
Li, D.,
Randhawa, V. K.,
Patel, N.,
Hayashi, M.,
and Klip, A.
(2001)
J. Biol. Chem.
276,
22883-22891 |
25. |
Foster, L. J.,
Li, D.,
Randhawa, V. K.,
and Klip, A.
(2001)
J. Biol. Chem.
276,
44212-44221 |
26. | Satoh, S., Gonzalez-Mulero, O. M., Clark, A. E., Kozka, I. J., Holman, G. D., and Cushman, S. W. (1991) Diabetes 40, 85A |
27. |
Satoh, S.,
Nishimura, H.,
Clark, A. E.,
Kozka, I. J.,
Vannucci, S. J.,
Simpson, I. A.,
Quon, M. J.,
Cushman, S. W.,
and Holman, G. D.
(1993)
J. Biol. Chem.
268,
17820-17829 |
28. | Somwar, R., Kim, D. Y., Sweeney, G., Huang, C., Niu, W., Lador, C., Ramlal, T., and Klip, A. (2001) Biochem. J. 359, 639-649[CrossRef][Medline] [Order article via Infotrieve] |
29. | Klip, A., Li, G., and Logan, W. J. (1984) Am. J. Physiol. 247, E494-E499 |
30. |
Huang, C.,
Somwar, R.,
Patel, N.,
Niu, W.,
Torok, D.,
and Klip, A.
(2002)
Diabetes
51,
2090-2098 |
31. | Rudich, A., Konrad, D., Török, D., Ben-Romano, R., Huang, C., Niu, W., Garg, R. R., Wijesekara, N., Germinario, R. J., Bilan, P. J., and Kilp, A. (2003) Diabetologia, in press |
32. | Murata, H., Hruz, P. W., and Mueckler, M. (2002) AIDS 16, 859-863[CrossRef][Medline] [Order article via Infotrieve] |
33. |
Hruz, P. W.,
Murata, H.,
Qiu, H.,
and Mueckler, M.
(2002)
Diabetes
51,
937-942 |
34. |
Somwar, R.,
Niu, W.,
Kim, D. Y.,
Sweeney, G.,
Ramlal, T.,
and Klip, A.
(2001)
J. Biol. Chem.
276,
46079-46087 |
35. |
Sweeney, G.,
Somwar, R.,
Ramlal, T.,
Volchuk, A.,
Ueyama, A.,
and Klip, A.
(1999)
J. Biol. Chem.
274,
10071-10078 |
36. |
Somwar, R.,
Koterski, S.,
Sweeney, G.,
Sciotti, R.,
Djuric, S.,
Berg, C.,
Trevillyan, J. M.,
Scherer, P. E.,
Rondinone, C. M.,
and Klip, A.
(2002)
J. Biol. Chem.
277,
50386-50395 |
37. | Yaffe, D. (1968) Proc. Natl. Acad. Sci. U. S. A. 61, 477-483[Medline] [Order article via Infotrieve] |
38. | Klip, A., Logan, W. J., and Li, G. (1982) Biochim. Biophys. Acta 687, 265-280[Medline] [Order article via Infotrieve] |
39. |
Tsakiridis, T.,
Vranic, M.,
and Klip, A.
(1994)
J. Biol. Chem.
269,
29934-29942 |
40. |
Wang, Q.,
Somwar, R.,
Bilan, P. J.,
Liu, Z.,
Jin, J.,
Woodgett, J. R.,
and Klip, A.
(1999)
Mol. Cell. Biol.
19,
4008-4018 |
41. | Ueyama, A., Yaworsky, K. L., Wang, Q., Ebina, Y., and Klip, A. (1999) Am. J. Physiol. 277, E572-E578[Medline] [Order article via Infotrieve] |
42. |
Kyriakis, J. M.,
and Avruch, J.
(1996)
J. Biol. Chem.
271,
24313-24316 |
43. | Ono, K., and Han, J. (2000) Cell. Signal. 12, 1-13[CrossRef][Medline] [Order article via Infotrieve] |
44. |
Ueki, K.,
Yamamoto-Honda, R.,
Kaburagi, Y.,
Yamauchi, T.,
Tobe, K.,
Burgering, B. M. T.,
Coffer, P. J.,
Komuro, I.,
Akanuma, Y.,
Yazaki, Y.,
and Kadowaki, T.
