1 Involvement of the p66Shc Protein in Glucose Transport

Articles in PresS. Am J Physiol Endocrinol Metab (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, Francesco Giorgino.
Department of Emergency and Organ Transplantation – Section on Internal Medicine,
Endocrinology and Metabolic Diseases, University of Bari, Bari, Italy.
Corresponding author:
Francesco Giorgino, M.D., Ph.D.
Department of Emergency and Organ Transplantation
Section on Internal Medicine, Endocrinology and Metabolic Diseases
University of Bari
Piazza Giulio Cesare, 11
I-70124 Bari
Italy
Phone/Fax +39 080 5478689
E-mail: [email protected]
Running head: p66Shc and glucose transport in skeletal muscle cells.
*contributed equally to the study.
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Copyright © 2008 by the American Physiological Society.
Abstract.
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 antisense-mediated
reduction (L6/p66Shcas) or adenovirus-mediated overexpression (L6/p66Shcadv) of the p66Shc
protein were examined. L6/Shcas 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 7-fold and 4-fold 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.
Key words: p66Shc, glucose transport, myoblasts, GLUT1, GLUT3, Erk.
Abbreviations: GLUT, glucose transporter; BCS, bovine calf serum; GFP, green fluorescent
protein; NP-40, Nonidet P-40.
2
Introduction.
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 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
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 are 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 pro-apoptotic signals and lifespan in
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mammals (29, 36). We 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 (33), 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 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).
In this study, we have 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 (i.) conveying MAP kinase-dependent signals to
the actin network, which ultimately affects glucose transporter trafficking, and (ii.) modulating
expression of GLUT1 and GLUT3 transporter proteins in skeletal muscle cells.
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Materials and Methods.
Cell Cultures and Antibodies.
L6 rat skeletal muscle myoblasts were cultured in MEM supplemented with 10% bovine calf
serum (BCS) (both from Invitrogen, Carlsbad, CA, USA), 2 mM L-glutamine, 100 U/ml penicillin,
100 mg/ml streptomycin, and non-essential amino acids in a 5% CO2 atmosphere at 37°C. To block
the MEK/Erk signaling pathway, cells were incubated with 20 μM PD98059 (Calbiochem, Merck
KGaA, Darmstadt, Germany) for the indicated times. To disassemble actin cytoskeleton, cells were
incubated with 2 μM cytochalasin D (Sigma, St. Louis, Missouri, USA) for 1 h.
Polyclonal GLUT1 and GLUT3 antibodies were purchased from Chemicon (Temecula, CA,
USA). Polyclonal Shc antibodies were from Transduction Laboratories (Lexington, KY, USA).
Anti-MAP kinase (Erk-1/2) antibodies were obtained from Zymed Laboratories (San Francisco,
CA, USA). Anti-phospho-p42/44 MAP kinase (Erk-1/2; Thr202/Tyr204) were obtained from New
England Biolabs (Beverly, MA, USA). Anti-GAPDH(FL-335) was from Santa-Cruz (Santa Cruz,
CA, USA).
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, USA). Efficient p66Shc overexpression was confirmed
by PCR amplification of the integrated plasmid, and by immunoblotting of total cell lysates with
Shc antibodies (33).
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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
(GFP) epitope. The resultant plasmid was linearized by digesting with restriction endonuclease Pme
I, and subsequently cotransformed into E. coli BJ5183 cells with an adenoviral backbone plasmid
pAdEasy-1. Recombinants were selected for kanamycin resistance, and recombination confirmed
by restriction endonuclease analyses. Finally, the recombinant plasmid was linearized by digesting
with restriction endonuclease Pac I and then transfected into adenovirus packaging cell lines
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. Following
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, USA), 1 μCi in 1 ml of buffer A,
to a concentration of 50 μM for 10 min at 20°C. Transport was stopped by placing the cells on ice
and rapidly washing 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 μM cytochalasin B.
Cell Fractionation.
