(Received for publication, December 16, 1996, and in revised form, February 20, 1997)
From the Departament de Bioquimica i Biologia Molecular, Facultat de Biologia, Universitat de Barcelona, Avda. Diagonal 645, 08028 Barcelona, Spain
Muscle cell differentiation caused a reduction of
glucose transport, GLUT1 glucose transporter expression, and GLUT1
mRNA levels. A fragment of 2.1 kilobases of the rat GLUT1 gene
linked to chloramphenicol acetyltransferase drove transcriptional
activity in myoblasts, and differentiation caused a decrease in
transcription. Transient transfection of 5 and 3
deletion constructs
showed that the fragment
99/
33 of the GLUT1 gene drives
transcriptional activity of the GLUT1 gene and participates in the
reduced transcription after muscle differentiation. Electrophoretic
mobility shift assays showed the binding of Sp1 protein to the fragment
102/
37 in the myoblast state but not in myotubes, and Sp1 was found
to transactivate the GLUT1 promoter. Western blot analysis indicated
that Sp1 was drastically down-regulated during myogenesis. Furthermore,
the forced over-expression of MyoD in C3H10T1/2 cells mimicked the effects observed during myogenesis, Sp1 down-regulation and reduced transcriptional activity of the GLUT1 gene promoter.
In all, these data suggest a regulatory model in which MyoD activation during myogenesis causes the down-regulation of Sp1, which contributes to the repression of GLUT1 gene transcription and, therefore, leads to the reduction in GLUT1 expression and glucose transport.
The formation of skeletal muscle during embryogenesis involves, first, commitment of mesodermal stem cells to the myogenic lineage. Myoblast cells, although undifferentiated and capable of continued proliferation, differentiate when they receive the appropriate environmental signals, fuse, and form multinucleate myotubes. At the same time as this morphological differentiation, a battery of adult muscle-specific genes whose products are required for the unique contractile and metabolic properties of the muscle fiber are activated (1, 2). The factors that regulate the expression of muscle-specific genes following commitment to terminal differentiation are well established. The best characterized are the members of the myogenic basic helix-loop-helix (bHLH)1 protein family or MyoD family, that function as master regulators of muscle cell fate during development (2, 3). Four members of the family have been cloned: MyoD (4), Myf5 (5), myogenin (6, 7), and Mrf4 (8-10). Each of these factors is expressed exclusively in skeletal muscle, and when expressed ectopically in a variety of non-muscle cell types, they activate the complete program of myogenic differentiation (2). All the members of the myogenic bHLH family activate the transcription of muscle-specific genes by binding to the E-box consensus sequence (CANNTG) in muscle gene promoters and enhancers. However, not all muscle genes contain functional E boxes in their regulatory promoter regions, and myogenic bHLH proteins can also activate transcription of muscle-specific genes that lack E boxes in their control regions (3). As expected, from these data, other muscle-specific transcription factors have been described to function as intermediates in the activation of gene expression during myogenesis, such as the M-CAT binding factor (11), and the myocyte enhancer factor 2 (MEF2) (12, 13).
Myogenesis is also associated with down-regulation of several
growth-regulated myoblast proteins, including c-Fos (14), - and
-actins (15, 16), and the differentiation inhibitor (Id) (17). In
contrast to the wealth of information regarding the mechanisms that
activate genes participating in the myotube phenotype, relatively
little is known regarding regulatory sequences or factors involved in
the control of the repression of the muscle embryo genes during muscle
cell differentiation. Thus, it has been described that an activating
transcription factor site is required for the expression of the Id2A
gene in muscle cells, and that the binding of nuclear factors to the
activating transcription factor site is decreased during myogenic
differentiation (18).
Glucose transporter expression is developmentally regulated in skeletal muscle (19, 20). Thus, during fetal and early postnatal life, GLUT1 is highly expressed in heart and skeletal muscles. Postnatal life is characterized by GLUT1 repression in muscle, which is concomitant with the induction of GLUT4 expression (19). Similarly, it has been reported that myogenesis leads to induction of GLUT4 expression and repression of GLUT1 expression (21, 22). Based on the fact that congenital hypothyroidism partially blocks GLUT1 repression associated with neonatal life (23) and that denervation up-regulates GLUT1 in skeletal muscle (20, 24-26), it is likely that thyroid hormones and muscle innervation play a role in the regulation of muscle GLUT1 expression in vivo. However, the detailed mechanisms that contribute to GLUT1 repression during perinatal development and myogenesis are largely unknown. Here we have examined the mechanisms that repress GLUT1 expression during myogenesis.