(1998)
J. Biol. Chem.
273,
5315-5322 |
45. |
Tanti, J. F.,
Grillo, S.,
Gremeaux, T.,
Coffer, P. J.,
Van Obberghen, E.,
and Le Marchand-Brustel, Y.
(1997)
Endocrinology
138,
2005-2010 |
46. |
Kohn, A. D.,
Summers, S. A.,
Birnbaum, M. J.,
and Roth, R. A.
(1996)
J. Biol. Chem.
271,
31372-31378 |
47. |
Cho, H.,
Mu, J.,
Kim, J. K.,
Thorvaldsen, J. L.,
Chu, Q.,
Crenshaw, E. B., 3rd,
Kaestner, K. H.,
Bartolomei, M. S.,
Shulman, G. I.,
and Birnbaum, M. J.
(2001)
Science
292,
1728-1731 |
48. |
Ciaraldi, T. P.,
Carter, L.,
Seipke, G.,
Mudaliar, S.,
and Henry, R. R.
(2001)
J. Clin. Endocrinol. Metab.
86,
5838-5847 |
49. | Joost, H. G., Weber, T. M., and Cushman, S. W. (1988) Biochem. J. 249, 155-161[Medline] [Order article via Infotrieve] |
50. | Hansen, P., Gulve, E., Gao, J., Schluter, J., Mueckler, M., and Holloszy, J. (1995) Am. J. Physiol. 268, C30-C35[Medline] [Order article via Infotrieve] |
51. | Wilson, C. M., and Cushman, S. W. (1994) Biochem. J. 299, 755-759[Medline] [Order article via Infotrieve] |
52. |
Hansen, P. A.,
Han, D. H.,
Marshall, B. A.,
Nolte, L. A.,
Chen, M. M.,
Mueckler, M.,
and Holloszy, J. O.
(1998)
J. Biol. Chem.
273,
26157-26163 |
53. | Zierath, J. R., Houseknecht, K. L., Gnudi, L., and Kahn, B. B. (1997) Diabetes 46, 215-223[Abstract] |
54. | Ryder, J. W., Kawano, Y., Chibalin, A. V., Rincon, J., Tsao, T. S., Stenbit, A. E., Combatsiaris, T., Yang, J., Holman, G. D., Charron, M. J., and Zierath, J. R. (1999) Biochem. J. 342, 321-328[CrossRef][Medline] [Order article via Infotrieve] |
55. | Clark, A. E., and Holman, G. D. (1990) Biochem. J. 269, 615-622[Medline] [Order article via Infotrieve] |
56. |
Harrison, S. A.,
Clancy, B. M.,
Pessino, A.,
and Czech, M. P.
(1992)
J. Biol. Chem.
267,
3783-3788 |
57. |
Shetty, M.,
Loeb, J. N.,
Vikstrom, K.,
and Ismail-Beigi, F.
(1993)
J. Biol. Chem.
268,
17225-17232 |
58. |
Barnes, K.,
Ingram, J. C.,
Porras, O. H.,
Barros, L. F.,
Hudson, E. R.,
Fryer, L. G.,
Foufelle, F.,
Carling, D.,
Hardie, D. G.,
and Baldwin, S. A.
(2002)
J. Cell Sci.
115,
2433-2442 |
59. | Klip, A., Li, G., and Walker, D. (1983) Can J. Biochem. Cell Biol. 61, 644-649[Medline] [Order article via Infotrieve] |
60. | Beguinot, F., Kahn, C. R., Moses, A. C., and Smith, R. J. (1986) Endocrinology 118, 446-455[Abstract] |
61. |
Konrad, D.,
Bilan, P. J.,
Nawaz, Z.,
Sweeney, G.,
Niu, W.,
Liu, Z.,
Antonescu, C. N.,
Rudich, A.,
and Klip, A.
(2002)
Diabetes
51,
2719-2726 |
62. | Somwar, R., Perreault, M., Kapur, S., Taha, C., Sweeney, G., Ramlal, T., Kim, D. Y., Keen, J., Cote, C. H., Klip, A., and Marette, A. (2000) Diabetes 49, 1794-1800[Abstract] |