University of Queensland Centre for Molecular and Cellular Biology Institute for Molecular Bioscience St. Lucia, 4072 Queensland, Australia
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ABSTRACT |
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INTRODUCTION |
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Great strides have been made in improving our understanding of muscle differentiation (myogenesis), hypertrophy, regeneration, and decay through the exploitation of the mouse muscle cell line, C2C12 [a subclone of C2 cells (7) deposited into ATCC (Manassas, VA)]. In this in vitro culture system, proliferating C2 myoblasts fuse into postmitotic multinucleated myotubes that acquire a muscle-specific phenotype. This cellular morphogenesis is accompanied by muscle-specific gene expression. RMSs are classified into two broad types, i.e. embryonal and alveolar, that have characteristic clinical, pathological, and histological features (8). Embryonal RMSs occur in young children and account for 60% of cases. The tumor is characterized by malignant spindle and round cells that contain skeletal muscle cross-striations and are located in specific sites, including the head/neck region, genitourinary tract, and orbit. In contrast, alveolar RMSs occur during adolescence as primary tumors of the extremities or trunk. This histological variant is characterized by the presence of fibrovascular septa that form alveolar like spaces filled with monomorphous malignant cells (5, 6, 8).
A group of basic helix-loop-helix (bHLH) proteins encoded by the myoD gene family (myoD, myf-5, myogenin, and MRF-4) and a second class of transcription factors, the myocyte enhancer factor-2 family (MEF2A-D), are required for proper muscle differentiation. The bHLH proteins and MEF2 transcription factors function in a cooperative manner to control the mutually exclusive events of division and differentiation (9, 10, 11).
Transcription is also regulated by the structural conformation of chromatin, a complex made up of DNA, histones, and other proteins. Chromatin structure is regulated by cofactors such as histone acetyltransferases (HATs) and deacetylases (HDACs). Histone acetylation and deacetylation affect accessibility of DNA to the transcriptional machinery, leading to transcriptional activation or repression, respectively. Both histone hyperacetylation and hypoacetylation have been associated with the neoplastic process, underscoring the need for improving our understanding of this process (12).
The cofactors PCAF and p300 that have HAT activity have been demonstrated to function as critical coactivators for the muscle-specific bHLH protein, MyoD, during myogenic commitment. However, skeletal muscle differentiation and the activation of muscle-specific gene expression are dependent on the concerted action of another bHLH factor, myogenin, and the MADS protein, MEF2, that function in a cooperative manner.
The cofactors GRIP-1 (glucocorticoid receptor interacting protein 1) and CBP/p300 have HAT activity and function as coactivators for MEF2C during myogenesis. (Refs. 13, 14, 15 and references therein). Class II HDACs (HDAC4 and 5) interact with the MEF2 proteins, and inhibit MEF2-dependent transactivation (16) and myoblast differentiation (17). The transcriptional activity of class II HDACs is controlled by compartmentalization (16) and 143-3-mediated subcellular localization (18). HDACs-4 and -5 are expressed constitutively in myoblasts and myotubes during C2C12 myogenesis, presenting a dilemma in terms of transcriptional control and the differentiation process (17). However, recently it has been demonstrated that myogenesis is controlled via differentiation-dependent nucleocytoplasmic trafficking of HDACs.
GRIP-1 belongs to the structurally related but genetically distinct SRC family (reviewed in Ref. 19) has three members variously denoted as SRC-1/N-CoA-1 (20, 21, 22), SRC-2/GRIP-1/TIF-2/N-CoA-2 (23, 24), and SRC-3/ACTR/pCIP/RAC-3/AIB1/TRAM1(22, 25, 26, 27, 28).
