Involvement of the p66Shc protein in glucose transport

Transcription

Am J Physiol Endocrinol Metab 296: E228–E237, 2009.
First published October 28, 2008; doi:10.1152/ajpendo.90347.2008.
Involvement of the p66Shc protein in glucose transport regulation in skeletal
muscle myoblasts
Annalisa Natalicchio,* Francesca De Stefano,* Sebastio Perrini, Luigi Laviola, Angelo Cignarelli,
Cristina Caccioppoli, Anna Quagliara, Mariangela Melchiorre, Anna Leonardini, Antonella Conserva,
and Francesco Giorgino
Department of Emergency and Organ Transplantation, Section on Internal Medicine, Endocrinology and Metabolic Diseases,
University of Bari, Bari, Italy
Submitted 27 June 2008; accepted in final form 15 October 2008
glucose transporter 1; glucose transporter 3; extracellular signal related
kinase
SKELETAL MUSCLE IS A MAJOR site of glucose disposal in vivo, and
this function is tightly regulated to guarantee efficient glucose
uptake and metabolism upon environment-driven changes in
energy demand. At the cellular level, transport of glucose
across the plasma membrane is the first rate-limiting step for
glucose metabolism and is mediated by facilitative glucose
transporter (GLUT) proteins, both in skeletal muscle fibers and
in cultured skeletal muscle cells (9, 32). The glucose transport
system has been extensively characterized in the rat L6 skeletal
muscle cell line (41, 50). While fully differentiated myotubes
express mainly GLUT1 and GLUT4, accounting for basal and
hormone-stimulated glucose uptake, respectively (11, 31), L6
myoblasts express mainly GLUT1, which is ubiquitous, and
* A. Natalicchio and F. De Stefano contributed equally to this work.
Address for reprint requests and other correspondence: F. Giorgino, Dept. of
Emergency and Organ Transplantation, Section on Internal Medicine, Endocrinology and Metabolic Diseases, Univ. of Bari, Piazza Giulio Cesare, 11,
I-70124 Bari, Italy (e-mail: [email protected]).
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GLUT3, which is typically found in fetal (15) and regenerating
muscle (11), as well as in neuronal cells (25). In the absence of
hormonal stimulation, in L6 myoblasts, GLUT1 is uniformly
localized to both the plasma membrane and an intracellular
pool of specialized vesicles, whereas GLUT3 is mainly intracellular (11). Regulation of the cellular content and localization
of GLUT1 and/or GLUT3, leading to increased glucose transport activity, represents an adaptive response to the increased
energy demand associated with various external stimuli, including exposure to insulin or IGF-I (19, 20, 32), hypoxia (2,
54), and oxidative stress (14, 21, 40). Even though distinct
protein members of the MAP kinase family were found to
affect the expression of various GLUT proteins in different
experimental conditions (13, 46, 53), the signaling events
regulating the glucose transport system in myoblasts remain to
be completely clarified.
The p66Shc protein is one of the three isoforms encoded by
the mammalian Shc locus (38). All three isoforms can be
tyrosine phosphorylated upon growth factor stimulation; however, p46/p42Shc is coupled to growth and survival signals,
whereas p66Shc also undergoes serine phosphorylation and
inhibits activation of the MEK/MAP kinase pathway triggered
by growth factor receptors (30, 33, 34). In addition, p66Shc has
been proposed as a cellular mediator of oxidative stress, and
this response appears to regulate proapoptotic signals and life
span in mammals (29, 36). We (33) have shown that selective
reduction of p66Shc in L6 myoblasts resulted in constitutive
activation of the MEK/MAP kinase signaling pathway, leading
to complete disruption of the actin network, which is critical
for the maintenance of cell shape and intracellular trafficking.
Indeed, both MAP kinase activity and the actin network regulate the glucose transport system. Transfection of constitutively active mutants of MEK into 3T3-L1 adipocytes resulted in increased basal glucose transport activity associated with increased expression of GLUT1 mRNA and
protein (53); furthermore, IGF-I-induced glucose transport
in retinal endothelial cells requires activation of MAP kinase (6). On the other hand, several studies have shown that
cellular glucose transport in insulin-sensitive cells relies on
the integrity of the actin and microtubule cytoskeleton.
Treatment of L6 myotubes, 3T3-L1 adipocytes, or rat adipocytes with specific cytoskeleton disrupters, such as cytochalasin D or latrunculin A, resulted in marked inhibition
of insulin-mediated GLUT4 translocation (35, 49, 51).
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0193-1849/09 $8.00 Copyright © 2009 the American Physiological Society
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Natalicchio A, De Stefano F, Perrini S, Laviola L, Cignarelli
A, Caccioppoli C, Quagliara A, Melchiorre M, Leonardini A,
Conserva A, Giorgino F. Involvement of the p66Shc protein in
glucose transport regulation in skeletal muscle myoblasts. Am J
Physiol Endocrinol Metab 296: E228 –E237, 2009. First published
October 28, 2008; doi:10.1152/ajpendo.90347.2008.—The p66Shc
protein isoform regulates MAP kinase activity and the actin cytoskeleton turnover, which are both required for normal glucose transport
responses. To investigate the role of p66Shc in glucose transport
regulation in skeletal muscle cells, L6 myoblasts with antisensemediated reduction (L6/p66Shcas) or adenovirus-mediated overexpression (L6/p66Shcadv) of the p66Shc protein were examined. L6/
Shc
as myoblasts showed constitutive activation of ERK-1/2 and disruption of the actin network, associated with an 11-fold increase in
basal glucose transport. GLUT1 and GLUT3 transporter proteins were
sevenfold and fourfold more abundant, respectively, and were localized throughout the cytoplasm. Conversely, in L6 myoblasts overexpressing p66Shc, basal glucose uptake rates were reduced by 30% in
parallel with a ⬃50% reduction in total GLUT1 and GLUT3 transporter levels. Inhibition of the increased ERK-1/2 activity with
PD98059 in L6/Shcas cells had a minimal effect on increased GLUT1
and GLUT3 protein levels, but restored the actin cytoskeleton, and
reduced the abnormally high basal glucose uptake by 70%. In conclusion, p66Shc appears to regulate the glucose transport system in
skeletal muscle myoblasts by controlling, via MAP kinase, the integrity of the actin cytoskeleton and by modulating cellular expression of
GLUT1 and GLUT3 transporter proteins via ERK-independent pathways.
P66SHC
AND GLUCOSE TRANSPORT IN SKELETAL MUSCLE CELLS
In this study, we investigated the role of p66Shc on glucose
uptake and glucose transporter expression and localization in
L6 myoblasts. We show that p66Shc plays an important role in
the regulation of cellular glucose uptake by 1) conveying MAP
kinase-dependent signals to the actin network, which ultimately affects glucose transporter trafficking; and 2) modulating expression of GLUT1 and GLUT3 transporter proteins in
skeletal muscle cells.
MATERIALS AND METHODS
AJP-Endocrinol Metab • VOL
enized with 20 strokes in a glass Dounce homogenizer (Type C;
Thomas, Philadelphia, PA) at 4°C. After centrifugation at 1,000 g for
3 min at 4°C to remove large cell debris and unbroken cells, the
supernatant from this first spin was separated from the pellet and then
centrifuged at 245,000 g for 90 min at 4°C to yield a pellet of total
cellular membranes (31, 50).