To isolate total cellular membranes, L6 skeletal muscle cells were collected into 5 ml of
icecold HES buffer (250 mM sucrose, 1 mM EDTA, 1 mM phenylmethylsulfonyl fluoride [PMSF],
μM pepstatin, 1 μM aprotinin, 1 μM leupeptin, 20 mM HEPES, pH 7.4) and subsequently
6
homogenized with 20 strokes in a glass Dounce homogenizer (Type C; Thomas, Philadelphia, PA,
USA) at 4°C. After centrifugation at 1,000g 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 245,000g 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 μg/ml leupeptin, 2 mM sodium orthovanadate, and
1% Nonidet P-40 (NP-40). Cell lysates were then centrifuged at 12,000g 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% SDS-polyacrylamide gels. The resolved proteins
were electrophoretically transferred to nitrocellulose membranes (Hybond-ECL, Amersham Life
Science, Inc., Arlington Heights, IL, USA) using a transfer buffer containing 192 mM glycine, 20%
(vol/vol) methanol, and 0.02% SDS. To reduce non-specific binding, the membranes were
incubated in TNA buffer (10 mM Tris-HCl, pH 7.8, 0.9% NaCl, 0.01% sodium azide)
supplemented with 5% BSA and 0.05% NP-40 for 2 h at 37°C, or in PBS supplemented with 3%
non-fat 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, Inc.,
Arlington Heights, IL, USA) and quantified by densitometric analysis using Quantity One® image
analysis software (BIO-RAD Laboratories, Hercules, CA).
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Immunofluorescence Analyses.
To visualize the actin cytoskeleton, L6 cells were grown on coverslips in complete medium
in the absence or presence of 20 μM PD98059 for 72 h, and/or 2 μM cytochalasin D for 1 h, as
indicated, then 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 (TRITC)-conjugated phalloidin (from Molecular Probes,
Eugene, OR, USA) for 20 min, and subsequently washed three times with PBS. Coverslips were
mounted on glass slides with Gel mount (Biomeda, Foster City, CA, USA). 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
cover-slides and incubated in the absence or presence of 20 μM PD98059 for 72 h and/or 2 μM
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 Fluor488 anti-rabbit antibodies in PBS (1:500) for
180 min at 25°C.
Measurements of GLUT1 and GLUT3 mRNAs by qRT-PCR.
Total RNA was isolated from L6 cells using RNeasy Mini Kit (Qiagen, Hilden, Germany).
Genomic DNA contamination was eliminated by DNase digestion (Qiagen, Hilden, Germany), and
1 μg of total RNA was used for cDNA synthesis using QuantiTect Reverse Transcription Kit
(Qiagen, Hilden, Germany). GLUT1, GLUT3, and rRNA 18S primers were designed using Primer
Express 3.0 (Applied Biosystems): glut1-For 5’ GCATCGTCGTTGGGATCCT 3’; glut1-Rev 5’
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CAAGTCTGCATTGCCCATGA 3’; glut3-For 5’ GGCTCTTTTTCTGTCGGACTCTT 3’; glut3Rev 5’ AAGGATGGCAATCAGGTTGACT 3’; 18S-For 5’TGATTAAGTCCCTGCCCTTTGT 3’;
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 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 ΔΔCt method. To
confirm amplification of specific transcripts, melting curve profiles were produced at the end of
each reaction. The mRNA level of GLUT1 and GLUT3 were normalized using rRNA 18S as
internal control.
Statistical Analyses.
All data are expressed as mean±SE. Data are expressed as percentage of control values. p
values <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.
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Results.
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, 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 house-keeping
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
approximately 11-fold, in L6/p66Shcas compared to 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 following 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 pre-treatment 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
10
transport was still 4-fold 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 undergoes significant augmentation when they differentiate into
myotubes (11, 31). GLUT1 protein levels were 7-fold higher in L6/p66Shcas compared to wild-type
L6 and L6/Neo myoblasts (p<0.05; Fig. 3A). Similarly, L6/p66Shcas myoblasts showed a 4-fold
increase in GLUT3 protein content compared to control cells (p<0.05; Fig. 3B). To assess whether
the changes in glucose transporter protein abundance could result from increased gene expression,
GLUT1 and GLUT3 mRNA levels were evaluated. GLUT1 mRNA levels were 3-fold higher in
L6/p66Shcas compared to L6/Neo myoblasts (p<0.05; Fig. 3C). Similarly, L6/p66Shcas myoblasts
showed a 2-fold increase in GLUT3 mRNA levels compared to 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 presence and absence of the MEK inhibitor in both control
and L6/p66Shcas myoblasts (Fig. 4B).