125I-labeled protein A and
[-32P]-dCTP were purchased from ICN.
2-Deoxy-D-[3H]glucose was obtained from
DuPont NEN. Hybond N was from Amersham Corp., and random primed DNA
labeling kit was from Boehringer Mannheim. Immobilon was obtained from
Millipore Corp.
-Globulin and most commonly used chemicals were from
Sigma. Dulbecco's modified Eagle's medium (DMEM), fetal-bovine serum,
glutamine, and antibiotics were obtained from Whittaker (Walkersville,
MD). L6E9 rat skeletal muscle cell line was kindly provided by Dr. B. Nadal-Ginard (Harvard University). C3H10T1/2 mouse cells stably
transfected with MyoD were obtained from Dr. V. Andrés (St.
Elizabeth's Medical Center, Boston).
The plasmid containing the 2106/+134 region of the rat GLUT1 genomic
sequence was obtained from Dr. M. Birnbaum (University of
Pennsylvania). pCAT-basic vector was obtained from Promega (Madison,
WI). pCMV-
-galactosidase vector was obtained from Dr. N. Brand
(National Heart & Lung Institute, London). Plasmid CMV-Sp1 was a
generous gift of Dr. R. Tjian (University of California, Berkeley).
Rat skeletal muscle L6E9 myoblasts were grown in monolayer culture in DMEM supplemented with 10% (v/v) fetal bovine serum, 1% (v/v) antibiotics (10,000 units/ml penicillin G and 10 mg/ml streptomycin), 2 mM glutamine, and 25 mM Hepes, pH 7.4. Confluent myoblasts were differentiated by lowering fetal bovine serum to a final concentration of 2% (v/v). Mouse C3H10T1/2 fibroblasts stably transfected with MyoD were grown as L6E9 cells in the presence of 0.5 mg/ml geneticin and differentiated in DMEM containing 5% (v/v) horse serum with antibiotics and geneticin.
Cells were washed 2 times with PBS, scraped, and homogenized in 2 ml of
ice-cold buffer containing 25 mM Hepes, 250 mM
sucrose, 4 mM EDTA, 1 trypsin inhibitor unit/ml of
aprotinin, 25 mM benzamidine, 0.2 mM
phenylmethylsulfonyl fluoride, 1 µM leupeptin, and 1 µM pepstatin, pH 7.4, using a Dounce A homogenizer.
Homogenates were processed as previously reported (27). Proteins were
measured by the method of Bradford (28) using -globulin as a
standard.
Before transport experiments, cells were incubated for 2 h in DMEM containing 0.2% bovine serum albumin. After this time, cells were washed, and transport solution (20 mM Hepes, 150 mM NaCl, 5 mM KCl, 5 mM MgSO4, 1.2 mM KH2PO4, 2.5 mM CaCl2, 2 mM pyruvate, pH 7.4) was added, together with 100 µM 2-deoxy-D-[3H]glucose (96 mCi/mmol). After 20 min, transport was stopped by addition of 2 volumes of ice-cold 50 mM glucose in PBS. Cells were washed 3 times in the same solution and disrupted with 0.1 M NaOH, 0.1% SDS. Radioactivity was determined by scintillation counting. Protein was determined by the Bradford method (28). Each condition was run in duplicate, and the nonspecific uptake (t = 0) was determined by incubation of the 2-deoxy-D-[3H]glucose in stop solution (50 mM glucose in PBS) instead of transport solution. In all cases, the value at t = 0 represented 4% of the basal transport activity at t = 20 min.