Immunoblotting. For preparing total cell lysates, L6 skeletal muscle
cells were washed with Ca2⫹/Mg2⫹-free PBS and then mechanically
detached in ice-cold lysis buffer containing 50 mM HEPES (pH 7.5),
150 mM NaCl, 1 mM MgCl2, 1 mM CaCl2, 10% glycerol, 10 mM
sodium pyrophosphate, 10 mM sodium fluoride, 2 mM EDTA, 2 mM
PMSF, 5 ␮g/ml leupeptin, 2 mM sodium orthovanadate, and 1%
Nonidet P-40. Cell lysates were then centrifuged at 12,000 g for 10
min, and the resulting supernatant was collected and assayed for
protein concentration using the Bradford dye binding assay kit with
BSA as a standard. For immunoblotting studies, equal amounts of
cellular proteins were resolved by electrophoresis on 10% SDSpolyacrylamide gels. The resolved proteins were electrophoretically
transferred to nitrocellulose membranes (Hybond-ECL; Amersham
Life Science, Arlington Heights, IL) using a transfer buffer containing
192 mM glycine, 20% (vol/vol) methanol, and 0.02% SDS. To reduce
nonspecific binding, the membranes were incubated in TNA buffer
(10 mM Tris 䡠 HCl, pH 7.8, 0.9% NaCl, and 0.01% sodium azide)
supplemented with 5% BSA and 0.05% Nonidet P-40 for 2 h at 37°C,
or in PBS supplemented with 3% nonfat dry milk for 2 h at room
temperature, as appropriate, and then incubated overnight at 4°C with
the indicated antibodies. The proteins were visualized by enhanced
chemiluminescence using horseradish peroxidase-labeled anti-rabbit
or anti-mouse IgG (Amersham Life Science) and quantified by densitometric analysis using Quantity One image analysis software (BioRad Laboratories, Hercules, CA).
Immunofluorescence analyses. To visualize the actin cytoskeleton,
L6 cells were grown on coverslips in complete medium in the absence
or presence of 20 ␮M PD98059 for 72 h and/or 2 ␮M cytochalasin D
for 1 h, as indicated, and then they were fixed with 4% paraformaldehyde and permeabilized with PBS supplemented with 0.1% Triton
X-100. Fixed cells were incubated with FITC-conjugated phalloidin
Oregon Green 514 or tetramethyl rhodamine isothiocyanate-conjugated phalloidin (Molecular Probes, Eugene, OR) for 20 min and
subsequently washed three times with PBS. Coverslips were mounted
on glass slides with gel mount (Biomeda, Foster City, CA). Pictures
were acquired on a Leica TCS SP2 laser scanning spectral confocal
microscope (Leica Microsystems, Heerbrugg, Switzerland), and all
images were taken at the same magnification.
To study the intracellular localization of GLUT1 and GLUT3, L6
cells were grown on glass coverslides and incubated in the absence or
presence of 20 ␮M PD98059 for 72 h and/or 2 ␮M cytochalasin D for
1 h, as indicated. Cells were fixed with 4% paraformaldehyde,
permeabilized with PBS supplemented with 0.1% Triton X-100 for 15
min, and then incubated with polyclonal GLUT1 or GLUT3 antibodies in PBS-3% BSA (1:100) overnight at 4°C. Cells were then washed
three times with PBS and incubated with Alexa Fluor 488 anti-rabbit
antibodies in PBS (1:500) for 180 min at 25°C.
Measurements of GLUT1 and GLUT3 mRNAs by quantitative
RT-PCR. Total RNA was isolated from L6 cells using RNeasy mini
kit (Qiagen, Hilden, Germany). Genomic DNA contamination was
eliminated by DNase digestion (Qiagen), and 1 ␮g of total RNA was
used for cDNA synthesis using QuantiTect reverse transcription kit
(Qiagen). GLUT1, GLUT3, and rRNA 18S primers were designed
using Primer Express 3.0 (Applied Biosystems) as follows: glut1For: 5⬘-GCATCGTCGTTGGGATCCT-3⬘; glut1-Rev: 5⬘CAAGTCTGCATTGCCCATGA-3⬘; glut3-For: 5⬘-GGCTCTTTTTCTGTCGGACTCTT-3⬘; glut3-Rev: 5⬘-AAGGATGGCAATCAGGTTGACT-3⬘; 18S-For: 5⬘-TGATTAAGTCCCTGCCCTTTGT-3⬘;
and 18S-Rev: 5⬘-GATCCGAGGGCCTCACTAAAC-3⬘. The PCR
reactions were carried out in an ABI PRISM 7500 System (Applied
Biosystems) under the following conditions: 50°C for 2 min, 95°C for
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Cell cultures and antibodies. L6 rat skeletal muscle myoblasts were
cultured in MEM supplemented with 10% BCS (both from Invitrogen,
Carlsbad, CA), 2 mM L-glutamine, 100 U/ml penicillin, 100 mg/ml
streptomycin, and nonessential amino acids in a 5% CO2 atmosphere
at 37°C. To block the MEK/ERK signaling pathway, cells were
incubated with 20 ␮M PD98059 (Calbiochem, Merck, Darmstadt,
Germany) for the indicated times. To disassemble actin cytoskeleton,
cells were incubated with 2 ␮M cytochalasin D (Sigma, St. Louis,
MO) for 1 h.
Polyclonal GLUT1 and GLUT3 antibodies were purchased from
Chemicon (Temecula, CA). Polyclonal Shc antibodies were from
Transduction Laboratories (Lexington, KY). Anti-MAP kinase (ERK1/2) antibodies were obtained from Zymed Laboratories (San Francisco, CA). Anti-phospho-p42/44 MAP kinase (ERK-1/2; Thr202/
Tyr204) was obtained from New England Biolabs (Beverly, MA).
Anti-GAPDH(FL-335) was from Santa Cruz Biotechnology (Santa
Cruz, CA).
Transfection studies. L6 myoblasts stably expressing reduced levels of the p66Shc protein isoform were generated as previously
described (33). Briefly, plasmids containing p66Shc-specific oligonucleotides in antisense orientation were transfected into L6 myoblasts
by liposome-mediated gene transfer using Lipofectamine (Invitrogen,
Carlsbad, CA). Efficient p66Shc overexpression was confirmed by
PCR amplification of the integrated plasmid and by immunoblotting
of total cell lysates with Shc antibodies (33).
To generate L6 myoblasts expressing augmented levels of the
p66Shc protein, the gene of interest was first cloned into a shuttle
vector pAdTrack-CMV containing a green fluorescent protein epitope.
The resultant plasmid was linearized by digesting with restriction
endonuclease PmeI and subsequently was cotransformed into Escherichia coli BJ5183 cells with an adenoviral backbone plasmid
pAdEasy-1. Recombinants were selected for kanamycin resistance,
and recombination was confirmed by restriction endonuclease analyses. Finally, the recombinant plasmid was linearized by digesting with
restriction endonuclease PacI and then transfected into the adenovirus
packaging cell line QBI293A. Scalar doses of adenovirus-containing
culture medium were used to biologically define the optimal infection
dose (⬎90% of infected cells) for L6 myoblasts.
