|
|
J Biol Chem. 2008 May 2; 283(18): 11935–11946. doi: 10.1074/jbc.M707389200. | PMCID: PMC2335360 |
Copyright © 2008, The American Society for Biochemistry and
Molecular Biology, Inc. Nuclear Respiratory Factor 1 Controls Myocyte Enhancer Factor 2A
Transcription to Provide a Mechanism for Coordinate Expression of Respiratory
Chain
Subunits *![[S with combining enclosing square]](/corehtml/pmc/pmcents/x0053x20DE.gif) Bindu Ramachandran,‡1 Gengsheng Yu,‡§1,2 and Tod Gulick‡§3 ‡Diabetes Research Laboratory, Department of Medicine, Massachusetts General Hospital, Charlestown, Massachusetts 02129 and the §Department of Medicine, Harvard Medical School, Boston, Massachusetts 02115 Received September 4, 2007; Revised January 24, 2008. Nuclear respiratory factors NRF1 and NRF2 regulate the expression of
nuclear genes encoding heme biosynthetic enzymes, proteins required for
mitochondrial genome transcription and protein import, and numerous
respiratory chain subunits. NRFs thereby coordinate the expression of nuclear
and mitochondrial genes relevant to mitochondrial biogenesis and respiration.
Only two of the nuclear-encoded respiratory chain subunits have evolutionarily
conserved tissue-specific forms: the cytochrome c oxidase (COX)
subunits VIa and VIIa heart/muscle (H) and ubiquitous (L) isoforms. We used
genome comparisons to conclude that the promoter regions of
COX6AH and COX7AH lack NRF sites but
have conserved myocyte enhancer factor 2 (MEF2) elements. We show that
MEF2A mRNA is induced with forced expression of NRF1 and that the
MEF2A 5′-regulatory region contains an evolutionarily conserved
canonical element that binds endogenous NRF1 in chromatin immunoprecipitation
(ChIP) assays. NRF1 regulates MEF2A promoter-reporters according to
overexpression, RNA interference underexpression, and promoter element
mutation studies. As there are four mammalian MEF2 isotypes, we used
an isoform-specific antibody in ChIP to confirm MEF2A binding to the
COX6AH promoter. These findings support a role for
MEF2A as an intermediary in coordinating respiratory chain subunit
expression in heart and muscle through a NRF1 → MEF2A →
COXH transcriptional cascade. MEF2A also bound the
MEF2A and PPARGC1A promoters in ChIP, placing it within a
feedback loop with PGC1α in controlling NRF1 activity. Interruption of
this cascade and loop may account for striated muscle mitochondrial defects in
mef2a null mice. Our findings also account for the previously
described indirect regulation by NRF1 of other MEF2 targets in muscle such as
GLUT4. The electron transport chain
(ETC) 4 consists of
four multisubunit enzyme complexes within the inner mitochondrial (mito)
membrane. These act in concert to transfer electrons from succinate or NADH to
molecular oxygen while pumping protons from the matrix to the intermembranous
space, establishing the electrochemical gradient required for oxidative
phosphorylation (OXPHOS) ( 1).
Nuclear genes encode all of the components of complex II, but the other
complexes have subunits encoded by both mito ( ETCmito) and
nuclear ( ETCnucl) genes
( 1,
2). Appropriate ETC subunit
stoichiometry requires the coordinate expression of genes on the two genomes
and an accounting for a variable number of mito genomes per cell
( 2,
3). This is orchestrated by the
nuclear respiratory (transcription) factors, NRF1 and NRF2
( 2– 5).
These structurally unrelated factors, encoded by nuclear genes, regulate the
transcription of TFAM, TFB1M, and TFB2M, nuclear genes of
the mito transcription factor Tfam (mtTFA)
( 6) and Tfbm specificity
factors ( 7). Tfam and Tfbm
proteins are imported into mito where they direct transcription from both
heavy and light strands of mito DNA (mtDNA). These transcripts are processed
to yield the various ETCmito mRNAs, as well as rRNAs,
tRNAs, and a primer for the RNA-dependent activity of DNA polymerase γ
and mtDNA replication. This NRF → [ TFAM, TFBM] →
ETCmito transcriptional cascade functions in parallel with
the direct control of promoters of overlapping sets of
ETCnucl genes by NRF1 and NRF2
( 2,
3,
8). As NRFs also regulate
nuclear genes encoding complex V (F 1F 0-ATPase) subunits,
heme biosynthetic enzymes, and mito protein import machinery
( 2,
3,
9,
10), they are the central
regulators of mito biogenesis and cellular respiration. This is underscored by
the mito deficiency and peri-implant lethality of nrf1 null mouse
embryos ( 11). Although the entire ETC can be regarded as functioning as a single unit,
the reaction catalyzed by cytochrome c oxidase (COX, complex IV)
involves the largest free energy change among ETC reactions
( 1). Subunits I, II, and III
that together form the catalytic core of COX are encoded in the mito genome,
whereas the remaining 10 subunits serve structural or regulatory functions and
are products of nuclear genes. Among ETC enzymes, COX alone has
tissue-specific subunit isoforms that are the products of separate nuclear
genes. Thus, there are both ubiquitous (L, for liver) and heart and
muscle-specific (H) isoforms of subunits VIa and
VIIa 5 in all mammals,
and some species also have L and H variants of COX VIII
( 2,
8,
12). Multiple COX VIa and VIIa
isoforms and cognate genes are also present in lower metazoan species
including Drosophila
( 13). Biochemical studies have
suggested that COX H may confer sensitivity of muscle COX activity
and respiration to cellular energy demands
( 14). In this study, we began with the question as to how the COX H
subunit isoforms are coordinately expressed with one another in a
tissue-specific manner but also in harmony with other nuclear-encoded proteins
that are critical to mito function in muscle. We show that MEF2A, the
gene encoding the transcription factor myocyte enhancer factor 2A (MEF2A), is
a target of NRF1 regulation. COXH subunit genes are in
turn targets of MEF2 ( 15,
16), and we show binding of
endogenous myocyte MEF2A protein to the cox6aH gene
promoter. We therefore propose that a transcriptional cascade exists (NRF1
→ MEF2A → COXH) that functions with the
NRF → [ TFAM, TFBM] → ETCmito cascade
and direct NRF → ETCnucl regulation to provide
coordinate control of respiratory chain component expression in muscle. Our
observations also place MEF2A with NRF1 and PGC1α in a mutually
reinforcing transcriptional network. RNA Analyses—Human tissue RNA was obtained from Ambion.