Many recent studies have implicated and confirmed the link between alterations in chromatin structure and differentiation, disease, and cancer. Chromatin remodeling by histone acetylation is generally associated with cell cycle arrest and differentiation, whereas deacetylation promotes cellular proliferation and growth. Many lines of evidence suggest a strong link between chromatin structure, normal and aberrant growth, and tumorigenesis. Recent studies implicate alterations in chromatin structure by histone hyperacetylation/deacetylation as playing an important role in either the genesis or suppression of cancer. Whether histone hyperacetylation or deacetylation is involved appears to depend on the specific target gene (reviewed in Refs. 12, 15, 29, 30) and references therein). For example, studies suggest that factors with HAT activity, e.g the SRC/p160 factors, are involved in transcriptional activation and exert antiproliferative effects. For example, translocations of SRC-2/TIF-2/GRIP-1 [i.e. coactivators with HAT activity] to genes implicated in chromatin structure and function have been documented in several leukemias: monocytic leukemia zinc finger protein gene (MOZ)-mediated monocytic leukemia; mixed lineage leukemia gene (MLL)-mediated acute lymphoblastic leukemias (31, 32, 33). Amplification and/or overexpression of AIB-1 is implicated in the onset, development, and prognosis of breast cancer (26).
Recently, we have observed that muscle differentiation is regulated by MEF2C and GRIP-1 (13). The mechanism involves direct interactions between MEF2 and GRIP-1. However, the subcellular localization and nucleocytoplasmic trafficking of the HAT complexes in the process of mammalian muscle differentiation and/or in skeletal muscle tumors have not been investigated. RMS cells have impaired myogenic factor activity, and it has been suggested that they lack some critical cofactors (34). In this study we have observed that RMS cells derived from skeletal muscle tumors harbor defects in the subcellular localization, expression, and activity of key transcription factors and SRC cofactors that control essential elements of the muscle-specific differentiation program.
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RESULTS |
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We used immunofluorescence to investigate the subcellular localization
and expression of the SRCs and MEF2 proteins during skeletal myogenesis
in culture. To detect and analyze the subcellular localization
pattern of SRC-2 we conducted immunofluorescence staining using
monoclonal anti-SRC-2 antibody. SRC-2 was endogenously expressed in all
low and high confluency proliferating C2C12 myoblasts grown in high
serum (DMEM supplemented with 20% FCS) (Fig. 1, A and B). The majority of staining was
restricted to the cell nucleus, with weak (but significant) staining
observed in the cytoplasm (Fig. 1
, A and B). The staining pattern in
the nucleoplasm was extranucleolar and appeared in prominent nuclear
dots in a punctate pattern characteristic of nuclear and splicing
bodies.
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We then conducted further immunofluorescence staining of cells after
7296 h of serum withdrawal. At this stage the cells form postmitotic
multinucleated cells (myotubes), where the nuclei line up end to end.
This demonstrated that SRC-2 was endogenously expressed in
differentiated postmitotic muscle cells and also localized to the
nucleus in prominent dot-like structures. (Fig. 1D). We also conducted
immunofluorescent analysis of SRC-1 and -3 using antibodies from
Santa Cruz Biotechnology, Inc. (Santa Cruz, CA) and
Affinity BioReagents, Inc. (Golden, CO) but the
level of staining was low to insignificant and consistent with the
Northern analysis (data not shown).
MEF2 is Expressed in the Nucleus during Skeletal Muscle
Differentiation: MEF2 Is Expressed in a Variegated/Mosaic Pattern in
Proliferating Myoblasts
We among others have previously demonstrated that when
proliferating C2C12 myoblasts are induced to biochemically and
morphologically differentiate into postmitotic multinucleated myotubes,
that the transition is associated with an increase in the levels of
MEF2 proteins during myogenesis.
To detect and analyze the subcellular localization pattern of MEF2, we
conducted immunofluorescence staining using polyclonal anti-MEF2
antibody. MEF2 was endogenously expressed in the nucleus of
proliferating C2C12 myoblasts grown in high serum (Fig. 1E). We
observed consistently that in a population of proliferating myoblasts
(which express low levels of MEF2), that only 15% of the cells
(nuclei) express MEF2. This can be seen by comparing Fig. 1E
to the
4,6-diamidino-2-phenylindole (DAPI)-stained nuclei in Fig. 1F
.