Glucose transport assay. Transport activity was determined in cells
cultured in 35-mm diameter wells. After incubation in serum-free
MEM for 16 h, cells were washed twice with 2 ml of buffer A (140
mM NaCl, 2.5 mM MgCl2, 1 mM CaCl2, 5 mM KCl, and 20 mM
HEPES, pH 7.4) at 37°C, and transport was started by adding
2-[3H]deoxy-D-glucose (NEN, Boston, MA), 1 ␮Ci in 1 ml of buffer
A, to a concentration of 50 ␮M for 10 min at 20°C. Transport was
stopped by placing the cells on ice and rapidly washing them three
times with ice-cold buffer A. Cells were lysed in 1 ml 0.05 N NaOH
for 45 min. Aliquots of this lysate were used for liquid scintillation
counting and determination of protein content. Nonspecific transport
was determined by performing the assay in the presence of 10 ␮M
cytochalasin B.
Cell fractionation. To isolate total cellular membranes, L6 skeletal
muscle cells were collected into 5 ml of ice-cold HES buffer (250 mM
sucrose, 1 mM EDTA, 1 mM PMSF, 1 ␮M pepstatin, 1 ␮M aprotinin,
1 ␮M leupeptin, 20 mM HEPES, pH 7.4) and subsequently homog-
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P66SHC
10 min, 40 cycles at 95°C for 15 s, and 60°C for 1 min. Relative RNA
levels were determined by analyzing the changes in SYBR green
fluorescence during PCR using the ⌬⌬CT method. To confirm amplification of specific transcripts, melting curve profiles were produced at
the end of each reaction. The mRNA levels of GLUT1 and GLUT3
were normalized using rRNA 18S as an internal control.
Statistical analyses. All data are expressed as means ⫾ SE. Data
are expressed as percentage of control values. P values of ⬍0.05 were
considered to represent statistical significance. Statistical analyses
were performed by unpaired Student’s t-tests or one-way ANOVA
tests, as appropriate.
RESULTS
Fig. 1. Effects of antisense-mediated inhibition of p66Shc protein expression
on glucose transport in L6 myoblasts. A: protein levels of Shc isoforms in
wild-type L6 myoblasts, L6 myoblasts transfected with the empty vector
(L6/Neo), and L6 myoblasts transfected with the p66Shc antisense (L6/
p66Shcas). Total cell lysates were analyzed by immunoblotting with Shc and
GAPDH antibodies. Top: representative immunoblot. Bottom: quantitation of
p66Shc (filled bars), p52Shc (gray bars), and p46Shc (open bars) protein levels in
wild-type L6, L6/Neo, and L6/p66Shcas myoblasts in 4 independent experiments. B: glucose uptake in L6 and L6/Neo (open bars) and L6/p66Shcas (filled
bars) myoblasts. 2-[3H]deoxy-D-glucose (2-DG) uptake rates were measured in
the basal state as described in MATERIALS AND METHODS. Values are means ⫾
SE of 5 experiments performed in triplicate. *P ⬍ 0.05 vs. L6 and L6/Neo.
levels were sevenfold higher in L6/p66Shcas compared with
wild-type L6 and L6/Neo myoblasts (P ⬍ 0.05; Fig. 3A).
Similarly, L6/p66Shcas myoblasts showed a fourfold increase
in GLUT3 protein content compared with control cells (P ⬍
0.05; Fig. 3B). To assess whether the changes in glucose
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Glucose transport in myoblasts with reduced p66Shc. To
investigate the potential role of p66Shc in the regulation of the
glucose transport system, basal glucose uptake was analyzed in
three independent clones of L6 myoblasts with antisensemediated reduction of p66Shc protein levels (L6/p66Shcas myoblasts, clones C6, D27, and D28; Fig. 1A). In the L6/p66Shcas
myoblasts, p66Shc protein levels were decreased by ⬃90%
compared with untransfected wild-type L6 myoblasts or L6/
Neo myoblasts transfected with the empty pCR3.1 vector
(L6/Neo and clones N1 and N5; P ⬍ 0.05), while the protein
levels of the other Shc isoforms, i.e., p52Shc and p46Shc, were
not significantly altered (Fig. 1A). Expression levels of the
housekeeping protein GAPDH were also similar in L6, L6/
Neo, and L6/p66Shcas myoblasts (Fig. 1A). Noteworthy, basal
2-[3H]deoxy-D-glucose uptake was found to be markedly
higher, by ⬃11-fold, in L6/p66Shcas compared with untransfected L6 and L6/Neo cells (P ⬍ 0.05; Fig. 1B).
The p66Shc protein exerts an inhibitory effect on the MEK/
MAP kinase signaling pathway, and thus L6 myoblasts with
selective reduction of p66Shc show markedly increased basal
levels of ERK-1/2 phosphorylation (33). To assess whether the
constitutive activation of MEK/MAP kinase could contribute
to the observed glucose transport changes in L6/p66Shcas
myoblasts, basal glucose uptake was assessed after exposure of
cells to the MEK inhibitor PD98059 for 72 h. The elevated
levels of basal ERK-1/2 phosphorylation, observed in L6/
p66Shcas myoblasts, were markedly reduced by PD98059, and
thus after treatment with this compound, ERK-1/2 phosphorylation was similarly low in L6/p66Shcas and control myoblasts
(Fig. 2A). In control cells, basal glucose uptake was not
affected by pretreatment of cells with the MEK inhibitor (Fig.
2B). By contrast, in L6/p66Shcas myoblasts, treatment with
PD98059 resulted in a 50% reduction of the high basal glucose
transport (P ⬍ 0.05 vs. untreated L6/p66Shcas cells; Fig. 2B);
however, glucose transport was still fourfold higher in
PD98059-treated L6/p66Shcas myoblasts compared with control cells (P ⬍ 0.05; Fig. 2B).
Glucose transporters in myoblasts with reduced p66Shc. To
investigate whether the enhanced basal glucose transport in
L6/Shcas myoblasts could be explained by p66Shc-related
changes in the amounts of glucose transporters, the protein
levels of GLUT1 and GLUT3, the two predominant glucose
transporter protein isoforms expressed in L6 myoblasts, were
measured in total cellular membranes by immunoblotting with
GLUT1 or GLUT3 antibodies, respectively. GLUT4 was not
analyzed because its expression is very low in the myoblast
stage of L6 cells, and it undergoes significant augmentation
when they differentiate into myotubes (11, 31). GLUT1 protein
P66SHC
p66Shcas
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p66Shcas
transporter protein abundance could result from increased
gene expression, GLUT1 and GLUT3 mRNA levels were
evaluated. GLUT1 mRNA levels were threefold higher in
L6/p66Shcas compared with L6/Neo myoblasts (P ⬍ 0.05;
Fig. 3C). Similarly, L6/p66Shcas myoblasts showed a twofold increase in GLUT3 mRNA levels compared with control cells (P ⬍ 0.05; Fig. 3D). Whether the constitutive
activation of MEK/MAP kinase could contribute to the
increased glucose transporter protein levels in L6/p66Shcas
myoblasts was investigated next. However, treatment with
PD98059 had no effect on GLUT1 protein levels in control
cells and did not significantly reduce GLUT1 in L6/p66Shcas
myoblasts (679 vs. 764% of control; P ⫽ 0.3; Fig. 4A).