Electrophoresis and ImmunoblottingSDS-polyacrylamide gel electrophoresis was performed in accordance with the method of Laemmli (29). Proteins were transferred to Immobilon as reported (30). Transfer was confirmed by Coomassie Blue staining of the gel after the electroblot. Rabbit Bb antiserum raised against the purified human erythrocyte glucose transporter (a gift of Dr. C. Carter-Su, University of Michigan) was used at a 1:400 dilution and was incubated with transferred protein overnight at room temperature in 1% nonfat dry milk, 0.02% sodium azide in PBS to detect GLUT1. An anti-Sp1 affinity-purified rabbit polyclonal antibody (PEP-2, Santa Cruz Biotechnology) was used at a 5 µg/ml dilution in 1% nonfat dry milk, 0.02% sodium azide in PBS and incubated overnight at 4 °C. Detection of the immune complexes with the rabbit polyclonal antibodies was accomplished using 125I-protein A for 4 h at room temperature or using the ECL Western blot detection system (Amersham Corp.). Immunoblots were performed under conditions in which autoradiographic detection was in the linear response range.
RNA Isolation and Northern Blot AnalysisTotal RNA was extracted using the acid guanidinium thiocyanate/phenol/chloroform method as described by Chomczynski and Sacchi (31). All samples had a A260/A280 ratio above 1.8. After quantification, total RNA (30 µg) was denatured at 65 °C in the presence of formamide, formaldehyde, and ethidium bromide to allow the visualization of RNA. RNA was separated on a 1.2% agarose-formaldehyde gel and blotted on Hybond N filters. The RNA in gels and in filters was visualized with ethidium bromide by UV transillumination to ensure the integrity of RNA, to check the loading of equivalent amounts of total RNA, and to confirm proper transfer. Northern blot was performed as reported (27). The rat cDNA probe for GLUT1 was a 2,521 fragment obtained from Dr. M. Birnbaum (University of Pennsylvania) and was labeled with [32P]dCTP by random oligonucleotide priming.
GLUT1 CAT Reporter ConstructsPlasmid 2,106/+134-CAT was
constructed by inserting a 2,240-bp EcoRI-XhoI
fragment containing the rat GLUT1 promoter region from positions
2,106 to +134 (relative to the transcription start site) into the
XbaI site of pCAT-basic vector (Promega). pCAT-basic was
digested with XbaI, filling in of the ends with dNTPs in the presence of the Klenow fragment, and treated with alkaline phosphatase. The GLUT1 promoter DNA fragment was filled in the presence of dNTPs and
Klenow fragment and ligated to the pCAT-basic vector. 5
deletions were
generated by cleaving with HindIII at
1672 (
1672/+134-CAT), with HindIII and BstEII
(
1203/+134-CAT), with HindIII and BanII
(
812/+134-CAT), with HindIII and SmaI
(
201/+134-CAT), and with HindIII and AocI
(
99/
33-CAT), adding Klenow and T4 DNA ligase. The
33/+134-CAT
construct was generated by obtaining a 28-bp
HindIII-BssHII DNA containing the fragment
38/
15 from a 106-bp AvaII fragment (position
38/+68)
of the GLUT1 promoter subcloned in Bluescript. The 28-bp fragment was
subcloned into the BssHII site of the
BssHII-G1CAT construct (
15/+134). The 3
deletion
constructs were generated by cleaving with XbaI and BfrI (
2106/+46-CAT) and with XbaI and
BssHII (+2106/
15-CAT).
250,000 L6E9 cells were grown in
10-mm diameter plates for 2 days in DMEM with 10% fetal bovine serum.
Monolayers were washed, and DNA transfection was performed by using the
CaPO4 coprecipitation procedure (32). One ml of calcium
phosphate DNA precipitate containing 10 µg of various deletion
promoter-chloramphenicol acetyltransferase constructs, 5 µg of
pCMV--galactosidase control vector, and 20 µg of Bluescript DNA
(pSK
, Stratagene), was added dropwise to the plate, and
medium was added 15 min later. After 16 h, the cells were washed
and incubated with 1 ml of 15% glycerol in Hepes-buffered saline for 3 min, washed with DMEM, and incubated with fresh complete medium for 72 h. For the myotubes, the medium was changed to differentiation medium (DMEM supplemented with 2% fetal bovine serum) after the 15 min
incubation with the DNA precipitate.