Importantly, although the staining of the protein is restricted to the
nucleus, the expression pattern in proliferating myoblasts is
mosaic/variegated (35, 36), i.e. it is either active
or completely inactive in individual cells, which may suggests this
transcription factor maybe regulated in a binary fashion (35, 36). In
contrast, MEF2 was expressed in more than 80% of the nuclei in
differentiated postmitotic myotube cells (Fig. 1
, G and H).
It should be noted that the MEF2 antibody used in these analyses was made against an epitope at the C terminus of MEF2A and has significant cross-reactivity to MEF2C, but not MEF2D or MEF2B. Previous MEF2 RNA and protein analysis using the cell culture model of skeletal myogenesis suggests MEF2D is expressed in the nuclei of proliferating myoblasts grown in high serum (whereas MEF2A and C are entirely absent) and myotubes (37). In contrast, MEF2A expression is induced only after serum withdrawal (38). Furthermore, MEF2C is expressed only in well differentiated postmitotic myotubes after more than72 h of serum withdrawal (39, 40). Our analysis in the context of these previous investigations suggests that early MEF2 expression is probably MEF2A and, during the progressive increase in MEF2 positive nuclei (coupled to the differentiation program) that MEF2A and subsequently MEF2C are expressed. This pattern of expression, i.e. the progressive increase in the proportion of nuclei expressing MEF2 from mosaic/variegated to total expression in all nuclei has been described as a characteristic of binary regulation (35, 36). The future availability of MEF2A and C-specific antibodies will help to determine whether this suggestion is correct.
GRIP-1 and MEF2 Are Colocalized in the Nuclei of Differentiated
Muscle Cells
We had previously demonstrated that the steroid receptor
coactivator, GRIP-1, coactivates MEF2C-mediated transcription, and
that the mechanism involves direct interaction between MEF2C and SRC-2
(13). This was demonstrated by a number of biochemical interaction
assays and two-hybrid analysis. In this study we examined the
expression of SRC-2 and MEF2 in muscle cells by double staining
myotubes with antibodies directed against SRC-2 and MEF2. Conventional
and confocal two-color immunofluorescence microscopy confirmed that the
SRC-2 (Fig. 2, A and D) and MEF 2 (Fig. 2
, B and E) proteins were endogenously expressed in differentiated
muscle cells. The colocalization of SRC-2 and MEF2 in differentiated
muscle cell nuclei was confirmed by merging these two images, which
showed considerable colocalization (Fig. 2
, C and F, yellow)
of the two proteins in the nucleus.
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PolyA+ RNA from C2C12 cells was isolated from proliferating myoblasts, confluent myoblasts, and postmitotic myotubes after 4, 8, 24, and 72 h of serum withdrawal and examined by Northern blot analysis. Similarly, RNA was isolated from alveolar RMS Rb-2061 cells grown in high serum after 72 h of serum withdrawal.
We have previously demonstrated in C2 cells (13) that SRC-2/GRIP-1 mRNA
was constitutively expressed in proliferating myoblasts as the cells
exited the cell cycle and fused to form postmitotic differentiated
multinucleated myotubes that had acquired a muscle-specific
phenotype (Fig. 3A). The
hybridization signal corresponded to the expected transcript size of
approximately 7.5 kb. It was necessary to use polyadenylated RNA since
the level of expression was quite low relative to
glyceraldehyde-3-phosphate dehydrogenase (GAPDH) or myogenin, and
exposure times required for the detection of SRC-2 were approximately
10-fold longer than that of GAPDH (Fig. 3A
). The SRC-1 mRNA was very
weakly expressed, however, and we could not detect the mRNA transcript
that encoded SRC-3. In alveolar RMS Rb-2061 cells, SRC-2 mRNA was not
detectable by Northern analysis; however, the mRNAs encoding SRC-1 and
SRC-3 were abundantly expressed relative to GAPDH.