Murine tissue and C2C12 and 10T1/2 cell RNA was isolated as described
( 17). Ribonuclease protection
assays (RPA) and radiolabeled cRNA probe syntheses were carried out as
described ( 17,
18). Human MEF2A and
murine mef2a, mef2c, and mef2d cRNA probes have been
described
( 17– 19).
Templates for other cRNA probes were PCR amplicon fragments subcloned into
pBluescript, oriented to permit cRNA production from the vector T7 promoter.
The 149-bp template for murine nrf1 cRNA used mouse heart RNA and
reverse transcription-PCR with primers
5′-ccc ggatCCCAGGCTCAGCTTCGGGCA-3′ and
5′-ccc gaattcGCTCTTCTGTGCGGACATCAC-3′. The underlined
letters are restriction sites used in subcloning. The uppercase letters are
cDNA sequences lowercase are extraneous to the cDNA but used to generate the
PCR amplicon and restriction site. Templates for murine cox subunits
were made using PCR on expressed sequence-tagged cDNAs or reverse
transcription-PCR using liver and heart RNA. Primers and expressed sequence
tag clones were: cox6aL (143 bp) IMAGE clone 3487598,
5′-gggcgc ggatcCTCGGATGTGGAAGGCCCTC-3′ and
5′-gggcgc gaattCTTGGTCCTGATGCGCAGG-3′;
cox6aH (120 bp) IMAGE clone 695934,
5′-gggcgc ggatCCAACACCTGGCGCCTC-3′ and
5′-gggcgc gaattcGGTGGTGATACGGGATGAAC-3′;
cox7aL (206 bp) 129 strain heart cDNA and IMAGE clone
1248366, 5′-gggcgc ggatccGAGGATAATGGGATGCCAG-3′ and
5′-gggcgc gaatTCAGATTCCTGGTCCATCG-3′;
cox7aH (171 bp) IMAGE clone 463628,
5′-gggcgc ggaTCCCAGGCTCTGGTCCGG-3′ and
5′-gggcgc gaattcGCCCCCCAGAGTCAGCGTC-3′;
cox7aR (102 bp) IMAGE clone 678445,
5′-gggcgc ggaTCCAGAAGGCTGATGGTTTCC-3′ and
5′-gggcgc gaatTCAGGCAGTAGATGGTCCCTC-3′. Reverse transcriptase reactions (Promega) used 1 μg of total RNA and an
oligo(dT) primer. Real-time PCR was performed using a MX-3000 multiplex
thermal cycler and SYBR Green with reaction conditions according to the master
mix reagent supplier (Stratagene). Quantitative polymerase chain reaction
(QPCR) primers for human cDNAs were: (GAPDH, 238 bp)
5′-GAGTCAACGGATTTGGTCGT-3′ and
5′-TTGATTTTGGAGGGATCTCG-3′; (NRF1, 281 bp)
5′-GTACAAGAGCATGATCCTGGA-3′ and
5′-GCTCTTCTGTGCGGACATC-3′; (MEF2A, 187 bp)
5′-GTGTACTCAGCAATGCCGAC-3′ and
5′-AACCCTGAGATAACTGCCCTC-3′; (CYCS, 106 bp)
5′-GTTGAAAAGGGAGGCAAGCA-3′ and
5′-TGTTCTTATTGGCGGCTGTG-3′; and (ACTB, 233 bp)
5′-GGACTTCGAGCAAGAGATGG-3′ and
5′-AGCACTGTGTTGGCGTACAG-3′. QPCR primers for murine cDNAs were:
(gapdh, 223 bp) 5′-CTGGAGAAACCTGCCAAGTA-3′ and
5′-TGTTGCTGTAGCCGTATTCA-3′; (cycs, 177 bp)
5′-TTCAGAAGTGTGCCCAGTGC-3′ and
5′-CTCCAAATACTCCATCAGGGTATC-3′; (cox5b, 217 bp)
5′-CAAGGTTACTTCGCGGAGTG-3′ and
5′-TCCTTGGTGCCTGAAGCTG-3′; and (actb, 160 bp)
5′-AAGAGCTATGAGCTGCCTGA-3′ and
5′-TACGGATGTCAACGTCACAC-3′. Mouse mef2a and nrf1
primer pairs were identical to the respective human MEF2A and
NRF1 primers. Real-time PCR results were analyzed using MX-3000
software, and the various mRNA quantities were normalized to that of
β-actin (ACTB or actb). In brief, the fractional
difference in the expression of a gene of interest (goi) mRNA in
experimental (e) versus control (c) samples was
determined using the formula ΔmRNA (arbitrary units) =
2{Δgoi
–Δactb}, where
Δgoi is the difference ([e] – [c]) in
cycle number at critical threshold for the goi and
Δactb is the difference ([e] – [c]) in cycle number at
critical threshold for actb. Fluorescence determinations were also
used to establish conditions (cycle number) to retrieve aliquots of parallel
reactions for direct visualization of amplicon levels by agarose gel
electrophoresis. All reported results were repeated in three independent small
interfering RNA (siRNA) transfections. Reporter Plasmids—p tkLuc and the
MEF2Ap 1-Luc and MEF2Ap 2-Luc deletion
series and p 1[m1 MEF2]-Luc and
p 2[m1 MEF2]-Luc have been described
( 19). tata-Luc (Gift
of Grace Gill) has been referred to as E1BLuc
( 20).