The cells were washed 2 times with PBS and were
harvested by scraping in 1 ml of STE (10 mM Tris-HCl, pH
7.5, containing 100 mM NaCl, and 1 mM EDTA).
The cells were collected by centrifugation in a microcentrifuge for 10 min, and the pellet was resuspended in 200 µl of 0.25 M
Tris, pH 7.5. The cells were lysed by 3 cycles of freezing and thawing
at 37 °C. After centrifugation in a microcentrifuge for 5 min at
4 °C, the supernatant was stored at 20 °C.
The CAT activity of 75 µl of cytoplasmic extract was measured by
incubating 0.1 µCi of 14C-chloramphenicol, 1.3 mM acetyl-CoA, 200 mM Tris-HCl, pH 7.5, and the
soluble extract for 3.5 h at 37 °C. At the end of the incubation, extraction into ethyl acetate and thin layer chromatography (33) were performed. The CAT activity was quantitated using an
InstantImager (Packard). -Galactosidase activity was measured as
described (34).
Preparation of
the nuclear protein extracts was performed as described by Ausubel
et al. (35). The DNA probe (fragment 102/
37) was
obtained by digesting the
201/+134-CAT construct with
AvaII, purifying the 66-bp fragment, and
32P-end-labeled using the Klenow fragment of the DNA
polymerase. The gel mobility shift assays were performed in a 12-µl
reaction volume, containing 2 µg of double-stranded poly(dI-dC),
15,000 cpm of labeled DNA probe, 5-10 µg of protein of nuclear
extracts, 10 mM Hepes, pH 7.9, 10% glycerol, 50 mM KCl, 0.1 mM EDTA, 0.1 mM
phenylmethylsulfonyl fluoride, and 0.25 mM dithiothreitol. The mixture (without the labeled DNA) was incubated for 15 min at room
temperature. After the addition of the labeled DNA, the reaction
mixture was incubated for another 10 min at 4 °C, immediately loaded
on a 7% polyacrylamide gel (30:0, 8 acrylamide-bis-acrylamide), and
run in 0.5 × TBE buffer (45 mM Tris, 45 mM boric acid, 1 mM EDTA, pH 8) for 3-4 h at 8 V/cm. The gels were dried and autoradiographed.
For competition assays, varying concentrations of unlabeled probe
(fragment 102/
37) or the following oligonucleotides were used in
the reaction mixture prior to addition of extract: oligonucleotide I
(
100/
82), 5
-CCTCAGGCCCCGCCCCCCG-3
; oligonucleotide I mutated, 5
-CCTCAGGCCCCGTACCCCG-3
; oligonucleotide
55/
42,
5
-GCGCGGGCCAATGG-3
; and oligonucleotides containing the
consensus site for Sp1 or AP2 (Promega).
Supershift experiments were performed by incubating nuclear extracts or commercial human recombinant Sp1 protein (Promega), poly(dI-dC), and end-labeled probe as detailed above and then incubated for 30 min at 4 °C in the presence of 1 µg of Sp1 antibody (PEP-2, Santa Cruz Biotechnology) or with an irrelevant antibody. The samples were loaded on a 4% polyacrylamide gel, dried, and autoradiographed.
Differentiation of L6E9 myoblasts into myotubes was
associated with a diminished rate of basal glucose transport (near 80% decrease) (Fig. 1). Under these conditions, the total
cellular content of GLUT1 glucose transporter protein was also markedly reduced (levels in myotubes accounted for 22 ± 6% of values
found in myoblasts) (Fig. 1). This is in keeping with previous
observations performed in L6 muscle cells (21, 22). A reduction in
GLUT1 mRNA levels was also detected in L6E9 myotubes compared with
myoblast cells (levels in myotubes accounted for 33 ± 3% of
values found in myoblasts) (Fig. 1). The reduction in GLUT1 protein and
mRNA occurred under conditions in which no changes in the cellular content of 1-integrin protein or rRNA were detected (data not shown).
To determine the basis for the repression of GLUT1 expression,
myoblasts or myotubes were incubated in the presence of actinomycin D
(5 µg/ml) for different time periods, and the levels of GLUT1 mRNA were assessed (Fig. 2A). Results
indicate that the half-life of GLUT1 mRNA species was near 5 h, and no differences between myoblast and myotube cells were detected
(Fig. 2A). Therefore, the reduction in GLUT1 mRNA levels
during muscle cell differentiation is not due to alterations in the
stability of GLUT1 mRNA.