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Differential Expression and Subcellular Localization of SRC-2 in
Alveolar (Rb-2061) and Embryonal (A-204) RMS Cells
We used immunofluorescence to investigate the subcellular
localization and expression of the steroid receptor coactivator, SRC-2,
in alveolar RMS Rb-2061 cells. To detect and analyze the subcellular
localization pattern of SRC-2, we conducted immunofluorescence staining
using monoclonal anti-SRC-2 antibody. SRC-2 was very weakly expressed
after 3 days of serum withdrawal in alveolar RMS Rb-2061 cells (Fig. 4A). In contrast to C2C12 cells, the
staining pattern did not appear in prominent nuclear dots
characteristic of nuclear and splicing bodies. The staining was rather
diffuse, and observed in the nucleus and cytoplasm. Merging the SRC-2
image (green) with DAPI image (blue) clearly
highlighted the expression of SRC-2 in the cytoplasm (Fig. 4B
).
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We then used immunofluorescence to investigate the subcellular
localization and expression of the steroid receptor coactivator, SRC-2,
in embryonal RMS A-204 cells. To detect and analyze the subcellular
localization pattern of SRC-2, we conducted immunofluorescence staining
using monoclonal anti-SRC-2 antibody. SRC-2 was strongly expressed in
proliferating embryonal RMS A-204 cells (Fig. 4E). In contrast to
alveolar RMS Rb-2061, the staining pattern in embryonal A-204 cells
appeared in prominent nuclear dots characteristic of nuclear and
splicing bodies. SRC-2 was predominantly expressed in the
nucleus. Comparing the SRC-2 image (green) (Fig. 4E
) with
the DAPI image (blue) (Fig. 4F
) clearly highlighted the
specific nature of SRC-2 staining, in the nucleus of embryonal RMS
A-204 cells.
SRC-1 and SRC-3 Are Abundantly Expressed in Embryonal and Alveolar
RMS Cells: Subcellular Localization of p160 Factors Is Defective
We used immunofluorescence to investigate the subcellular
localization and expression of the steroid receptor coactivators, SRC-1
and 3, in the alveolar (Rb-2061) and embryonal (A-204) RMS cells
after 72 h of serum withdrawal. To detect and analyze the
subcellular localization pattern of SRC-1 and -3, we conducted
immunofluorescence staining. SRC-1 in alveolar RMS Rb-2061 (Fig. 5, A and B) and embryonal RMS A-204 cells
(Fig. 5
, C and D) were expressed in RMS cells. The staining was rather
diffuse and was observed in cytoplasm and nucleus. The staining pattern
did not appear in prominent nuclear dots characteristic of nuclear and
splicing bodies. Merging the SRC-1 image (green) with the
DAPI image (blue) clearly highlighted the significant SRC-1
staining in the cytoplasmic expression (Fig. 5
, B and D). Similarly,
significant expression of SRC-3 is highlighted in alveolar RMS Rb-2061
(Fig. 5
, E and F) and embryonal RMS A-204 cells. (Fig. 5
, G and H).
These data suggest SRC -1 and -3 are aberrantly localized in alveolar
and embryonal RMS cells, in contrast to the nuclear-specific
pattern of SRC-2 staining in embryonal RMS A-204 cells (Fig. 4E
).
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We then analyzed the subcellular localization pattern of MEF2 in
embryonal RMS cells. Immunofluorescence staining using polyclonal
anti-MEF2 antibody demonstrated MEF2 was endogenously expressed in
alveolar RMS cells growth medium (RPMI supplemented with 10% FCS) and
after serum withdrawal (differentiation medium, RPMI supplemented with
2% HS) (Fig. 6, panels H and K, respectively). The staining pattern of
MEF2 in alveolar and embryonal RMS cells grown in high serum was
similar. In contrast to the accumulation of MEF2 in the cytoplasm after
serum withdrawal in Rb 2061 cells (Fig. 6E
, yellow
arrows), MEF2 accumulated in the nucleus after serum
withdrawal in the embryonal RMS A-204 cells (Fig. 6K
). This suggested
the trafficking of MEF2 into the nucleus was regulated by serum
withdrawal and perhaps occurred in a density-dependent manner.