p 1[m1 NRF1]-Luc and p 1[m2 NRF1]-Luc
were created using PCR mutagenesis, exploiting the MEF2A promoter
FspI site within the NRF1 element ( TGC ↓ GCACGCGCA). Mutations
corresponded to those used in mobility shift assay probes
(). A reverse
luciferase primer was used in combination with 5′-gggcgc tgg ↓
cCACGgGCAGCACA-3′ (m1 NRF1) or 5′-gggcgc ttt
↓ aaACttGCAGCACA-3′ (m2 NRF1) on a
MEF2A-Luc template. Amplicons were cut with NcoI and either MscI
(m1 NRF1) or DraI (m2 NRF1), and the NcoI-blunt fragments
were substituted into FspI-and NcoI-cut S- MEF2Ap 1-Luc. A
BamHI-XbaI fragment was substituted into S- MEF2Ap 2-Luc to
give p 2[m1 NRF1]-Luc. This strategy was also used with
p 1[m1 MEF2]-Luc and p 2[m1 MEF2]-Luc
( 19) to construct reporters
containing dual NRF1 and MEF2 element mutations,
p 1[m1 MEF2/m1 NRF1]-Luc and
p 2[m1 MEF2/m1 NRF1]-Luc. II-E-Luc was made by
isolating and subcloning a RPCI-98-01L01 BAC clone EcoRI fragment, using this
as a template with forward vector and reverse
(5′-gggcgc gtcgacTCTCCTCTCATTGCGTTTTCC-3′) primers in
PCR, and subcloning the EcoRI- and SalI-restricted amplicon into
tata-Luc. An analogous construct containing an EWG element mutation,
II-E[m1 EWG]-Luc, was made by combining PCR with vector primers and
5′-GGTTTTT TGCc CATGg GCTCCTTCCCCACAG-3′ and
5′-GAAGG AGCc CATGg GCAAAAAACCGTAAACTC-3′. Bold
letters indicate the binding site for the relevant transcription factor. Eukaryotic Expression Vectors—pCDNA-MEF2A α2/β,
-MEF2C α2/β, and -MEF2D α2/β have been described
( 17,
18). pAc, pAc-EWG, and
pAc-EWG ΔN144 were gifts of Grace Gill (Tufts University) and
have been described ( 20). The
human NRF1 coding region was PCR-amplified from IMAGE cDNA clone 591311
template using primers (forward)
5′-gggcgc aagcttgccaccATGGAGGAACACGGAGTGAC-3′ and
(reverse) 5′-gaggtg gcggccgcttcaCTGTTCCAATGTCACCAC-3′,
and the HindIII- and NotI-restricted amplicon was subcloned to make
pCDNA-NRF1. pCDNA-NRF1 ΔN85 was made using PCR on this
template with forward primer
5′-gggcgc aagcttgccaccATGGCAACAGGAAAGAAACG-3′ and a
reverse vector primer followed by substitution of a HdIII- and
EcoRI-restricted amplicon. pCDNA-NRF1 VP16 was constructed using PCR
with reverse primer
5′-gaggcgc cctgcaggCTGTTCCAATGTCACCACC-3′ to delete the
NRF1 stop codon and then reintroducing the coding region into a
modified pCDNA3 containing sequences encoding the herpes simplex virus VP16
transactivation domain downstream of an SbfI site. pCDNA3-EWG and
-EWG ΔN144 and pAc-NRF1 and -NRF1 ΔN85 were
constructed by swapping coding regions between pAc and pCDNA vectors. A
derivative of pCDNA3 containing a viral internal ribosome entry site and
enhanced green fluorescent protein was used in pCDNA-NRF1 VP16/GFP.
pCDNA-PGC1α included the IMAGE clone 30094033 SalI to NotI insert. Mobility Shift Assays—The NRF1 coding region from pCDNA-NRF1
was subcloned into pET28 (Novagen) to give pET-NRF1. pET-NRF1 myc
was created by substituting a NRF1 insert lacking a stop codon from
pCDNA-NRF1 VP16 with a SbfI to NotI fragment containing sequence
encoding the c- myc epitope (EQKLISEEDLN) and a stop codon. In
vitro transcription/translation reactions used these plasmid templates
with the T nT system (Promega). Muscle cell nuclear extracts were
prepared, and electrophoretic mobility shift assays were conducted as
described previously ( 19).
Probe and competitor oligo sequences were: MEF2A NRF1,
5′-gatccGGTAG TGCGCACGCGCAGCACAa-3′ and
5′-gatctTGTGC TGCGCGTGCGCACTACCg-3′; MEF2A NRF1
m1 NRF1,
5′-gatccGGTAG TGCc CACGg GCAGCACAa-3′ and
5′-gatctTGTGC TGCc CGTGg GCACTACCg-3′;
MEF2A NRF1 m2 NRF1,
5′-gatccGGTAG TGCaa ACtt GCAGCACAa-3′ and
5′-gatctTGTGC TGCaa GTtt GCACTACCg-3′;
DMef2 EWG, 5′-gatccTTTTT TGCGCATGCGCTCCTTCa-3′ and
5′-gatctGAAGG AGCGCATGCGCAAAAAAg-3′; and MEF2C
NRF1, 5′-gatccTTCGG TGCGCGCGCGAATGCGCAAGCCCa-3′ and
5′-gatctGGGCT TGCGCATTCGCGCGCGCACCGAAg-3′. Chromatin Immunoprecipitation—Three 15-cm dishes of C2C12 or
HEK 293 cells were grown to confluence prior to cross-linking in
situ, nuclear isolation, DNA shearing, preclearing, and
immunoprecipitation. Procedures used were modified from the ChIP-IT kit
(Active Motif) using enzymatic DNA shearing as described previously in detail
( 19). Results were taken only
when validated with both positive and negative precipitation and PCR controls.
Processing of three independent cross-linked samples gave similar results.
Primers for human sequences were: chromosome (chr.) 12 intergenic (174 bp),
5′-atggttgccactggggatct-3′ and
5′-tgccaaagcctaggggaaga-3′; GAPDH, chr. 12 (166 bp),
5′-tactagcggttttacgggcg-3′ and
5′-tcgaacaggaggagcagagagcga-3′; MEF2A, chr. 15 (199 bp)
5′-accgagaggataattcagtcctg-3′ and
5′-acatccgcgcacggatc-3′; PPARGC1A, chr. 4 (204 bp),
5′-gagatggacaatgaagaacagtg-3′ and
5′-agttcccaggagatgtacacg-3′; and GLUT4, chr. 17 (242 bp),
5′-aaggcgtcatctccctgtc-3′ and
5′-aactctgcgggtctggac-3′. Primers for murine sequences were:
intergenic, chr. 6 (248 bp), 5′-aacctcatggttgccacag-3′ and
5′-accacgagatctgtaggcaag-3′; gapdh, chr. 6 (207 bp),
5′-agctactcgcggctttacg-3′ and
5′-tcacctggcactgcacaag-3′; mef2a, chr. 7 (233 bp),
5′-accgagagcagaaatatacccta-3′ and
5′-gagccgcctccttcagc-3′; ppargc1a, chr. 5 (124 bp),
5′-gagcacattaaattaacctcagtgg-3′ and
5′-ccagctcatttcctttacttgac-3′; glut4, chr. 11 (215 bp),
5′-taaggctccatctcctttgc-3′ and
5′-gtatgggctacatgtacttgcc-3′; and ( cox6ah, chr. 7, (179
bp) 5′-ggatctcctgccagtcaagac-3′ and
5′-ttagaggcagagccattgtca-3′. Immunoprecipitations used rabbit
anti-NRF1 (Abcam, ab34682), mouse anti-RNA polymerase II (Upstate Biologicals,
clone 8WG16), rabbit anti-MEF2A
( 19), or control IgG. Cultured Cell Transfection—C2C12 cells were maintained and
differentiated as described
( 17– 19).