Next, myoblast or myotube cells were transiently transfected with a
fragment of the GLUT1 gene promoter (2106/+134) fused to the reporter
gene CAT. Cells transfected with the reporter gene showed very low
levels of CAT activity, similar to those shown by non-transfected cells
(Fig. 2B). Transfection with the construct
2106/+134-CAT
in L6E9 myoblasts caused a 11.5-fold increase in CAT activity (Fig.
2B). Furthermore, CAT activity detected in myotubes was
substantially reduced (60% decrease) compared with values in myoblasts
(Fig. 2B). These results indicate that the fragment
2106/+134 of the GLUT1 gene contains information that is relevant to
transcriptional activity in the myoblast and that allows repression in
response to myogenesis.
To determine the
cis-elements involved in the transcriptional activity of the GLUT1
promoter, 5 deletion constructs of the GLUT1 promoter fused to the CAT
reporter gene were generated and transiently transfected in myoblast
and myotube L6E9 cells (Fig. 3).
Deletion from 2106 to
812 of the GLUT1 gene caused no significant
alterations in CAT activity (Fig. 3), and deletion from
812 to
201
caused nearly 60% stimulation of CAT activity, suggesting a repressor
element (Fig. 3). The transcriptional activity of the
201 construct
was maximal and only a slight decrease was noted after deletion from
201 to
99. However, deletion of a further 66 base pairs (from
99
to
33) led to a marked reduction (80%) in the transcriptional
activity.
The repression of transcriptional activity due to myogenesis was
maximal in the 99/+134 construct although some differences were still
found in the
33/+134 construct (Fig. 3). The
33/+134 construct
contains the TATA box, located at
32/
27 relative to the
transcription start site (36).
To rule out the participation of the 3-end of the fragment of the
GLUT1 promoter which lies 3
of the transcription start site,
additional constructs were generated by 3
deletion (Fig. 4). Deletion of 88 base pairs lying between +134 and +46
caused a 60% reduction in transcriptional activity (Fig. 4),
suggesting elements important for the transcriptional activity. Under
these conditions, myogenesis reduced the transcriptional activity of all constructs studied (Fig. 4).
These data indicate that the fragment 99/
33 is responsible for the
transcriptional activity of the GLUT1 promoter. Furthermore, this
fragment, together with the fragment containing the TATA box, seems to
confer sensitivity to muscle cell differentation.
The
99/
33 fragment of the GLUT1 gene contains one consensus Sp1 site,
two AP-2-like sites, and one CAAT box (Fig. 6). To determine whether
nuclear proteins bind to the fragment
99/
33 of the GLUT1 promoter,
a DNA fragment encompassing the sequence
102/
37 was radioactively
labeled, and EMSA assays were performed in the presence of nuclear
extracts obtained from L6E9 myoblasts or myotubes (Fig.
5). A number of specific bands was detected in nuclear
extracts (Fig. 5). Some of them, named A1, A2 and B, were restricted to
myoblasts, and others (complexes D1 and D2) were more abundant in
myoblasts than in myotubes. In contrast, complexes C2 and D3 were more
abundant in myotubes than in myoblasts (Fig. 5). C1 was the only
complex to show a similar abundance in extracts from myoblasts and from
myotubes (Fig. 5).
To map the DNA elements that allowed the binding of the different
complexes, EMSA assays were performed in the presence of an excess of
unlabeled oligonucleotides (Fig. 6). EMSA assays performed in the presence of unlabeled oligonucleotide 100/
82 (oligonucleotide I), which contains the canonical Sp1 site, and an
overlapping AP-2-like site displaced, in a concentration-dependent manner, complexes A1 and A2 found in myoblast extracts (Fig.
7A). No bands were displaced in the presence
of the canonical sequence corresponding to the AP-2 site or with
oligonucleotide
55/
42, which contains the CAAT box (data not
shown).