Embryonal RMS A-204 cells express SRC-2 in the nucleus; furthermore,
serum withdrawal leads to a predominant localization of MEF2 in the
nucleus. MEF2 functions through the formation of a complex that also
includes the cofactor SRC-2. The formation of this complex mediated by
direct protein-protein interaction is critical to the activation and
progression of the differentiation program (13). Hence, we proceeded to
investigate the subcellular localization of the MEF2 cofactor SRC-2 in
the RMS A-204 cells by double immunostaining after serum withdrawal.
The expression of MEF2 and SRC-2 in embryonal A-204 cells was further
highlighted by double staining cells with antibodies directed against
SRC-2 and MEF2. Immunostaining clearly showed that SRC-2 (Fig. 6, G and
J, green) was localized to the nucleus of all cells cultured
in growth and differentiation medium. Two-color immunofluorescence and
merging (Fig. 6
, I and L, yellow) highlighted the nuclear
expression of SRC-2 and the predominant localization of MEF2 from the
cytoplasm to the nucleus after serum withdrawal.
We investigated this differential expression and localization of
MEF2 by Western analysis of total lysates and nuclear extracts from C2
cells and alveolar RMS Rb 2061 cells. We observed that MEF2 proteins
were expressed and induced in both C2C12 cells and RMS cells after
serum withdrawal (Fig. 6M). However, RMS cells abundantly expressed
MEF2A and the faster migrating MEF2C species (
50 kDa). Total
expression of MEF2 in both cell types after serum withdrawal, as
measured by immunostaining and Western analysis, was similar.
Curiously, Western analysis of nuclear extracts (Fig. 6N
) from C2 and
alveolar RMS Rb-2061 cells indicated that significant amounts of the
faster migrating MEF2C species from the RMS cells was not shuttled into
the nucleus after serum withdrawal. This observation was
consistent with the immunofluorescent analysis. In
summary, this indicates that MyoD is exclusively localized to the
nucleus; however, the partner protein, MEF2, is predominantly localized
to the cytoplasm. This confirmed that the trafficking of MEF2C into the
nucleus of alveolar RMS Rb-2061 cells is impaired.
Similarly, Western analysis of the embryonal RMS A-204 cells
demonstrates that these cells accumulate large amounts of MEF2C in the
nucleus after serum withdrawal (Fig. 6P), in contrast to the alveolar
cells. This correlated with the expression of SRC-2 in the nuclei of
these cells detected by immunostaining.
The Transcriptional Activity of the Myogenic Factors Is Impaired in
Alveolar RMS Rb-2061 Cells: MEF2 Function Is Significantly Compromised
in Alveolar RMS Rb-2061 Cells
We compared the relative activity of MEF2C, MyoD, and myogenin in
muscle and RMS cells using the GAL4 hybrid assay. We observed that
MEF2C, MyoD, and myogenin induced transcription by approximately 25-,
50-, and 120-fold (relative to GAL0, arbitrarily set to 1) in C2 muscle
cells. In comparison, in alveolar RMS Rb-2061 cells, MEF2C, MyoD, and
myogenin induced transcription by approximately 4-, 22-, and 45-fold.
This suggested that in RMS cells, the transcriptional activity of MEF2C
is significantly impaired (>5-fold). MyoD and myogenin activity is
reduced about 2-fold (Fig. 7A).