HeLa and HEK 293 cells were maintained in Dulbecco's modified Eagle's medium
with 10% fetal calf serum. S2 cells were maintained at 25 °C in DS2 medium
with 10% fetal bovine serum. HEK 293 cells in 15-cm dishes were transfected
with 10 μgof pCDNA-NRF1 VP16/GFP prior to GFP +
versus GFP – fluorescence-activated cell sorting
after 48 h. Promoter reporter assays in mammalian cells in 12-well plates were
performed as described using Superfect (Qiagen)
( 19). S2 cells in 6-well
plates were transfected at 50% confluence. Triplicate wells received 3 μg
of reporter plasmid and 1 μg of expression vector, and cells were harvested
for luciferase assays after 48 h. 10T1/2 cells stably transformed with a
MyoD-ER fusion were maintained in medium with G418 and 10% charcoal-stripped
fetal bovine serum, and myogenesis was induced using 10 n m
estradiol as described
( 17). RNA Interference—siRNA transfections were performed using
HiPerFect (Qiagen) according to the supplier instructions. C2C12 cells were
transfected every 24 h for 2 days with subdivisions to maintain cell
subconfluence, and RNA was harvested after a total elapsed time of 72 h. HEK
293 cells were transfected once 48 h before RNA harvesting at 80% cell
confluence. 75 ng/well (12-well plate) or 150 ng/well (6-well plate) of dsRNA
was used in transfections. Double-stranded siRNAs all had 19 bp corresponding
to the targeted mRNA and 3′ UU extensions on each strand. Human siRNA
sense sequences were: GAPDH, 5′-GUCAACGGAUUUGGUCGUAUU-3′;
NRF1, 5′-GAAACGGCCUCAUGUAUUUUU-3′,
5′-UAGUAUAGCUCAUCUUGUAUU-3′,
5′-CACAUUGGCUGAUGCUUCAUU-3′, and
5′-GCUAUUGUCCUCUGUAUCUUU-3′. Mouse siRNA sequences were:
gapdh, 5′-GUGUGAACCACGAGAAAUAUUUU-3′; nrf1,
5′-GAAUGAACGCCACCGAUUUUU-3′,
5′-CAGUAUAGCUCAUCUCGUAUU-3′,
5′-UGAAAUAAGCCUCCCGAUAUU-3′, and
5′-CAACAGGGAAGAAACGGAAUU-3′. A control siRNA that fails to
recognize any human or mouse targets (Qiagen catalog No. 1027280) was used as
a negative control. An Alex Fluor 488-labeled negative control ds-siRNA
(Qiagen) was used initially to optimize transfection conditions. Miscellaneous Reagents and Procedures—All plasmid segments
derived from PCR were verified by dideoxy sequencing
( 21). Mouse anti- myc
epitope (clone 9E10) was from United States Biologicals, and anti-NRF1
antiserum was the generous gift of Richard Scarpula (Northwestern). MEF2 Regulatory Elements Are Common to Heart/Muscle-specific
Respiratory Chain Subunit Gene Promoters—We examined the
distribution of cox6a and cox7a isoform mRNA in adult mouse
tissues using RPA to discriminate between the highly similar mRNA sequences.
Consistent with previous reports
( 22),
cox6aH was present exclusively in skeletal muscle and
heart, whereas cox6aL was expressed in all other tissues
examined ().
cox7aH was also expressed exclusively in striated muscle,
and although cox7aL was highest in non-muscle tissues, it
was present in skeletal muscle and quite abundant in heart.
cox7aR, a third isoform of this subunit
( 23), was expressed
ubiquitously at a relatively low level. Using 10T1/2 MyoD-ER cells wherein
myogenesis is induced by estrogen
( 24), isoform switching from
cox6aL to cox6aH coincided with the
appearance of a myosin heavy chain skeletal muscle differentiation marker
( not shown). The COX6A, COX7A, and COX8 (H and L forms of each) are
compact genes, each having a single promoter and splicing pattern and one
initiation codon within the first exon
( 8). We compared
5′-flanking region sequences of each of these genes among mammals to
identify conserved elements (). Transgenic promoter-reporter studies in mice have shown that
MEF2 and E box elements provide heart- and muscle-specific expression of mouse
cox6aH
( 16). The corresponding region
of other mammalian COX6AH genes is highly conserved
( 25). Specifically, the region
has either one canonical MEF2 element or two sites separated by 7–10 bp
(supplemental Table S1). COX7AH and
COX8H gene regulatory regions also have functional MEF2
elements ( 15,
16). No sequences
corresponding to NRF1 or NRF2 elements were detected in
COXH promoters. By contrast, both the
COX6AL and COX7AL promoters have
putative or documented NRF1 or NRF2 elements
( 2,
3,
8), but none has a MEF2 site.
Although the COX7AR gene promoter region has not been
functionally characterized, we found NRF1 and MEF2 elements within 1.2 kb of
the transcription start site (TSS). No other transcription factor binding
sites that are conserved and selectively present in the COXH
versus COXL promoters were found. Forced Expression of NRF1 Induces MEF2A Expression—Four
mammalian MEF2 genes encode MEF2
factors 6
( 18,
26). These genes have
overlapping but distinct patterns of expression, and the respective protein
products also have both unique and redundant functions
( 26,
27). Our in silico
analyses and the prior mouse cox6ah promoter-reporter transgene
studies ( 16) led us to suspect
that COXH transcription is regulated by one or more MEF2
factors. Coordinated expression with other ETC components could then be given
by a NRF if it controls expression of the cognate MEF2 gene(s).
nrf1 expression is known to precede expression of the mef2
genes during mouse embryonic development and to be present in differentiating
muscle ( 11), establishing one
prerequisite for such a cascade. Overexpression of native NRF1 does not typically give robust activation of
target genes either in vivo
( 28) or in cultured mammalian
cells ( 2). We therefore forced
expression of a NRF1 fusion to the strong transactivating domain from viral
VP16 (NRF1 VP16) to determine whether MEF2 gene expression
might be controlled by NRF1. Enhanced green fluorescent protein was
co-expressed with NRF1 VP16 from a viral promoter using an internal
ribosome entry site between the fusion and marker coding sequences
(). Human HEK
293 cells were used because they express all MEF2 isotype mRNAs at
modest levels under control conditions
( 29). Transfected cells
(GFP +) were separated from untransfected cells
(GFP –) by flow cytometry, and gene expression was compared in
the populations. MEF2A mRNA was strongly induced in the
NRF1 VP16/GFP + cells as confirmed by RPA
(), implicating
MEF2A as a target of NRF1 regulation. MEF2A Promoter 1 Contains a Conserved Canonical NRF1
Element—We mapped the 5′-regulatory region of human
MEF2A. The gene has two closely approximated alternative first exons,
A 1 and A 2, and cognate promoters ~65 kb upstream of
the first coding exon 1 (). A MEF2 element overlies the major promoter 1 TSS
(TSS p 1), and we have shown that this site provides for
MEF2A transcriptional autoregulation
( 19). Of particular interest
to our hypothesis, a canonical NRF1
( 2,
4) element is centered at
–47 relative to TSS p 1
( and
supplemental Fig. S1). Two E boxes and two potential NRF2 sites
( 2,
5) are located further upstream
( and
supplemental Fig. S1). The MEF2A 5′-regulatory region within
~0.8 kb upstream of TSS p 1 is highly conserved among mammals,
and the MEF2, NRF, and E box elements are each conserved in sequence and
location (supplemental Fig. S1). The proximal promoter 1 location of the
putative NRF1 element and its evolutionary conservation strongly suggested
functional relevance. We performed both mobility shift and chromatin immunoprecipitation (ChIP)
assays to verify NRF1 binding to the putative MEF2A promoter element.