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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 fill completely 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, similarly 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 appropriate
regulation of the glucose transport system in rat myoblasts, control L6 cells were studied following
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
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change in the cellular localization of both GLUT1 and GLUT3, which appeared to fill up
completely the cytoplasm (Fig. 6A), similarly 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 following
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 (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 following overexpression of p66Shc with a
specific recombinant adenoviral vector (L6/p66Shcadv myoblasts). The adenovirus-mediated p66Shc
gene transfer resulted in an 8-fold increase of p66Shc protein levels compared to non-infected 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, similarly to control
cells (Fig. 7B). In addition, both the actin cytoskeleton and basal Erk phosphorylation appeared to
be unchanged in L6 myoblasts overexpressing p66Shc as compared with control cells (Fig. 7C, and
data not shown).
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Basal glucose uptake was reduced by approximately 20% in L6/p66Shcadv as compared to
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 to 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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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, similarly 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
expression (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 down-regulated while basal Erk activity
was not apparently altered in p66Shc overexpressing myoblasts (Fig. 7; Supplementary Material, Fig.
15
1) supports the latter hypothesis. It has been recently reported 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 (45). 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 p53dependent apoptosis (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 tyrosinephosphorylated 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 FAK-p130Cas 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
subsequent promote the cytoskeletal contraction necessary for cell movement (17). In line with
these observations, up-regulation 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 up-regulation of Erk activity was found to be inappropriate for maintenance of
16
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 the actin
network and cell phenotype and was also associated with a 50% to 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 alpha-actinin-4 (7), SNAP23 (8),
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 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
17
in p66Shc protein levels, leading to a high p46Shc/p52Shc to p66Shc expression ratio, correlates with
increased proliferative activity and poor prognosis (5). Interestingly, also the abnormalities in the
glucose transport system observed in p66Shcas cells 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
modulates 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.
18
19
Acknowledgements.
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.
20
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26
Figure Legends.
Figure 1.
Effects of antisense-mediated inhibition of p66Shc protein expression on glucose transport in
L6 myoblasts. Panel 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. A representative immunoblot is shown on the top. The quantitation of p66Shc (black
bars), p52Shc (gray bars) and p46Shc (white bars) protein levels in wild-type L6, L6/Neo, and
L6/p66Shcas myoblasts in four independent experiments is shown on the bottom. Panel B. Glucose
uptake in L6, L6/Neo (open bars), and L6/p66Shcas (filled bars) myoblasts. 2-deoxy-D-glucose (2DG) uptake rates were measured in the basal state as described under Materials and Methods.
Values are mean±SE of five experiments performed in triplicate. *p<0.05 vs. L6 and L6/Neo.
Figure 2.
Effects of MEK inhibition by PD98059 on Erk phosphorylation and glucose transport in
L6/p66Shcas myoblasts. Panel A. Wild-type L6, L6/Neo, and L6/p66Shcas myoblasts were incubated
in complete medium with 20 μM 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 shown
are representative of three independent experiments. Panel B. Wild-type L6, L6/Neo, and
L6/p66Shcas myoblasts were incubated in complete medium with 20 μM PD98059 for 72 h or left
untreated. [3H]2-deoxy-D-glucose (2-DG) uptake was measured as described under Materials and
Methods. Values represent the mean±SE of five experiments performed in triplicate. #p<0.05 vs.
same cell clone not treated with PD98059; *p<0.05 vs. control cells (L6, L6/Neo) subjected to same
treatment.
27
Figure 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 (Panel A) and GLUT3 (Panel 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 represent the mean±SE of five independent experiments. *p<0.05 vs. control L6 and L6/Neo
cells. Panels 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 (Panel C) and
GLUT3 (Panel 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 four independent experiments. #p<0.05 vs. control L6/Neo cells.
Figure 4.