Similarly, GLUT3 protein levels were comparable in the
Fig. 3. Effects of antisense-mediated inhibition of p66Shc protein expression on GLUT1
and GLUT3 glucose transporters. Representative immunoblots (top) and quantification
from multiple experiments (bottom) of the protein content of GLUT1 (A) and GLUT3 (B)
transporters in total membranes from wild-type
L6, L6/Neo (clones N1 and N5), and L6/
p66Shcas (clones C6, D27, and D28) myoblasts.
Values are means ⫾ SE of 5 independent experiments. *P ⬍ 0.05 vs. control L6 and L6/Neo
cells. C and D: GLUT1 and GLUT3 mRNA
expression levels. Total RNA was extracted
from L6/Neo and L6/p66Shcas myoblasts, and
mRNA expression levels of GLUT1 (C) and
GLUT3 (D) were determined by quantitative
real-time RT-PCR. The mRNA level was normalized for each target gene against 18S ribosomal RNA as internal control. Values are
means ⫾ SE of cells from 4 independent experiments. #P ⬍ 0.05 vs. control L6/Neo cells.
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Fig. 2. Effects of MEK inhibition by PD98059 on ERK phosphorylation and glucose transport in L6/p66Shcas myoblasts. A: wildtype L6, L6/Neo, and L6/p66Shcas myoblasts were incubated in
complete medium with 20 ␮M PD98059 for 72 h or left untreated.
Cells lysates were analyzed by immunoblotting with phospho-p42/
p44 MAP kinase (Thr202/Tyr204) or MAP kinase antibodies to
study ERK-1/2 phosphorylation (top) and total protein content
(bottom), respectively. Data represent 3 independent experiments.
B: wild-type L6, L6/Neo, and L6/p66Shcas myoblasts were incubated in complete medium with 20 ␮M PD98059 for 72 h or left
untreated. 2-DG uptake was measured as described in MATERIALS
AND METHODS. Values are means ⫾ SE of 5 experiments performed in triplicate. *P ⬍ 0.05 vs. control cells (L6, L6/Neo)
subjected to same treatment; #P ⬍ 0.05 vs. same cell clone not
treated with PD98059.
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P66SHC
presence and absence of the MEK inhibitor in both control
and L6/p66Shcas myoblasts (Fig. 4B).
Integrity of the actin cytoskeleton and regulation of the
glucose transport system. Constitutive activation of ERK-1/2
in the basal state in L6/p66Shcas myoblasts results in abnormalities of the cell morphology due to disruption of actin fibers
and cytoskeleton (33), and this could potentially affect the
intracellular distribution of glucose transporters and overall
glucose transport activity. In control myoblasts, in which actin
fibers could be easily detected (Fig. 5, A and B), GLUT1
appeared to be uniformly distributed in the perinuclear region
(Fig. 5A). By contrast, L6/p66Shcas myoblasts showed complete disassembly of the actin network (Fig. 5, A and B) and
large amounts of cellular GLUT1, which appeared to completely fill the reduced cell cytoplasm (Fig. 5A). Similarly, the
GLUT3 signal appeared to be localized in the perinuclear
region in control cells (Fig. 5B), and it showed an intense
localization throughout the cytosol of the L6/p66Shcas myoblasts (Fig. 5B). Inhibition of the MEK/MAP kinase pathway
with PD98059 resulted in restoration of the actin cytoskeleton
in the L6/p66Shcas myoblasts (Fig. 5, A and B); this was
associated with partial relocalization of GLUT1 and GLUT3
transporters, which appeared to be distributed in the perinuclear region and more diffusely along the actin filaments,
similar to the control myoblasts. Therefore, in L6 myoblasts
with reduced p66Shc levels, the increased basal glucose uptake
was associated with increased protein content as well as disruption of the actin cytoskeleton, possibly resulting in altered
cellular localization of GLUT1 and GLUT3 glucose transporters. Inhibition of the constitutive activation of ERK-1/2 in the
L6/p66Shcas myoblasts did not significantly reduce the abnormally elevated levels of GLUT1 and GLUT3 glucose transporters but partially corrected the increase in glucose transport,
likely through reorganization of the actin network.
To verify whether the integrity of the actin cytoskeleton is
indeed essential for the appropriate regulation of the glucose
transport system in rat myoblasts, control L6 cells were studied
after incubation with the actin-disrupting agent cytochalasin D,
which provoked a marked disorganization of the actin network
(Fig. 6A). Treatment with cytochalasin D also induced a
change in the cellular localization of both GLUT1 and GLUT3,
which appeared to completely fill up the cytoplasm (Fig. 6A),
similar to what was observed in the L6/p66Shcas myoblasts
(Fig. 6B), and an augmentation of basal glucose transport rates
by 1.5-fold (P ⬍ 0.05; Fig. 6C). Thus inhibition of actin
polymerization partially replicated the effects of p66Shc reduction on the glucose transport system. To further confirm the
role of the actin cytoskeleton in glucose transport regulation in
the L6/p66Shcas myoblasts, glucose uptake was also assessed
in these cells after exposure to either the MEK inhibitor
PD98059, cytochalasin D, or both of these compounds (Fig.
6D). Glucose uptake was reduced by 50% after ERK inhibition
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Fig. 4. Effects of MEK inhibition by PD98059 on GLUT1 and
GLUT3 protein levels. Total protein content of GLUT1 (A) and
GLUT3 (B). Cells were treated with 20 ␮M PD98059 in complete medium for 72 h or left untreated. Total membranes were
obtained as described in MATERIALS AND METHODS, and equal
amounts of membrane protein were resolved by 10% SDSPAGE and subjected to immunoblotting with GLUT1 or GLUT3
antibodies, respectively. A representative immunoblot (top) and
the quantification of multiple experiments (bottom) are shown.
Values are means ⫾ SE of 4 independent experiments. *P ⬍
0.05 vs. control L6 and L6/Neo cells.
P66SHC
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(P ⬍ 0.05) but remained unaltered when cells were incubated
in the presence of cytochalasin D (Fig. 6D; P ⫽ 0.35 vs.
untreated L6/p66Shcas myoblasts). Importantly, the addition of
cytochalasin D to PD98059-treated cells resulted in a significant increase in glucose transport rates (P ⬍ 0.05), which were
not different than those of untreated cells (Fig. 6D). Therefore,
the integrity of the actin cytoskeleton is essential for the
regulation of the glucose transport system, and changes in
glucose transport rates in myoblasts with reduced p66Shc levels
occur, at least in part, via ERK-mediated disassembly of the
cytoskeleton integrity.