In the former, one specific retarded complex was formed in resolved binding
reactions of recombinant epitope-tagged NRF1 (NRF1myc) and a dsDNA
probe containing the element (). This complex was not competed by otherwise analogous
probes containing either two (m1NRF1) or four (m2NRF1)
substitutions within the GC core repeats of the element
(). The complex
was supershifted in reactions containing anti-myc antibody,
confirming a recombinant NRF1 component. Specific complexes were also formed
with the probe and C2C12 myoblast and myotube nuclear protein extracts, and
these were supershifted in resolved reactions containing anti-NRF1 antibody
(). Endogenous nuclear NRF1 interaction with the MEF2A promoter
element was verified using ChIP. PCR primers were designed to amplify the
region surrounding TSS p 1 that includes the NRF1 and MEF2 elements
(), or a
downstream region. Anti-NRF1 antibody specifically co-precipitated the
MEF2A promoter fragment from HEK 293 (human) cells
(). Similar
findings were obtained in ChIP assays using C2C12 (murine) cells and primers
designed to detect the analogous region of mef2a (not shown). We had
previously shown that MEF2A factors also bind this promoter region
( 19). NRF1 is therefore
positioned to directly regulate MEF2A transcription and to influence
its autoregulation. Forced NRF1 Expression Induces MEF2A Promoter Activity—To
evaluate the influence of the NRF1 element and factor on MEF2A
promoter 1 and 2 activities, we used the MEF2Ap 1-Luc and
MEF2Ap 2-Luc reporters, respectively
( 19)
(). In general,
target gene promoters are only modestly sensitive to NRF1 overexpression in
mammalian cells ( 30). By
contrast, forced expression of NRF1 (or its ortholog, EWG) has been reported
to have robust activity in cultured Drosophila cells
( 20,
30). We therefore began by
examining the effects of NRF1 expression on the MEF2A reporters in
Drosophila S2 cells. Our focus on the NRF1 element led us to use the
S- MEF2Ap 1-Luc and S- MEF2Ap 2-Luc
constructs that contain only proximal promoter fragments in order to exclude
superfluous upstream enhancers
( 19). We expressed either
human NRF1 or its Drosophila ortholog, EWG, the product of erect
wing ( 20,
31), or a fusion or mutant of
either () from
the Drosophila β-actin promoter. In these studies,
MEF2Ap 1-Luc activity was induced 17-fold and
MEF2Ap 2-Luc 25-fold in cells expressing full-length NRF1
(). Forced
expression of EWG gave less potent effects, but an EWG mutant deleted for the
N terminus (EWG ΔN144)
( 20) activated both
MEF2A promoters to a similar extent as NRF1. These transcription
factors gave up to 400-fold stimulation of
[NRF1 MEF2A] 3- tata-Luc, a reporter
containing 3 copies of the NRF1 site, but had minimal effect on the parent
tata-Luc control. The MEF2A promoters are highly responsive
to forced expression of NRF1 in the nuclear milieu provided by this cell
type. To evaluate MEF2A promoter responses to NRF1 overexpression in
mammalian cells, reporter activity was studied in cells expressing intact
NRF1, a deletion mutant analogous to EWG ΔN144
(NRF1 ΔN86), or the NRF1 VP16 fusion. In HeLa cells,
NRF1 expression produced ~3-fold activation of
MEF2Ap 1-Luc and ~4-fold activation of
MEF2Ap 2-Luc (). Although modest, these responses are equal to or
greater than those reported for other promoter reporters in mammalian cells
( 7,
30). As expected, more robust
effects were produced by NRF1 VP16, which stimulated these reporters
12- and 15-fold, respectively. These studies suggested that NRF1 co-regulates
the MEF2A gene promoters through its proximal promoter 1 element. Myocyte mef2a mRNA Is Reduced with nrf1 RNA Interference—We
used RNAi to study the effects of nrf1 knockdown on expression of
endogenous mef2a mRNA in muscle cells. C2C12 cells were transfected
with dsRNA targeted to nrf1. Control cells received an siRNA that
targets gapdh, one that fails to hybridize with any known message, or
no dsRNA in a mock transfection. Transfection conditions were optimized using
Alexa Fluor 488-tagged control siRNA. Because myoblasts proliferate rapidly,
we performed serial transfections with a low concentration (5 nm)
of the siRNAs. Total cell RNA, harvested at cell confluence after three
transfections at doubling time intervals, was used in reverse transcriptase
reactions. Validated primer pairs specific for gapdh and
nrf1 were used in quantitative PCR, with values normalized to sample
β-actin (actb) mRNA levels. Messenger RNAs of these targets were
each selectively down-regulated to 8 and to 15% of control levels,
respectively, with transfection of the cognate siRNA, as can be seen in
stained electrophoretic gels of the reaction products
(). Levels of the
non-cognate siRNA-transfected cell and control cell mRNAs were not affected
and were indistinguishable from those in mock-transfected cells. Having established the conditions under which nrf1 expression in
myocytes could be down-regulated, we tested the consequences for
mef2a expression. As controls, we examined effects on the expression
of established targets of NRF1, including the cycs and cox5b
genes that encode the somatic form of cytochrome c and the cytochrome
c oxidase Vb subunit, respectively
( 4). Validated primer pairs for
mef2a, cycs, and cox5b were used, with the former pair
designed to detect all splicing variants of mef2a
( 18). In an experimental
series, the respective mef2a, cycs, and cox5b mRNA
expression levels were reduced in nrf1 siRNA-transfected cells to 33,
30, and 25% of control levels (). Together with the NRF1 overexpression study findings, this
clearly demonstrates a strong correlation between cellular NRF1 levels and
MEF2A gene expression. MEF2A Promoter Activity Is Sensitive to NRF1 Element Mutation and to
RNAi-mediated Down-regulation of nrf1 Expression—C2C12
transfections with a deletion series of both MEF2A
promoter-reporters, including short (S), intermediate (I), and long (L)
promoter constructs, showed that preferential activity in myocytes was
maintained in the (S) reporters
( 19). These constructs each
retain nearly 50% of full-length promoter function
(). MEF2A reporters with an NRF1 element
mutation corresponding to one that failed in binding competition assays
(p 1- and p 2-[m1 NRF1]-Luc) had markedly
compromised activity compared with the cognate wild-type promoter reporters,
consistent with positive control of MEF2A transcription from this
element. Similar results were obtained in HEK 293 cells ( not shown).