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To further substantiate these results we tested the activity of a
MEF2-dependent reporter with three tandem copies of the MEF2 binding
sites upstream of a basal E1b promoter in C2C12 and RMS Rb-2061 cells
cultured in differentiation medium (Fig. 7C). These experiments clearly
demonstrated again that the activity of a MEF2-dependent reporter is
reduced by 4-fold in alveolar RMS Rb-2061 cells. In conclusion, our
studies demonstrate that the activity and localization of MEF2C in
alveolar RMS cells are impaired.
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DISCUSSION |
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Muscle cells abundantly express SRC-2 (and weakly express SRC-1). Alveolar RMS-Rb 2061 cells derived from malignant skeletal muscle tumors very weakly express the steroid receptor coactivator, SRC-2, and more abundantly express SRC-1 and SRC-3. In contrast, the embryonal A-204 cells express SRC-2; however, they also express SRC-1 and SRC-3. Furthermore, SRC-1 and -3 are inappropriately localized in cells derived from alveolar and embryonal RMS. This observation correlates with a number of other studies on steroid receptor cofactor expression, cancer development, and progression. For example, normal brain tissue is positive for SRC-2(GRIP-1/TIF-2) and SRC-1; however, meningiomas (associated with breast cancer) abundantly express SRC-3 and SRC-1 (41). SRC-3 (AIB-1) was originally identified in a search for genes whose expression and copy number were elevated in human breast cancers (26). SRC-3 amplification and increased expression correlate with steroid receptor-positive breast and ovarian tumors (42). SRC-3 amplification in association with other coamplifications (e.g. ERBB2, MYC, CCND1, or FGFR1) suggest poor prognosis and worsened outcome (43, 44, 45). Pancreatic carcinoma and gastric tumors are also linked to SRC-3 amplification/overexpression and chromosomal aberration (46, 47). Our study suggests that elevated SRC expression may also be a marker for RMSs.
Aberrant subcellular localization of SRCs, and of critical transcription factors in RMSs, rather than over/mis-expression, suggests that inappropriate/dysfunctional trafficking of essential transcription factors and cofactors is associated with the RMS phenotype. However, we cannot rule out increased export or reduced degradation of cytoplasmic MEF2. Interestingly, inappropriate localization of SRC-2 and dysfunctional trafficking of MEF2 in alveolar cells relative to embryonal RMS cells may correlate with the poor prognosis for individuals with alveolar RMS.
Class II HDACs (HDAC-4 and -5) interact with the MEF2 proteins and inhibit MEF2-dependent transactivation and myoblast differentiation (17). The transcriptional activity of class II HDACs is controlled by compartmentalization (16, 17) and 143-3 protein- mediated subcellular localization (18). HDACs-4 and -5 are expressed constitutively in myoblasts and myotubes during C2C12 myogenesis, presenting a dilemma in terms of transcriptional control and the differentiation process (16). However, recently it has been demonstrated that myogenesis is controlled via differentiation-dependent nuclear export of HDACs. The shuttling of HDACs and the nucleocytoplasmic trafficking of these transcriptional repressors are regulated by phosphorylation. Calcium-calmodulin-dependent protein kinase stimulates myogenesis, blocks HDAC-mediated inhibition of MEF2, and induces export of the class II HDACs from the nucleus. Export of these repressors from the nucleus is mediated by the 143-3 proteins in a phosphorylation-dependent manner.
These results raise the scenario that kinase signaling regulates trafficking of the SRCs into the nucleus. The mitogen-activated protein kinases (MAPKs), p38 and ERK5, stimulate MEF2 activity by direct phosphorylation of the C-terminal activation domain (46). Analogously, SRC-3 has been demonstrated to be a phosphoprotein targeted by MAPK (47). Furthermore, MAPK activation of SRC-3 facilitates the recruitment of other cofactors and associated HAT activity (47). These observations in the context of defective SRC cofactor localization in RMS cells (this work) suggest that protein kinase signaling mediates the active shuttling of the SRCs into the nucleus. This hypothesis is strongly supported by several observations: 1) myogenesis is dependent on p38 MAPK activation; 2) RMS cells are deficient in p38 MAPK; 3) MAPK kinase 6 expression in RMS cells stimulates MEF2 activity; and 4) ectopic expression of an activated MAPK kinase 6, which induces p38 MAPK, restored MyoD and MEF2 function and led to terminal differentiation.