Effects of MEK inhibition by PD98059 on GLUT1 and GLUT3 protein levels. Total protein
content of GLUT1 (Panel A) and GLUT3 (Panel B). Cells were treated with 20 μM PD98059 in
complete medium for 72 h or left untreated. Total membranes were obtained as described under
Materials and Methods, and equal amounts of membrane protein were resolved by 10% SDS-PAGE
and subjected to immunoblotting with GLUT1 or GLUT3 antibodies, respectively. A representative
immunoblot (top) and the quantification of multiple experiments (bottom) are shown. Values
represent the mean±SE of four independent experiments. ∗p<0.05 vs. control L6 and L6/Neo cells.
Figure 5.
Effects of MEK inhibition by PD98059 on intracellular localization of GLUT1 and GLUT3.
Actin cytoskeleton and intracellular localization of GLUT1 (Panel A) and GLUT3 (Panel 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, then incubated with (+) or without
28
(-) 20 μM PD98059 for 72 h, as indicated. Cells were fixed and then incubated with GLUT1 or
GLUT3 antibodies (green), as described under Materials and Methods. TO-PRO-3 (blue) and FITCphalloidin (red) were used to visualize the nuclei and actin cytoskeleton, respectively. Images
representative of three experiments are shown.
Figure 6.
Effects of Cytochalasin D on the glucose transport system. Panel 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 mM cytochalasin D for 1 h. Cells were fixed as described under
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 are representative of three
independent experiments. Panel B. L6/p66Shcas myoblasts analyzed by indirect
immunofluorescence with antibodies to GLUT1 or GLUT3, in green, shown for comparison. TOPRO-3 (blue) was used to visualize the nuclei. Panel C. Glucose uptake in control L6 myoblasts in
the presence or absence of cytochalasin D. Cells were treated with or without 2 μM cytochalasin D
in complete medium for 1 h, and 2-deoxy-D-glucose (2-DG) uptake was measured as described
under Materials and Methods. Values represent the mean±SE of three experiments performed in
triplicate. *p<0.05 vs. untreated cells. Panel 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 μM PD98059 for 72 h and/or 2 μM cytochalasin
D for 1 h. 2-deoxy-Dglucose uptake rates were then measured as described under Materials and
Methods. #p<0.05 vs. same cell clone not treated with PD98059; +p<0.05 vs. same cell clone treated
with PD98059.
29
Figure 7.
Effects of adenovirus-mediated overexpression of p66Shc on the glucose transport system in L6
myoblasts. Panel A. Overexpression of p66Shc in L6 myoblasts. Total cell lysates from wild-type L6
myoblasts (white bars), L6 myoblasts infected with the empty vector (L6/Mock, gray bar), and L6
myoblasts infected with the p66Shc adenovirus (L6/p66Shcadv, black bar) were analyzed by
immunoblotting with Shc antibodies. A representative immunoblot (left) and the quantitation of
multiple experiments (right) are shown. Data represent the mean±SE of four independent
experiments. ∗p<0.05 vs. control (L6, L6/Mock) cells. Panel B. Morphology of confluent control
L6/Mock and L6/p66Shcadv myoblasts under light microscopy. Magnification, x 200. Panel 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
under Materials and Methods. The GFP signal in green identifies the infected cells. Panel D. [3H]2deoxy-D-glucose (2-DG) uptake in L6 (white bars), L6/Mock (gray bars) and L6/p66Shcadv
myoblasts (black bars). 2-DG uptake was measured as described under Materials and Methods.
Values represent the mean±SE of three independent experiments performed in triplicate. Panels E
and F. Total cellular content of GLUT1 and GLUT3 glucose transporters. Wild-type L6 (white bar),
L6/Mock (gray bar), and L6/p66Shcadv (black bars) myoblasts were grown in complete medium to
∼90% confluence. Then, total membranes were obtained and analyzed by immunoblotting with
GLUT1 (Panel E) or GLUT3 (Panel F) antibodies, as indicated. A representative immunoblot (left)
and the quantification of three independent experiments (right) are shown. ∗p<0.05 vs. control (L6
and L6/Mock) cells.