This element therefore appears to be crucial to full promoter activity in a
variety of cell types in which MEF2A is expressed. To confirm that RNAi-mediated NRF1 underexpression reduced mef2a
mRNA by a transcriptional mechanism, we evaluated MEF2A
promoter-reporter activity in C2C12 and HEK 293 cells. In each case, cells
were transfected with control or NRF1 siRNAs as for the mRNA studies followed
after 48 h by co-transfection of the S-MEF2Ap2-Luc
reporter. NRF1 knockdown led to significant reductions in MEF2A
promoter activity to 42 and 25% of controls in C2C12 and 293 cells,
respectively (). The rapid proliferation rate of these cells probably
means that this understates the impact of NRF1 down-regulation on
MEF2A promoter activity. Taken together with the NRF1 element
mutation studies, these results confirm that NRF1 plays a critical role in
MEF2A transcription in two diverse cell types, including
differentiating muscle cells. PGC1α Co-activates the MEF2A Promoters from the NRF1 and
MEF2 Elements—The close proximity of the MEF2A promoter
NRF1 and MEF2 elements suggested the possibility of an interaction. Indeed,
the activities of reporters with or without mutations in the MEF2 and NRF1
sites were consistent with element cooperation
( ).
Under conditions where activity of MEF2Ap 2-Luc was
~5-fold higher than that of p 2[m1 NRF1]-Luc and
3-fold higher than a reporter with a MEF2 site mutation
(p 2[m1 MEF2]-Luc)
( 19), it was nearly 20-fold
higher than a reporter with mutations in both elements. Forced expression of
MEF2A α2/β and NRF1 ΔN86 synergistically activated
the MEF2A promoter in HeLa cells, also suggesting cooperation of the
elements (). No
direct protein-protein interaction between MEF2 and NRF1 was noted in a
mammalian two-hybrid system ( not shown), eliminating one possible
mechanism for element synergy. NRF1 and MEF2 are each known to interact with PPARγ coactivator
1α (PGC1α)
( 32– 36).
This suggested that MEF2A transcription might be sensitive to the
abundance of this cofactor. We therefore examined the sensitivity of
MEF2A promoter activity to forced expression of this coactivator. As
shown in ,
PGC1α stimulated MEF2Ap 2-Luc activity ~4-fold in
C2C12 cells, which is similar to PGC1α activities on other promoters
such as those of the TFBM genes
( 7). This effect is mediated
from the NRF1 and MEF2 elements, as mutation of either site resulted in
attenuation in the PGC1α effect, and the response was nearly abolished
on p 2[m1 NRF1/m1 MEF2]-Luc. Co-recruitment of
PGC1α is one possible mechanism for cooperative control of
MEF2A transcription by MEF2 and NRF1. Endogenous Muscle Cell MEF2A Binds the Promoters of Genes Encoding COX
VIaH, PGC1α , and MEF2A—There are four
MEF2 isotype genes in all mammals. These genes have distinct but
overlapping expression patterns and functions
( 18,
26,
27). These genes encode either
one protein (MEF2B) 6 or multiple splicing variant proteins (MEF2A,
MEF2C, and MEF2D) that have a common N-terminal DNA-binding and dimerization
domain ( 17,
18), such that all MEF2
proteins can heterodimerize promiscuously. Expression of the MEF2
isotypes is induced at different stages of cultured myoblast differentiation,
but each is highly expressed in differentiated myotubes
( 18). We therefore used C2C12
myotubes as substrate for ChIP assays to confirm occupation of gene promoters
relevant to our hypothesis by MEF2A. We developed a highly specific anti-MEF2A antibody that recognizes all
splicing variants of MEF2A but no other MEF2 form. This antibody had been used
by us previously to demonstrate MEF2A binding to the MEF2 element in the
MEF2A promoter ( 19).
We performed ChIP assays with C2C12 myotubes to verify that MEF2A proteins
also bind the cox6aH and ppargc1a gene promoters
at the respective MEF2 element regions. The gapdh gene promoter,
which is not regulated or bound by MEF2 proteins, served as a negative
control. As shown in ,
MEF2A co-precipitated with chromatin containing the cox6aH
and ppargc1a gene fragments but failed to precipitate gapdh.
Thus, in this setting of co-expressed MEF2 isotype and splicing variant
proteins, MEF2A forms occupy the promoters of MEF2A, COXH, and
PPARGC1A. This work does not exclude co-occupation by other MEF2
forms. Mito density is particularly rich in cardiac and skeletal muscle, where
OXPHOS is brisk. The ETC is unique in these tissues by virtue of the
expression of tissue-specific complex IV subunits, COX VIa H and COX
VIIa H. Although the functions of these and other non-catalytic COX
subunits are incompletely understood, there is evidence to suggest that
COX H confer sensitivity of COX to the cellular energy state. In
specific, activity of COX isolated from striated muscle is activated by ADP,
whereas enzyme purified from other tissues is not
( 14). In addition, an antibody
that specifically recognizes COX VIa H neutralizes this ADP
sensitivity. It is therefore speculated that COX VIa H provides for
rapid adaptation of ETC activity to energy stores in muscle, where energy
demands can vary acutely and dramatically
( 1,
14). We explored a mechanism
that could orchestrate tissue-specific expression of COX H subunits
at levels commensurate with other ETC components and the mito and mtDNA
density in muscle. We found that COXH gene promoters have
evolutionarily conserved MEF2 elements, that myocyte MEF2A occupies the
cox6ah promoter, and that MEF2A expression is in turn
regulated by NRF1. We therefore propose that a NRF1 → MEF2A
→ COXH cascade functions in parallel with the NRF
→ TFAM → ETCmito and NRF →
ETCnucl pathways to control respiration in striated muscle
(). We have provided multiple lines of evidence that the closely approximated
alternative MEF2A promoters are co-regulated by NRF1, the first limb
of the proposed transcriptional cascade. We confirmed the evolutionary
conservation of a canonical NRF1 binding site in MEF2A promoter 1.