Sp1 Binds and Transactivates the GLUT1 Promoter and Is Repressed during Myogenesis
Next, we focused on the nature of the complexes
A1 and A2, characteristic of myoblast extracts. Based on the presence
of a canonical Sp1 site in position 92/
87, we searched whether
binding of factors to this site might be responsible for complexes A1 and A2. EMSA assays were performed in the presence of an excess of
unlabeled oligonucleotides containing a canonical Sp1 site (Fig.
7A). Under these conditions, complexes A1 and A2 were
displaced very efficiently in the presence of 100-fold molar excess of
the oligonucleotide Sp1 (Fig. 7A). Furthermore, the
incubation in the presence of an excess of oligonucleotide
100/
82
in which the Sp1 site was mutated (Fig. 7A, oligonucleotide
Imut) failed to displace complexes A1 and A2 (Fig.
7A). Sp1 protein belongs to a family of zinc-finger
transcription factors (37, 38), and the formation of complexes in
band-shift assay is sensitive to the presence of Zn2+ or
EDTA in the medium (39). Based on this, gel-retardation analyses were
performed in the absence or presence of Zn2+ or EDTA. The
addition of Zn2+ to the medium increased the
formation of complexes A1 and A2 in a
concentration-dependent manner (Fig. 7B). In
contrast, addition of EDTA caused a concentration-dependent
inhibition of complexes A1 and A2 (Fig. 7B).
To confirm the binding of Sp1 protein, super shift assays were also
performed. To this end, EMSA assays were carried out in the presence of
an anti-Sp1 antibody. Due to the utilization of a different percentage
of polyacrylamide and due to a longer electrophoresis run, the complex
A1 previously seen as a single broad band was resolved as two distinct
complexes (named A1 and A1) (Fig. 8). In these studies,
recombinant Sp1 protein was also incubated with labeled fragment
102/
37 (Fig. 8). Results indicate that recombinant Sp1 forms a
complex with fragment
102/
37, which shows a retardation similar to
complex A1, and the formation of this complex was prevented in the
presence of an excess of oligonucleotide I (
100/
82) (Fig. 8). In
addition, anti-Sp1 antibody eliminated part of complex A1 and generated
a complex showing a greater retardation. A similar super-retarded band
was observed when recombinant Sp1 was incubated with anti-Sp1 antibody
(Fig. 8).
These results indicate that endogenous Sp1 present in extracts from
L6E9 myoblasts binds to the GLUT1 promoter. To determine whether Sp1
modulates the transcriptional activity of the GLUT1 promoter, L6E9
myoblasts or myotubes were co-transfected with the construct
2106/+134-CAT and the cDNA coding for Sp1. Sp1 caused a large
transactivation of the GLUT1 promoter activity (4.9-fold increase) in
myoblasts (Fig. 9). In addition, Sp1 activated the GLUT1
promoter activity in myotubes; however, the transcriptional activity
detected in myoblasts was much greater than in myotubes (Fig. 9).
We have found that Sp1 protein binds to the GLUT1 gene promoter in
myoblasts but not in myotubes. To determine the nature of the
mechanisms involved, we determined the level of Sp1 protein in nuclear
extracts obtained from L6E9 myoblasts and myotubes (Fig.
10). Sp1 protein was observed in Western blot as two
bands with an apparent molecular masses of 105 and 95 kDa, which is in
keeping with previous observations (40, 41). The content of Sp1 protein
in myoblasts was much greater than in myotubes (Fig. 10) (levels of Sp1
protein in myotubes accounted for 27 ± 10% of values found in
myoblasts). This effect was specific since the abundance of the
transcription factor STAT-1 was similar in preparations from myoblasts
or myotubes (data not shown).
Over-expression of MyoD Represses Sp1 and Inhibits the Transcriptional Activity of the GLUT1 Gene Promoter
The best
characterized factors that regulate the terminal differentiation of the
muscle cells are the members of the MyoD family. To determine whether
MyoD plays a role in the regulation of GLUT1 gene expression during
myogenesis, we studied the effect of the stable over-expression of MyoD
in C3H10T1/2 cells. The stable expression of MyoD in these cells caused
a marked reduction in the transcriptional activity of the GLUT1
promoter as assessed by transient transfection of the 2106/+134-CAT
construct (CAT activity levels in C3H10T1/2 wild type and
C3H10T1/2-MyoD were 43 ± 3 and 15 ± 3, respectively,
expressed as arbitrary units and corrected per
-galactosidase
activity).