30
A
GAPDH
p66
p52
p46
L6
N1
N5
C6
L6/Neo
D27
D28
L6/p66Shcas
Shhc protein content (% of L6)
160
140
p66
p52
p46
120
100
80
60
40
*
20
*
*
0
L6
N1
N5
C6
L6/Neo
D27
D28
L6/p66Shcas
pmol 2-DG uptake x mgg prot-1 x min-1
B
2000
*
1500
1000
*
*
D27
D28
500
0
L6
N1
N5
L6/Neo
C6
L6/p66Shcas
A
p Erk 1
p-Erk-1
p-Erk-2
Erk-1
Erk-2
-
PD98059
+
-
+
L6
-
N1
+
-
N5
+
-
+
C6
L6/Neo
D28
L6/p66Shcas
B
pmol 22-DG uptake x mg prot1 x min-1
2000
*
1500
*
1000
*
*
#
*
#
500
0
PD98059
*
#
- +
- +
- +
- +
- +
- +
L6
Neo1
Neo5
C6
D27
D28
L6/Neo
L6/p66Shcas
A
B
GLUT1
N1
N5
900
C6
D27
*
*
D28
*
600
300
0
L6
N1
N5
L6/Neo
C6
D27
D28
L6
GLLUT3 protein content
(% of L6)
GLLUT1 protein content
(% of L6)
L6
GLUT3
N5
C6
D27
D28
600
*
*
*
D27
D28
300
0
L6
L6/p66Shcas
N1
N5
L6/Neo
C6
L6/p66Shcas
D
12
10
8
6
4
2
0
GLUT1 mRNA
#
L6/Neo
L6/p66
p Shcas
GLUT3/18S mRNA
A
GLUT1/18S mRNA
A
C
N1
3.0
25
2.5
2.0
1.5
1.0
0.5
0.0
GLUT3 mRNA
#
L6/Neo
L6/p66
p Shcas
A
*
PD98059 -
+
-
L6
+
-
N1
+
N5
-
+
C6
+
D28
L6/p66Shcas
L6/Neo
900
GLUT1 protrein levels
(% of L6)
-
GLUT1
*
*
*
*
600
300
0
-
PD98059
-
+
L6
+
N1
-
+
N5
-
+
C6
- +
D28
B
GLUT3
PD98059
-
+
-
L6
+
-
N1
+
-
C6
+
D28
L6/p66Shcas
L6/Neo
GLUT3 protein levels
(% of L6/Neo)
450
*
*
300
*
*
150
0
PD98059
-
+
L6
-
+
N1
-
+
C6
- +
D28
A
GLUT1
L6/Neo (N1)
Phalloidin
Merge
GLUT1
L6/p66Shcas (D28)
Phalloidin
Merge
-
-
47.62 μm
47.62 μm
+
+
47.62 μm
47.62 μm
B
GLUT3
L6/Neo (N1)
Phalloidin
Merge
GLUT3
L6/p66Shcas (D28)
Phalloidin
Merge
47.62 μm
47.62 μm
+
47.62 μm
+
47.62 μm
A
B
L6 + Cytochalasin D
Phalloidin
GLUT1
Merge
L6/p66Shcas
GLUT1
m
Phalloidin
GLUT3
Merge
GLUT3
m
D
L6
30000
pmol 2-DG uptake x mg prot-1 x min-1
pmol 2-DG uptake x mg prot-1 x min-1
C
25000
20000
15000
10000
5000
0
Cytochalasin D
-
+
L6/p66Shcas
140000
120000
100000
80000
60000
40000
20000
0
Cytochalasin D
PD98059
-
+
-
+
+
+
p66Shc protein levels
(%
% of L6)
A
p66
p52
p46
L6
B
L6/Mock
L6/Mock
L6/
p66Shcadv
1200
1000
800
600
400
200
0
*
L6
C
L6/p66Shcadv
L6/Mock
L6/Mock
L6/
p66Shcadv
L6/p66Shcadv
47.62μm
47.62μm
pm
mol 2-DG uptake x mg prrot-1 x min-1
D
120
100
*
80
60
40
20
0
L6
L6/Mock L6/p66Shcadv
GLUT1
L6
L6/Mock
L6/p66Shcadv
GLUT1 protein levels
(%
% of L6)
E
140
120
100
80
60
40
20
0
GLUT3
L6
L6/Mock
L6/p66Shcadv
GLUT3 protein levels
(% of L6))
F
140
120
100
80
60
40
20
0
*
L6
L6/Mock L6/p66Shcadv
*
L6
L6/Mock L6/p66Shcadv