Both mobility shift assays with muscle cell nuclear extracts and myotube ChIP
assays verified endogenous nuclear NRF1 protein association with this
promoter. NRF1 homodimers recognize a 12-bp site, YGCGCAYGCGCR, and there is
little tolerance for sequence degeneracy
( 4). By consequence of this and
the presence of multiple CpG doublets, canonical NRF1 sites are predicted to
appear in a mammalian genome at a frequency of only 10 –8
bp. This location at –47 relative to the promoter 1 TSS (and ~600 bp
upstream of the promoter 2 TSS) and our DNA binding study results strongly
support a role for this site in regulating MEF2A transcription. We
saw no change in NRF1 binding to the site during myoblast differentiation
using either assay. This was not surprising because this factor is
constitutively bound to its DNA targets
( 37,
38). A second line of evidence in support of MEF2A as a target of NRF1
was provided by experiments using forced expression of NRF1, N-terminal
deletion mutants of NRF1 and its Drosophila ortholog EWG, or a
NRF1 VP16 fusion protein. We initially showed that the
MEF2A mRNA level in cultured HEK cells was strongly induced with
expression of NRF1 VP16. We used the VP16 transactivation domain
fusion in these exploratory studies because the forced expression of native
NRF1 is known to induce rather small changes in target gene expression. For
example, NRF1 overexpression led to only an ~2-fold increases in
TFAM promoter activity
( 32), and a synthetic minimal
promoter containing four NRF1 elements is typically used to demonstrate
transactivity ( 30,
32). Likewise, transgenic
expression of NRF1 in muscle to a level that exceeded the endogenous level by
a factor of 10 produced only 1.5- and 2-fold increases in cytochrome c
(cycs) and δ-aminolevulinic acid synthase (δ -alas)
mRNAs, respectively ( 28).
NRF1 VP16 was therefore a valuable tool in the detection of
MEF2A as a potential NRF1 target. Unlike the case in cultured
mammalian cells, exogenous expression of NRF1 or EWG has been shown by other
investigators to produce potent activation of target gene promoters in
Drosophila cells ( 20,
30). We were able to exploit
this model system to develop further evidence for the authenticity of
MEF2A as a target of NRF1. We also saw modest but significant effects
of NRF1 overexpression on MEF2A promoter-reporters in mammalian cells
that were similar to those reported for other promoters. Evidence that specifically addressed the role of NRF1 in controlling
MEF2A transcription in muscle came from studies of the function of
MEF2A promoter-reporters in C2C12 myocytes. Mutation of the NRF1
element produced a drastic decrease in reporter activity. Furthermore,
nrf1 RNAi in this and other cell types led to coincident reductions
in nrf1 and mef2a mRNA levels, as well as diminished
MEF2A promoter activity. We believe that the composite of evidence
reported here confirms that MEF2A transcription is controlled by
NRF1. MEF2A is expressed in neurons, adipocytes, in various immune
cell types, and in smooth muscle, in addition to skeletal and cardiac
myocytes. We have previously provided evidence that MEF2A has only
two promoters, p 1 and p 2, which do not have
tissue-specific activities
( 19). Because NRF1 is
ubiquitously expressed ( 39)
and we have shown here that p 1 and p 2 are coregulated by
this factor, NRF1 can probably control MEF2A expression in non-muscle
cells as well as in skeletal muscle and heart. The effect of NRF1
RNAi on MEF2A expression and promoter activity in HEK 293 cells
supports this contention. In differentiating mouse myoblasts in vitro, there is a quantum
burst in mef2a expression at the time of cell cycle withdrawal
( 18). There is recent evidence
that the transactivity of NRF1 is sensitive to cell cycle-regulated
phosphorylation ( 40). In
specific, Cdk4 or Cdk6, in association with the regulatory subunit cyclin D1,
can phosphorylate NRF1 Ser-47, which leads to diminished transactivity.
Disinhibition then occurs with cell cycle withdrawal because of the associated
reduction in cyclin D1 levels and NRF1 poS47.
Negative regulation of NRF1 activity by this mechanism is consistent with
previous studies of the EWG ΔN144 N-terminal deletion mutant
( 20), as well as with our
present findings with this and the analogous NRF1 ΔN86 mutant.
In each case, the mutants were more potent transactivators of NRF1-responsive
promoters than the full-length native factors in proliferating cells. We
speculate that cyclin D1/Cdk4 activity on NRF1 may account for or be involved
in the upsurge in mef2a expression noted at cell cycle withdrawal.
This stated, NRF1 transactivity and DNA binding are also positively
regulated by incompletely characterized serum-responsive phosphorylation
events ( 30,
38) and possibly in response
to cellular redox state ( 41).
In addition, this factor may repress transcription of some or all target genes
under some circumstances ( 37).
This suggests additional regulatory complexity with relevance to
MEF2A transcription that will require further study. nrf1 mRNA is detectable very early in embryonic development and is
ubiquitously expressed in developing and mature tissues
( 11,
39). Disruption of murine
nrf1 leads to embryonic failure at a very early stage (3.5–6.5
days postcoitus) ( 11), prior
to the appearance of mef2a during normal development. Tests of the
role of NRF1 in controlling temporal and spatial expression of MEF2A in
vivo will therefore require regulated interference with nrf1
expression and/or conditional nrf1 knockout, complemented by paired
transgenic MEF2A and MEF2A[m NRF1] promoter
studies. In addition to the NRF1 element, the MEF2A
5′-regulatory region has other conserved elements (supplemental Fig.
S1). A canonical MEF2 element overlies the major TSS of one of the alternative
TATA-less promoters, and MEF2 activity at this site confers transcriptional
autoregulation and sensitivity of MEF2A expression to stress
signaling ( 19). Additional
sites of potential relevance to both muscle-specific expression and mito
function are also present, including E boxes and putative NRF2 sites.
Explorations of the functions of each of these sites are progressing in our
laboratory. Despite evidence presented here that NRF1 controls MEF2A
expression as part of a transcriptional cascade, we cannot exclude a role for
other MEF2 genes in this network. Indeed, one of the several
MEF2C promoters does have conserved NRF1 elements (supplemental Fig.