EMSA assays revealed the presence of highly retarded complexes in
nuclear extracts from C3H10T1/2 cells that showed similar mobility to
complexes A1 and A2 from L6E9 myoblasts. Furthermore, these complexes
were competed with an oligonucleotide containing the consensus sequence
for Sp1 binding (Fig. 11) and with oligonucleotide I
(100/
82) (data not shown). Stable over-expression of MyoD caused
the disappearance of the complex binding to the Sp1 element (Fig. 11)
and the formation of low-retarded complexes (Fig. 11). Furthermore,
Western blot assays of Sp1 protein from nuclear extracts obtained from
wild-type C3H10T1/2 and C3H10T1/2-MyoD cells indicated a dramatic
down-regulation of Sp1 protein after MyoD over-expression (Fig. 11).
These results suggest a role of MyoD in the down-regulation of Sp1
associated with myogenesis.
In this study, we have demonstrated that GLUT1 is repressed in
muscle cells during differentiation as a consequence of alterations in
transcriptional activity of the GLUT1 gene, which seems to involve the
fragment 99/
33 and the fragment 5
proximal to the transcription
start site containing the TATA box. Furthermore, the levels of GLUT1
mRNA and the transcriptional activity of the GLUT1 promoter were
similarly reduced in response to myogenesis; this suggests that the
alterations in the transcriptional activity of the GLUT1 promoter are
sufficient to account for the GLUT1 repression. In the region
99/
33
of the GLUT1 gene, we have identified the binding of Sp1 to the GLUT1
gene promoter; this seems to be important from a functional viewpoint
since Sp1 transactivates, in transient transfection assays, the
transcriptional activity of GLUT1 promoter. In contrast to the current
view stating that Sp1 is a ubiquitous factor, we have found that
myogenesis leads to a drastic reduction in the formation of a
DNA-protein complex involving Sp1, which is due to a marked
down-regulation of Sp1 expression. Our results also indicate that MyoD
over-expression down-regulates Sp1 expression in cells, which is in
parallel to a reduction in the transcriptional activity of the GLUT1
gene. Based on this, we propose the model depicted in Fig.
12. According to this, the transcriptional activity of
the GLUT1 gene is high in proliferating myoblasts, in part due to a
high expression of the activator Sp1. Muscle cell differentiation is
associated with activation of MyoD transcription factors, which act as
master regulators leading to activation of many muscle-specific genes. In our model, Myo D activation leads to the repression of Sp1 expression in muscle cells. In turn, Sp1 down-regulation causes inactivation of the transcriptional activity of the GLUT1 gene and,
therefore, leads to GLUT1 down-regulation and to a diminished rate of
glucose transport. A prior report indicates that an Sp1-site is
required for the expression of Id (an inhibitory factor of MyoD
function) in muscle cells (18). Based on this, we additionally postulate that Sp1 down-regulation contributes to the repression of Id
found during myogenesis and which is known to participate in the
activation of myogenic transcription factors (17, 42).
Prior studies on the regulation of the GLUT1 gene have exclusively
focused on the functional role of two enhancer elements found in the
mouse GLUT1 gene. The first enhancer has been located 2.7 kilobases
upstream of the transcription start site, whereas the second is in the
second intron of the gene (43). These enhancers permit the activation
of the transcription in response to growth factors, insulin, or hypoxia
(43-45). Here, we have analyzed the properties of the proximal
promoter of the rat GLUT1 gene. Our results indicate the presence of a
repressor element located in the fragment 812/
201 that is active
both in myoblasts and in myotubes. Our data also suggest that the
fragment close to the TATA box or the TATA box itself participates in
the repression of GLUT1 transcription during myogenesis. In this
regard, it has been reported that factors such as TEF-1 or NC2 block
the formation of TBP-TATA complexes and inhibit gene transcription (46,
47). Whether some of these factors play a role through the TATA box in
the regulation of the GLUT1 gene during myogenesis remains unknown.