Our current studies are directed at examining whether MAPK activation stimulates the localization of MEF2 and the SRC cofactors into the nucleus. Our preliminary data suggest O-tetradecanoylphorbol-13-acetate treatment completely restores MEF2C activity in alveolar RMS cells (data not shown). Time will tell whether drug intervention targeted at protein kinases, and nucleocytoplasmic trafficking of SRC cofactors is a plausible therapeutic strategy.
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MATERIALS AND METHODS |
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For isolating total proteins, cells were lysed in lysis buffer from LUC-Lite luciferase assay kit (Packard Instruments, Meriden, CT) and cleared by centrifugation. Proteins were dialyzed against PBS to reduce the detergents concentration, and the protein concentration were determined by DECA protein assay kit (Pierce Chemical Co., Rockford, IL).
Immunoblotting
Aliquots of 30 µg nuclear protein or 50 µg total
protein were used for each sample. The proteins were blotted onto
nitrocellulose membrane at 160220 mA overnight in Towbin
buffer (25 mM Tris at pH 8.3, 200 mM glycine,
20% methanol, 0.4% SDS) after being resolved on 7% or 10%
SDS-PAGE gels. The blots were blocked in blocking solution (5% skim
milk, 0.05% Tween 20 in PBS) at 37 C for 1 h and then incubated
with 0.1 µg/ml MEF2 or TIF2 antibodies diluted in blocking solution
at room temperature for at least 2 h or at 4 C overnight. Blots
were washed several times in PBST (0.05% Tween in PBS) after the
primary antibody incubation and then incubated with secondary antibody
diluted in blocking solution at room temperature for 1 h. After
the secondary antibody incubation the blots were washed in PBST for at
least 20 min with several changes of PBST and then viewed with
chemiluminescence (ECL; Amersham Pharmacia Biotech,
Arlington Heights, IL).
Northern Analysis
Total RNA from different differentiation stages of C2C12 and RMS
cells was prepared by using an acid guanidinium thiocyanate-based
method (50). Briefly, cells were harvested in PBS containing 5
mM EDTA and then lysed in solution D (4 M
guanidinium thiocyanate, 25 mM sodium citrate, pH 7.0,
0.5% sarcosyl, 0.1 M 2-mercaptoethanol.). Cell lysates
were kept at -20 C before RNA was extracted. To extract total RNA,
water-saturated phenol was added to the lysate and then vortexed.
Chloroform was added and vortexed vigorously for 30 sec before
centrifugation at 4 C to separate the aqueous phase from the organic
phase. RNA in the aqueous phase was precipitated by isopropanol
precipitation and was resuspended in diethyl pyrocarbonate
(DEPC)-treated water.
Total RNA was further purified using the Oligotex mRNA Mini Kit (QIAGEN, Chatsworth, CA), and an aliquot of 5 µg poly (A)+ RNA was used for each sample. RNA was denatured at 70 C for 10 min in RNA sample buffer (50% formamide, 2 M formaldehyde, 1x MOPS, 50 µg/ml ethidium bromide), and quickly chilled on ice before being run on a 1% agarose-2 M formaldehyde denaturing gel in 1x 3-[N-morpholino]propanesulfonic acid (MOPS) buffer (20 mM MOPS, 5 mM sodium acetate, 1 mM EDTA). The fractionated RNA was blotted onto nylon membranes (Hybond N; Amersham Pharmacia Biotech) overnight with a vacuum blotter (LKB, Rockville, MD) to ensure that the high molecular weight RNA was transferred completely. The nylon membranes were UV cross-linked and baked at 80 C after blotting. The nylon membranes were prehybridized in 5x SSPE, 5x Denhardts solution, 0.1% SDS, 100 µg/ml herring sperm DNA, 5% glycine overnight at 42 C before the addition of specific probes. Specific probes were labeled by a random priming method using specific cDNAs as templates and were further purified with NICK column (Pharmacia Biotech, Piscataway, NJ). Prehybridized blots were hybridized with approximately 107 cpm of specific probe for another 2448 h in prehybridization buffer at 42 C. After hybridization the blots were washed in 1x or 0.5x SSC for 3060 min at 65 C and were visualized by autoradiography.