Glucose transport system in myoblasts with increased
p66Shc. To confirm the involvement of p66Shc in the regulation
of the glucose transport system in rat skeletal muscle cells, L6
myoblasts were examined after overexpression of p66Shc with
a specific recombinant adenoviral vector (L6/p66Shcadv myoblasts). The adenovirus-mediated p66Shc gene transfer resulted
in an eightfold increase of p66Shc protein levels compared with
noninfected or mock-infected control cells (P ⬍ 0.05; Fig. 7A).
L6/p66Shcadv myoblasts showed normal cellular morphology
and appeared to grow as a typical monolayer of elongated cells,
similar to control cells (Fig. 7B). In addition, both the actin
cytoskeleton and basal ERK phosphorylation appeared to be
unchanged in L6 myoblasts overexpressing p66Shc compared
with control cells (Fig. 7C, and data not shown).
Basal glucose uptake was reduced by ⬃20% in L6/
p66Shcadv compared with control noninfected and mock-infected L6 myoblasts (P ⬍ 0.05; Fig. 7D). To evaluate whether
the reduced glucose transport rates in L6 myoblasts with
augmented p66Shc levels were associated with altered glucose
transporter proteins, the protein levels of GLUT1 and GLUT3
transporters were measured in total cellular membranes by
immunoblotting with specific antibodies. GLUT1 protein
was found to be significantly reduced in L6/p66Shcadv
compared with control myoblasts (P ⬍ 0.05; Fig. 7E). In
addition, GLUT3 protein levels were markedly lower than
control in p66Shc-overexpressing cells (P ⬍ 0.05; Fig. 7F).
Therefore, in L6 myoblasts, overexpression of p66Shc results
in changes in the glucose transport system, including lower
basal transport and decreased protein levels of GLUT1 and
GLUT3 transporters, that are opposite to those observed
when p66Shc is knocked down.
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Fig. 5. Effects of MEK inhibition by PD98059 on intracellular localization of GLUT1 and GLUT3. Actin cytoskeleton and intracellular localization of GLUT1
(A) and GLUT3 (B) analyzed by indirect immunofluorescence. Control L6/Neo (N1) and L6/p66Shcas (D28) cells were grown on glass slides in complete medium
to ⬃80% confluence and then incubated with (⫹) or without (⫺) 20 ␮M PD98059 for 72 h, as indicated. Cells were fixed and then incubated with GLUT1 or
GLUT3 antibodies (green), as described in MATERIALS AND METHODS. TO-PRO-3 (blue) and FITC- phalloidin (red) were used to visualize the nuclei and actin
cytoskeleton, respectively. Images represent 3 experiments.
E234
P66SHC
DISCUSSION
The p66Shc protein isoform has been shown to regulate
cellular apoptosis and survival (29, 36), the cytoskeleton architecture (33), and cellular responses to oxidative stress (29,
36). In this study, we provide evidence for an important role of
p66Shc in the regulation of glucose uptake in rat skeletal muscle
cells. Antisense-mediated reduction of p66Shc protein content
in L6 myoblasts resulted in markedly increased glucose transport (Fig. 1). Conversely, in p66Shc-overexpressing L6 cells,
basal glucose uptake was reduced (Fig. 7). These data, for the
first time, establish a link between p66Shc and glucose metabolism in insulin-sensitive cells.
The absolute cellular levels of glucose transporters are
critical determinants of the efficiency of the glucose transport
system in L6 myoblasts (18). In previous work (39, 53),
overexpression of GLUT1 in both L6 myoblasts and 3T3-L1
adipocytes was associated with markedly increased basal glucose transport rates, similar to the results of this study. GLUT1
and GLUT3 expression was found to be markedly increased in
cells with selective reduction of p66Shc (Fig. 3) and significantly decreased in p66Shc-overexpressing myoblasts (Fig. 7).
This directly relates to changes in basal glucose transport,
which was increased in L6/p66Shcas (Fig. 1) and decreased in
L6/p66Shcadv cells (Fig. 7). Signaling through the MEK/MAP
kinase pathway has been shown to control GLUT gene expresAJP-Endocrinol Metab • VOL
sion (53), and L6/p66Shcas myoblasts have constitutive activation of the MEK/ERK pathway (33; this study). However,
inhibition of the elevated levels of ERK-1 and ERK-2 phosphorylation in L6/p66Shcas myoblasts did not significantly
modify GLUT1 or GLUT3 proteins levels (Fig. 4). It is
possible that MEK inhibition achieved with PD98059 for 72 h
was still not sufficient to restore ERK to normal activity levels
and/or to allow degradation of excess transporter proteins,
which are characterized by relatively long half-lives (16, 42).
However, these results suggest that other p66Shc-related but
ERK-independent signals may be required for the regulation of
GLUT gene expression. The observation that GLUT1 and
GLUT3 transporters were downregulated while basal ERK
activity was not apparently altered in p66Shc-overexpressing
myoblasts (Fig. 7; Supplemental Figs. 1 and 2; supplemental
data for this article are available online at the Am J Physiol
Endocrinol Metab website) supports the latter hypothesis. It
has been recently reported (45) that GLUT1 and GLUT4
promoters are inhibited by the tumor suppressor protein p53
and that point mutations in p53 either partially or completely
abolish its inhibitory effects on GLUT1 and GLUT4 gene
expression. This may explain the increase in glucose transporter expression associated with certain varieties of cancer
characterized by loss of p53 function (45). Since ablation of
p66Shc is associated with impaired p53- dependent apoptosis
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Fig. 6. Effects of cytochalasin D on the glucose transport system. A: intracellular localization of GLUT1 and GLUT3. Control L6 myoblasts were grown on glass slides, arrested at
80% confluence and incubated in the presence
of 2 ␮M cytochalasin D for 1 h. Cells were
fixed as described in MATERIALS AND METHODS
and analyzed by indirect immunofluorescence
with antibodies to GLUT1 (top) or GLUT3
(bottom), as indicated in green. TO-PRO-3
(blue) and FITC-phalloidin (red) were used to
visualize the nuclei and actin cytoskeleton,
respectively. Images represent 3 independent
experiments. B: L6/p66Shcas myoblasts analyzed by indirect immunofluorescence with antibodies to GLUT1 or GLUT3, in green, shown
for comparison. TO-PRO-3 (blue) was used to
visualize the nuclei. C: glucose uptake in control L6 myoblasts in the presence or absence of
cytochalasin D. Cells were treated with or
without 2 ␮M cytochalasin D in complete medium for 1 h, and 2-DG uptake was measured
as described in MATERIALS AND METHODS. Values are means ⫾ SE of 3 experiments performed in triplicate. *P ⬍ 0.05 vs. untreated
cells. D: glucose uptake in L6 myoblasts with
reduction of p66Shc (L6/p66Shcas myoblasts,
results from clones C6 and D28 pooled together). Cells were incubated in complete medium with or without 20 ␮M PD98059 for 72 h
and/or 2 ␮M cytochalasin D for 1 h. 2-DG
uptake rates were then measured as described
in MATERIALS AND METHODS. #P ⬍ 0.05 vs.
same cell clone not treated with PD98059;
⫹P ⬍ 0.05 vs. same cell clone treated with
PD98059.
P66SHC
E235
(47), it will be important to investigate whether the p66Shc
protein may affect glucose transporter expression via p53
signaling. Other intracellular kinases, such as p70S6 kinase and
p38 MAP kinase, have been also implicated in the regulation of
glucose transporter expression in L6 myoblasts (13, 46).