Furthermore, we have identified the fragment 99/
33 of the GLUT1
gene that is responsible for the transcriptional activity of the GLUT1
promoter. We have also identified the presence of different complexes
found in myoblasts or myotube nuclear extracts and that bind to the
99/
33 region. Specifically, we have identified high-retardation
complexes (named A1 and A2) that are restricted to myoblasts. Based on
the selective competition of these bands to oligonucleotides containing
the Sp1 binding site, the sensitivity of these complexes to
Zn2+ and EDTA, the fact that they show a similar
retardation to recombinant Sp1, and that there is a super-shift in the
presence of an antibody against Sp1, we propose that Sp1 participates
in the formation of these complexes.
In our study, we have found that Sp1 stimulates the transcriptional
activity of the GLUT1 gene 5-fold in transient transfection assays.
These data, together with the fact that there is a high expression
level of Sp1 protein in nuclear extracts obtained from myoblasts and
that Sp1 binds to the fragment 99/
33 selectively in myoblasts,
strongly support the hypothesis that Sp1 regulates GLUT1 transcription
in muscle cells. Interestingly, the binding of Sp1 protein to the GLUT1
promoter correlates with a high GLUT1 gene expression in a variety of
experimental conditions.2 The transient
transfection of Sp1 into myotubes led to a stimulation of the GLUT1
rate of transcription that was markedly lower than that obtained in
myoblasts. These results suggest a defect of Sp1 action in myotube
cells. In this regard, it has been shown that Sp1 forms heteromeric
complexes with several cellular proteins. The TATA-binding protein
protein-associated protein TAF110 binds Sp1 and functions as a
co-activator in Sp1-dependent transcription (37). Sp1 also
interacts with the cellular protein YY1 (48, 49), with the RelA subunit
of NF-
B (50), and with the bovine papilloma virus (51). Based on the
fact that the cotransfection of a retinoblastoma expression vector is
able to modulate the transactivation of responsive genes by Sp1, the
function of Sp1 has been linked to that of the retinoblastoma protein
(52, 53) through retinoblastoma control elements. Some mechanisms of
inhibition of Sp1 action have also been reported. Thus, the
cell-cycle-regulatory protein 107 can be found endogenously associated
with Sp1 and, in cotransfection assays, p107 specifically represses
Sp1-dependent transcription (41). Furthermore, G10BP
protein or Sp3 competes with Sp1 for G-rich sequences and inhibits Sp1
action (54-56). Based on all these items of information, it might be
postulated that myogenesis not only leads to Sp1 down-regulation but
also causes alteration in the biological potency of Sp1, which might be
explained either by alterations in the proteins that allow the
formation of active complexes or due to the presence of Sp1-binding inhibitory proteins.
Sp1 is thought to be a ubiquitously expressed transcription factor that plays a primary role in the regulation of a large number of genes, including constitutive housekeeping genes and inducible genes (57). Recent studies indicate that Sp1 is a limiting factor in cultured cells and that overexpression of Sp1 increases expression from promoters containing GC-box elements (58). Additionally, there is evidence that Sp1 expression is regulated. Thus, Sp1 levels increase during SV40 infection of the CV1 cells (58). Furthermore, it has been reported that Sp1 expression varies greatly in different cell types and changes in Sp1 occur during development in vivo (59). Thus, high levels of Sp1 expression were found in spermatids, T cells, epithelial cells and hematopoietic cells (59). Interestingly, the expression of Sp1 mRNA in heart and skeletal muscles was higher in early neonatal mice than in adult animals (59). Along these lines, we have found in this study that myogenesis greatly alters the expression of Sp1 in muscle cells. Furthermore, our results support the view that MyoD causes down-regulation of Sp1 in stable transfection assays. In summary, we propose that activation of MyoD transcription factors causes down-regulation of Sp1 protein, and this might be one the mechanisms that lead to GLUT1 repression and accomplishment of the differentiation program in the muscle cell. The mechanisms underlying Sp1 down-regulation in response to MyoD transcription factors are currently under study in our laboratory.
We thank R. Rycroft for editorial support. We also thank Dr. C. J. Ciudad and V. Noé (Universitat de Barcelona) for thorough discussions of super-shift assays and Dr. J. Stephens (Boston University Medical School) for many suggestions.