The SRC-1 probe encompasses amino acids 1,1381,441 and 180 bp
of the 3'- untranslated region, the GRIP-1 probe spans amino acids
5451,157, and the RAC-3 probe contains amino acids 507937. The
MyoD, myogenin, p21, and cyclin D1 probes were EcoRI
digestion fragments isolated from pEMSV-myoD, pEMSV-myogenin, pCMW35,
and pGEX-3X-CYL1 plasmids, respectively. GAPDH cDNA was amplified by
RT-PCR and gel isolated for use as a template for random priming. cDNA
probes were radioactively labeled by random priming. DNA fragments
(50100 ng) were boiled with 20 ng of random primers (pdN6;
Pharmacia Biotech) at 100 C for 10 min and quickly chilled
on ice for another 10 min. Denatured DNA was then incubated overnight
with 510 µl of [-p32]-dCTP, 62.5 µM of
dATP, dGTP, dTTP, and 5 U of Klenow enzyme (New England Biolabs, Inc., Beverly, MA) in 1x EcoPol buffer (New England Biolabs, Inc.). Probes were purified using NICK columns
(Pharmacia Biotech) according to the manufacturers
instructions.
Fluorescence Immunohistochemistry
For fluorescence immunohistochemistry analysis, C2C12 and RMS
cells were grown on cover slides held in six-well dishes. Cell were
washed in PBS once and then fixed in 100% methanol at -20 C for 5
min. After fixation cells were washed once in PBS and then quenched in
PBS containing 50 mM NH4Cl to avoid
the deleterious effect of the methanol on the antibodies. Cells were
then blocked in blocking solution (0.2% fish skin gelatin and 0.2%
BSA diluted in PBS) at room temperature for 10 min before being
incubated with primary antibody diluted 1:200 in blocking solution at
room temperature for 30 min. Then cells were washed with PBS four times
before being incubated with Bodify-, Texas red-, or fluorescein
isothiocyanate (FITC)-conjugated secondary antibodies (Molecular Probes, Inc., Eugene, OR) diluted 1:2,000 in blocking solution
at room temperature for 30 min and then washed four times with PBS
again. To visualize the nuclei, cells were incubated with DAPI (1:2,000
dilution in PBS) at room temperature for 10 min after the secondary
antibody incubation and were then washed thoroughly with
PBS.
Transient Transfection and GAL4 Hybrid Analysis
C2C12, and RMS cell line SJRH30/RMS13 were passaged into 12-well
plates and transfected at 6080% confluence with 1,000 ng of
reporter, G5E1b-LUC, and 330 ng of GAL-MEF2C, GAL-MyoD, or GAL-myogenin
in the presence and absence of GRIP-1 SJRH30) by liposome (Dotap and
Dosper, Roche Molecular Biochemicals, Indianapolis,
IN)-mediated method for 612 h. Cells were harvested 48 h after
transfection for assay of luciferase activity. Each experiment
represented at least two sets of independent triplicates to overcome
the variability inherent in transfection experiments.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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This investigation was supported by the National Health and Medical Research Council (NHMRC) of Australia. G.E.O.M. is an NHMRC Principal Research Fellow. The Centre for Molecular and Cellular Biology, IMB, is part of the Special Research Centre for Functional and Applied Genomics that is supported by the Australia Research Council (ARC).
Received for publication October 26, 2000. Revision received February 19, 2001. Accepted for publication February 20, 2001.
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REFERENCES |
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