Whether these other signaling components may be modulated
by p66Shc has not been yet determined.
In the L6 myoblasts with reduced p66Shc, constitutive activation of MEK/MAP kinase signaling results in disorganization of the actin cytoskeleton (33; this study) and altered
subcellular localization of GLUT1 and GLUT3 (Fig. 5). The
Shc proteins have been shown to promote cytoskeleton rearrangement in response to growth factor stimulation through the
ERK pathway in multiple cell types (10, 22). In addition, the
p46Shc and p52Shc isoforms bind to and are tyrosine phosphorylated by integrin-activated tyrosine kinases, such as FAK and
Src (43, 44), and similarly promote ERK activation. Thus
growth factor- and integrin-triggered signals converge on the
Shc/ERK pathway and dynamically regulate actin fiber assembly, together with signals transduced through the FAKp130Cas complex. The link between ERK and the cytoskeleton
is also demonstrated by the finding that activated ERK can
directly phosphorylate and activate myosin light chains and
subsequently promote the cytoskeletal contraction necessary
for cell movement (17). In line with these observations, upregulation of MEK activity in rat kidney cells causes disruption
of the actin cytoskeleton through MEK-dependent inhibition of
the Rho-ROCK-LIM kinase pathway, which promotes actin
stress fiber stabilization and actomyosin-based cell contractility
(37). Similarly, in this study, persistent upregulation of ERK
activity was found to be inappropriate for maintenance of
normal organization of the actin cytoskeleton in L6/p66Shcas
myoblasts; conversely, inhibition of the abnormally elevated
MEK/ERK activity with the MEK-inhibitor PD98059 restored
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Fig. 7. Effects of adenovirus-mediated overexpression of p66Shc on the glucose transport
system in L6 myoblasts. A: overexpression of
p66Shc in L6 myoblasts. Total cell lysates from
wild-type L6 myoblasts (open bar), L6 myoblasts infected with the empty vector (L6/
Mock, gray bar), and L6 myoblasts infected
with the p66Shc adenovirus (L6/p66Shcadv,
filled bar) were analyzed by immunoblotting
with Shc antibodies. A representative immunoblot (left) and the quantitation of multiple
experiments (right) are shown. Data are
means ⫾ SE of 4 independent experiments.
*P ⬍ 0.05 vs. control (L6, L6/Mock) cells.
B: morphology of confluent control L6/Mock
and L6/p66Shcadv myoblasts under light microscopy. Magnification: ⫻200. C: actin cytoskeleton of control L6/Mock and L6/p66Shcadv myoblasts. Cells were stained with TO-PRO-3 (blue)
and phalloidin (red) to visualize the nuclei and
actin cytoskeleton, respectively, as described in
MATERIALS AND METHODS. Green fluorescent protein (GFP) signal in green identifies the infected cells. D: 2-DG uptake in L6 (open bar),
L6/Mock (gray bar), and L6/p66Shcadv myoblasts (filled bar). 2-DG uptake was measured
as described in MATERIALS AND METHODS. Values are means ⫾ SE of 3 independent experiments performed in triplicate. E and F: total
cellular content of GLUT1 and GLUT3 glucose transporters. Wild-type L6 (open bars),
L6/Mock (gray bars), and L6/p66Shcadv (filled
bars) myoblasts were grown in complete medium to ⬃90% confluence. Then, total membranes were obtained and analyzed by immunoblotting with GLUT1 (E) or GLUT3 (F)
antibodies, as indicated. A representative immunoblot (left) and the quantification of 3
independent experiments (right) are shown.
*P ⬍ 0.05 vs. control (L6 and L6/Mock) cells.
E236
P66SHC
ulates the cellular levels of GLUT1 and GLUT3, and this also
contributes to the changes in glucose uptake. Thus the p66Shc
may represent an effector of glucose transport in skeletal
muscle cells and prove to play an important role in the adaptive
responses to environmental factors.
GRANTS
This work was supported by grants from the Ministero dell’Università e
Ricerca (Italy), the Cofinlab 2000 –Centro di Eccellenza “Genomica comparata: geni coinvolti in processi fisiopatologici in campo biomedico e agrario”
(Italy), and an educational grant from Pfizer Italia srl (ARADO Program) to
F. Giorgino.
REFERENCES
1. Barres R, Gremeaux T, Gual P, Gonzales T, Gugenheim J, Tran A, Le
Marchand-Brustel Y, Tanti J-F. Enigma interacts with adaptor protein
with PH and SH2 domains to control insulin-induced actin cytoskeleton
remodeling and glucose transporter 4 translocation. Mol Endocrinol 20:
2864 –2875, 2006.
2. Bashan N, Burdett E, Hundal HS, Klip A. Regulation of glucose
transport and GLUT1 glucose transporter expression by O2 in muscle cells
in culture. Am J Physiol Cell Physiol 262: C682–C690, 1992.
3. Boado RJ, Black KL, Pardridge WM. Gene expression of GLUT3 and
GLUT1 glucose transporters in human brain tumors. Brain Res Mol Brain
Res 27: 51–57, 1994.
4. Cormont M, Le Marchand-Brustel Y. The role of small G-proteins in
the regulation of glucose transport. Mol Membr Biol 18: 213–220, 2001.
5. Davol PA, Bagdasaryan R, Elfenbein GJ, Maizel AL, Frackelton AR
Jr. Shc proteins are strong, independent prognostic markers for both
node-negative and node-positive primary breast cancer. Cancer Res 63:
6772– 6783, 2003.
6. DeBosch BJ, Deo BK, Kumagai AK. Insulin-like growth factor-1 effects
on bovine retinal endothelial cell glucose transport: role of MAP kinase.
J Neurochem 81: 728 –734, 2002.
7. Foster LJ, Rudich A, Talior I, Patel N, Furtado LM, Bilan PJ, Mann
M, Klip A. Insulin dependent interactions of proteins with GLUT4
revealed through stable isotope labeling by amino acids in cell culture
(SILAC). J Proteome Res 5: 64 –75, 2006.
8. Foster LJ, Yaworsky K, Trimble SW, Klip A. SNAP23 promotes
insulin-dependent glucose uptake in 3T3-L1 adipocytes: possible interaction with cytoskeleton. Am J Physiol Cell Physiol 276: C1108 –C1114,
1999.
9. Furler SM, Jenkins AB, Storlien LH, Kraegen EW. In vivo location of
the rate-limiting step of hexose uptake in muscle and brain tissue of rats.
Am J Physiol Endocrinol Metab 261: E337–E347, 1991.
10. Giancotti FG, Ruoslahti E. Integrin signaling. Science 285: 1028 –1032,
1999.
11. Guillet-Deniau I, Leturque A, Girard J. Expression and cellular localization of glucose transporters (GLUT1, GLUT3, GLUT4) during differentiation of myogenic cells isolated from rat foetuses. J Cell Sci 107:
487– 496, 1994.
12. Hatanaka M. Transport of sugars in tumor cell membranes. Biochim
Biophys Acta 355: 77–104, 1974.
13. Ho RC, Alcazar O, Fujii N, Hirshman MF, Goodyear LJ. p38␥ MAPK
regulation of glucose transporter expression and glucose uptake in L6
myotubes and mouse skeletal muscle. Am J Physiol Regul Integr Comp
Physiol 286: R342–R349, 2004.
14. Katz A. Modulation of glucose transport in skeletal muscle by reactive
oxygen species. J Appl Physiol 102: 1671–1676, 2007.
15. Kayano T, Fukumoto H, Eddy RL, Fan YS, Byers MG, Shows TB,
Bell GI. Evidence for a family of human glucose transporter-like proteins.
Sequence and gene localization of a protein expressed in fetal skeletal
muscle and other tissues. J Biol Chem 263: 12245–12248, 1988.
16. Khayat ZA, McCall AL, Klip A. Unique mechanism of GLUT3 glucose
transporter regulation by prolonged energy demand: increased protein
half-life. Biochem J 333: 713–718, 1998.
17. Klemke RL, Cai S, Giannini AL, Gallagher PJ, de Lanerolle P,
Cheresh DA. Regulation of cell motility by mitogen-activated protein
kinase. J Cell Biol 137: 481– 492, 1997.
18. Klip A, Pâquet MR. Glucose transport and glucose transporters in muscle
and their metabolic regulation. Diabetes Care 13: 228 –243, 1990.
296 • FEBRUARY 2009 •
www.ajpendo.org
the actin network and cell phenotype and was also associated
with a 50 –75% reduction of the abnormally high basal glucose
uptake (Fig. 2). This may be explained by the fact that an intact
actin cytoskeleton is necessary for intracellular localization and
appropriate plasma membrane insertion of glucose transporters
(23, 48). Multiple proteins have been reported to interact with
glucose transporters and the actin fibers, providing the molecular scaffold for cytoskeleton-regulated glucose transporter
localization and trafficking; these include ␣-actinin-4 (7),
SNAP23 (8), and Enigma (1) and small GTPases such as Ras,
Rad, Rho, Arf, and Rab isoforms (4).
Indeed, the results of this study suggest that basal glucose
uptake in rat myoblasts is more strictly dependent on the
integrity of the actin network than on the absolute levels of
glucose transporter proteins. In control L6 myoblasts, disruption of the cytoskeleton with cytochalasin D was associated
with abnormal intracellular localization of GLUT1 and GLUT3
and significantly increased glucose transport rates (Fig. 6). In
cells with reduced p66Shc levels treated with the MEK inhibitor, restoration of the actin cytoskeleton markedly reduced the
high glucose uptake (Fig. 2), whereas if the actin network was
maintained in a disrupted state by simultaneous incubation
with cytocalasin D, the increased glucose uptake rates remained elevated (Fig. 6). Conversely, in cells with increased
cellular levels of p66Shc, the actin network did not appear to be
perturbed and changes in glucose transport rates were rather
modest (Fig. 7). L6 myoblasts with reduction of p66Shc and
overactivation of MEK/ERK signaling exhibited a transformed
phenotype, with rounded shape; complete disruption of the
actin stress fibers, focal adhesions, and cell cytoskeleton (33);
and resistance to stress-induced apoptosis (Natalicchio A. and
Giorgino F., unpublished results). Importantly, in human breast
cancer tissues, a relative reduction in p66Shc protein levels,
leading to a high p46Shc/p52Shc-to-p66Shc expression ratio,
correlates with increased proliferative activity and poor prognosis (5). Interestingly, the abnormalities in the glucose transport system observed in p66Shcas cells also evoke some features of cancer cells. Tumorigenesis is associated with enhanced cellular uptake and metabolism of glucose, which is
required for adaptation to higher energy supply (12). Increased
glucose transport in transformed cells has been associated with
increased and deregulated expression of glucose transporter
proteins, mainly GLUT1 and/or GLUT3 (3, 26 –28, 52). In
addition, oncogenic transformation of cultured mammalian
cells causes a rapid increase of glucose transport and GLUT1
expression, due to specific effects involving GLUT1 promoter
enhancer elements (24). Induction of the GLUT1 isoform is
therefore instrumental in cellular adaptation to stress conditions, including cancer, in which energy requirements are
increased (21). In future work, it will be important to
determine whether changes in cellular p66Shc levels may
contribute to the increase in glucose transporter expression
and glucose uptake observed in transformed cells.
In conclusion, the p66Shc protein exerts a constitutive inhibitory effect on ERK-1/2 activity, which is required for appropriate regulation of actin cytoskeleton turnover controlling the
normal cellular localization of GLUT1 and GLUT3 in skeletal
muscle myoblasts. Loss of p66Shc results in constitutive activation of ERK-1/2, leading to altered cell cytoskeleton and
profound modifications of the cellular glucose uptake capacity.
In addition, via other yet unidentified pathways, p66Shc mod-
P66SHC
37. Pawlak G, Helfman DM. MEK mediates v-Src-induced disruption of the
actin cytoskeleton via inactivation of the Rho-ROCK-LIM kinase pathway. J Biol Chem 277: 26927–26933, 2002.
38. Pelicci G, Lanfrancone L, Grignani F, McGlade J, Cavallo F, Forni G,
Nicoletti I, Grignani F, Pawson T, Pelicci PG. A novel transforming
protein (SHC) with an SH2 domain is implicated in mitogenic signal
transduction. Cell 70: 93–104, 1992.
39. Robinson R, Robinson LJ, James DE, Lawrence JC Jr. Glucose
transport in L6 myoblasts overexpressing GLUT1 and GLUT4. J Biol
Chem 268: 22119 –22126, 1993.
40. Rudich A, Kozlovsky N, Potashnik R, Bashan N. Oxidant stress reduces
insulin responsiveness in 3T3-L1 adipocytes. Am J Physiol Endocrinol
Metab 272: E935–E940, 1997.
41. Sargeant R, Mitsumoto Y, Sarabia V, Shillabeer G, Klip A. Hormonal
regulation of glucose transporters in muscle cells in culture. J Endocrinol
Invest 16: 147–162, 1993.
42. Sargeant RJ, Pâquet MR. Effect of insulin on the rates of synthesis and
degradation of GLUT1 and GLUT4 glucose transporters in 3T3-L1 adipocytes. Biochem J 290: 913–919, 1993.
43. Schlaepfer DD, Hauck CR, Sieg DJ. Signaling through focal adhesion
kinase. Prog Biophys Mol Biol 71: 435– 478, 1999.
44. Schlaepfer DD, Hunter T. Focal adhesion kinase overexpression enhances ras-dependent integrin signaling to ERK2/mitogen-activated protein kinase through interactions with and activation of c-Src. J Biol Chem
272: 13189 –13195, 1997.
45. Schwartzenberg-Bar-Yoseph F, Armoni M, Karnieli E. The tumor
suppressor p53 downregulates glucose transporters GLUT1 and GLUT4
gene expression. Cancer Res 64: 2627–2633, 2004.
46. Taha C, Tsakiridis T, McCall A, Klip A. Glucose transporter expression
in L6 muscle cells: regulation through insulin- and stress-activated pathways. Am J Physiol Endocrinol Metab 273: E68 –E76, 1997.
47. Trinei M, Giorgio M, Cicalese A, Barozzi S, Ventura A, Migliaccio E,
Milia E, Padura IM, Raker VA, Maccarana M, Petronilli V, Minucci
S, Bernardi P, Lanfrancone L, Pelicci PG. A p53-p66Shc signalling
pathway controls intracellular redox status, levels of oxidationdamaged
DNA and oxidative stress-induced apoptosis. Oncogene 21: 3872–3878,
2002.
48. Tsakiridis T, Tong P, Matthews B, Tsiani E, Bilan PJ, Klip A, Downey
GP. Role of the actin cytoskeleton in insulin action. Microsc Res Tech 47:
79 –92, 1999.
49. Tsakiridis T, Vranic M, Klip A. Disassembly of the actin network
inhibits insulin-dependent stimulation of glucose transport and prevents
recruitment of glucose transporters to the plasma membrane. J Biol Chem
269: 29934 –29942, 1994.
50. Walker PS, Ramlal T, Sarabia V, Koivisto UM, Bilan PJ, Pessin JE,
Klip A. Glucose transport activity in L6 muscle cells is regulated by the
coordinate control of subcellular glucose transporter distribution, biosynthesis, and mRNA transcription. J Biol Chem 265: 1516 –1523, 1990.
51. Wang Q, Bilan PJ, Tsakiridis T, Hinek A, Klip A. Actin filament
participate in the relocalization of phosphatidyl 3-kinase to glucose transporter-containing compartments and in the stimulation of glucose uptake
in 3T3-L1 adipocytes. Biochem J 331:917–928, 1998.
52. Yamamoto T, Seino Y, Fukumoto H, Koh G, Yano H, Inagaki N,
Yamada Y, Inoue K, Manabe T, Imura H. Over-expression of facilitative glucose transporter genes in human cancer. Biochem Biophys Res
Commun 16: 223–230, 1990.
53. Yamamoto Y, Yoshimasa Y, Koh M, Suga J, Masuzaki H, Ogawa Y,
Hosoda K, Nishimura H, Watanabe Y, Inoue G, Nakao K. Constitutively active mitogen-activated protein kinase kinase increases GLUT1
expressions and recruits both GLUT1 and GLUT4 at the cell surface in
3T3-L1 adipocytes. Diabetes 49: 332–339, 2000.
54. Zhang JZ, Behrooz A, Ismail-Beigi F. Regulation of glucose transport
by hypoxia. Am J Kidney Dis 34: 189 –202, 1999.
296 • FEBRUARY 2009 •
www.ajpendo.org
19. Klip A, Ramlal T, Bilan PJ, Marette A, Liu Z, Mitsumoto Y. What
signals are involved in the stimulation of glucose transport by insulin in
muscle cells? Cell Signal 5: 519 –529, 1993.
20. Koivisto UM, Martinez-Valdez H, Bilan PJ, Burdett E, Ramlal T, Klip
A. Differential regulation of the GLUT-1 and GLUT-4 glucose transport
systems by glucose and insulin in L6 muscle cells in culture. J Biol Chem
266: 2615–2621, 1991.
21. Kozlovsky N, Rudich A, Potashnik R, Bashan N. Reactive oxygen
species activate glucose transport in L6 myotubes. Free Radic Biol Med
23: 859 – 869, 1997.
22. Lai KM, Pawson T. The ShcA phosphotyrosine docking protein sensitizes cardiovascular signaling in the mouse embryo. Genes Dev 14:
1132–1145, 2000.
23. Liu Z, Zhang YW, Chang YS, Fang FD. The role of cytoskeleton in
glucose regulation. Biochemistry 71: 476 – 480, 2006.
24. Macheda ML, Rogers S, Best JD. Molecular and cellular regulation of
glucose transporter (GLUT) proteins in cancer. J Cell Physiol 202:
654 – 662, 2005.
25. Maher F, Vannucci S, Takeda J, Simpson IA. Expression of mouseGLUT3 and human-GLUT3 glucose transporter proteins in brain. Biochem Biophys Res Commun 182: 703–711, 1992.
26. Matsuzu K, Segade F, Matsuzu U, Carter A, Bowden DW, Perrier
ND. Differential expression of glucose transporters in normal and pathologic thyroid tissue. Thyroid 14: 806 – 812, 2004.
27. Medina RA, Owen GI. Glucose transporters: expression, regulation and
cancer. Biol Res 35: 9 –26, 2002.
28. Meneses Am Medina RA, Kato S, Pinto M, Jaque MP, Lizama I,
Garcia Mde L, Nualart F, Owen GI. Regulation of GLUT3 and glucose
uptake by the cAMP signalling pathway in the breast cancer cell line
ZR-75. J Cell Physiol 214: 110 –116, 2008.
29. Migliaccio E, Giorgio M, Mele S, Pelicci G, Reboldi P, Pandolfi PP,
Lanfrancone L, Pelicci PG. The p66shc adaptor protein controls oxidative stress response and life span in mammals. Nature 402: 309 –313,
1999.
30. Migliaccio E, Mele S, Salcini AE, Pelicci G, Venus Lai KM, SupertiFurga G, Pawson T, Di Fiore PP, Lanfrancone L, Pelicci PG. Opposite
effects of the p52Shc/p46Shc and p66Shc splicing isoforms on the EGF
receptor-MAP kinase-fos signaling pathway. EMBO J 16: 706 –716, 1997.
31. Mitsumoto Y, Klip A. Development regulation of the subcellular distribution and glycosylation of GLUT1 and GLUT4 glucose transporters
during myogenesis of L6 muscle cells. J Biol Chem 267: 4957– 4962,
1992.
32. Mueckler M. Family of glucose-transporter genes. Implications for glucose homeostasis and diabetes. Diabetes 39: 6 –11, 1990.
33. Natalicchio A, Laviola L, De Tullio C, Renna LA, Montrone C,
Perrini S, Valenti G, Procino G, Svelto M, Giorgino F. Role of the
p66Shc isoform in insulin-like growth factor I receptor signaling through
MEK/Erk and regulation of actin cytoskeleton in rat myoblasts. J Biol
Chem 279: 43900 – 43909, 2004.
34. Okada S, Kao AW, Ceresa BP, Blaikie P, Margolis B, Pessin JE. The
66-kDa Shc isoform is a negative regulator of the epidermal growth
factor-stimulated mitogen-activated protein kinase pathway. J Biol Chem
272: 28042–28049, 1997.
35. Omata W, Shibata H, Li L, Takata K, Kojima I. Actin filaments play
a critical role in insulin-induced exocytolytic recruitment but not in
endocytosis of GLUT4 in isolated rat adipocyres. Biochem J 346: 321–
328, 2000.
36. Pacini S, Pellegrini M, Migliaccio E, Patrussi L, Ulivieri C, Ventura A,
Carraro F, Naldini A, Lanfrancone L, Pelicci PG, Baldari CT.
P66SHC promotes apoptosis and antagonizes mitogenic signaling in T
cells. Mol Cell Biol 24: 1747–1757, 2004.
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Involvement of the p66Shc protein in glucose transport

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