S2), suggesting that this isotype may also be a target. MEF2C proteins may
therefore participate in the proposed transcriptional cascade in some cell
types or circumstances. However, neither MEF2C, MEF2B, nor
MEF2D mRNA was induced with forced expression of NRF1 VP16
under conditions where cultured cell MEF2A expression was strongly
stimulated. MEF2A may therefore play a unique role among the isotypes
as intermediary in communicating NRF1 activity. This is compatible with the
mef2a–/– mouse phenotype. These animals have
deficient and disorganized myocardial mito and a neonatal sudden death
syndrome, consistent with a defect in oxidative metabolism
( 42). Although it is clear that COXH expression is controlled
from promoter MEF2 elements, any of the four mammalian MEF2 isotype proteins
could conceivably act at these sites. We had previously found that available
anti-MEF2 antibodies do not display MEF2 isotype specificities
( 19). We used a new validated
isotype-specific antibody and myotube chromatin to demonstrate by ChIP that
endogenous MEF2A occupies the cox6ah promoter. Because disruption or
knockdown of one MEF2 gene results in the dysregulation of other
isotypes ( 42,
43), 6 we believe
that this is the best evidence for a direct regulation of
COXH genes by MEF2A, the second limb of our hypothesized
transcriptional cascade. mef2a is expressed in striated muscle
precursors in the developing embryo after day 9 post-coitus, and expression is
maintained in differentiated tissues after birth
( 26,
42,
44,
45). In differentiating mouse
myoblasts in vitro, mef2a expression increases dramatically upon cell
cycle withdrawal ( 18),
coincident with the appearance of coxH
( 22). Thus, MEF2A abundance in
muscle in vivo and in vitro closely mimics expression of the
heart/muscle forms of COX VIa and VIIa. This contrasts with the expression
pattern of mef2c, which is present at high levels during embryonic
development but subsequently diminishes drastically
( 26,
44,
45), and mef2d, which
is an early marker of the myogenic lineage and is maintained in differentiated
skeletal muscle
( 44– 46).
This stated, the MEF2A N-terminal MADS/MEF2 region that mediates dimerization
and sequence-specific DNA binding is shared among all MEF2 proteins
( 26). We therefore cannot
exclude co-regulation of the COXH genes by other MEF2
forms or by heterodimers of MEF2A with MEF2B, MEF2C, or MEF2D at different
stages of muscle development or differentiation. MEF2A is highly expressed in skeletal muscle and heart, but it is
also present at other sites including neural, adipose, and immune cells
( 17,
18,
26). Selective expression of
COXH genes may therefore require muscle-specific
activities of either MEF2A or its collaborating factors. Certain splicing
variants of MEF2A are expressed only in heart and
muscle 7
( 17,
18), providing one potential
mechanism for a muscle-specific MEF2A activity. Signaling that controls MEF2
protein or cofactor modifications may also be operative, particularly among
pathways regulated by intramyocellular Ca 2+ transients
( 47). Myogenic basic
helix-loop-helix factors could also contribute to muscle-specific expression,
because these factors can use DNA-bound MEF2 as scaffolding to transactivate
MEF2 target genes ( 48), and
these factors could also regulate MEF2A transcription from the
promoter E boxes. The strict specificity of COX6AH expression
suggests that gene silencing must also occur in non-muscle cells, the
mechanisms for which remain to be determined. The control of MEF2A transcription by NRF1 has implications beyond
the coordinated co-expression of ETC subunits in muscle. Specifically, other
genes that are regulated by MEF2A are implicated by our findings as potential
indirect targets of NRF1. The GLUT4 gene provides one important
example that has particular relevance to muscle and cardiac metabolism. Glut4
is the insulin-responsive facilitated glucose transporter form that provides
the major route for glucose uptake in striated muscle
( 49– 51).
Both cultured myocyte and transgenic promoter-reporter studies have
established that MEF2A regulates glut4 expression through a promoter
MEF2 element ( 49,
50). Muscle tissue
Glut4-level, insulin-stimulated glucose transport and MEF2A protein
concentration were each found to be increased ~2-fold in mice carrying a
transgene that overexpresses NRF1 in skeletal muscle compared with control
tissue ( 28). Our detection of
MEF2A as a target of NRF1 suggests the relevant mechanism, NRF1
→ mef2a → glut4, because glut4 is not a
direct target of NRF1. Given the tissue expression patterns of both
NRF1 and MEF2A, many more genes are likely to be regulated
by such a cascade in both muscle and non-muscle sites. PGC1α was originally identified as a coactivator of PPARγ and
NRF1 in brown adipose tissue, but it is now recognized as a key regulator of
mito biogenesis in various tissues
( 32,
52). PGC1α also
coactivates MEF2 factors ( 33,
35,
36), and its forced expression
in muscle increases mito density and OXPHOS and produces a
fast-twitch/glycolytic to slow-twitch/oxidative phenotype conversion
( 53). It is established that
PGC1α co-activates transcription from a PPARGC1A promoter MEF2
site ( 33,
35). We have shown here with
myotube ChIP that MEF2A factors bind this PPARGC1A element where they
are poised to recruit PGC1α in this autoregulatory loop. We have also
shown that PGC1α co-activates the MEF2A promoter from its MEF2
and NRF1 elements, confirming that PGC1α can also feed back on
MEF2A transcription. Taken together, this indicates that MEF2A,
NRF1, and PPARGC1A and their respective protein products form a
mutually reinforcing network of auto- and cross-regulation capable of
directing mito biogenesis and OXPHOS capacity in muscle
(). The NRF1
promoter has not been characterized, although the 5′-exons of human
NRF1 have been identified
( 54). There is a canonical
MEF2 binding site upstream of the human NRF1 TSS, 7
suggesting the possibility of direct reciprocal MEF2A and
NRF1 regulation. We are exploring whether this vector is also part of
this transcriptional network. The regulation of MEF2A transcription by NRF1 and PGC1α is
highly relevant to metabolic dysregulation in diabetes. Muscle glut4
mRNA, Glut4 protein, and insulin-stimulated glucose uptake are reduced in
animal models of diabetes, and there is a coincident down-regulation of
mef2a mRNA and MEF2A protein abundance
( 50,
51,
55). In these models, a
hypercatabolic state leads to a decline in the [AMP]/[ATP] ratio and a
coincident reduction in 5′-AMP-activated protein kinase (AMPK) activity
( 55– 58).
Because expression of both mef2a and of glut4 can be
restored with the administration of a small molecular activator of AMPK
( 55), this pathway is
implicated as a crucial sensor linking cell energy state with the capacity for
nutrient uptake and metabolism
( 55,
56). There is recent evidence
to suggest that AMPK activity regulates nuclear DNA binding activity of both
NRF1 ( 59) and MEF2
( 60). Neither factor appears
to be a direct target of AMPK
( 60). AMPK may therefore
target co-repressor(s) of one or both factors to promote dissociation and
de-repression of targets such as MEF2A
( 19,
61,
62). As one alternative, AMPK
activity could indirectly influence the subnuclear locus or co-activator
associations of these factors. In any case, down-regulated MEF2A
expression may be a primary mechanism by which NRF1 and PGC1α targets
are coordinately down-regulated in humans with diabetes and insulin resistance
( 63). We saw no evidence for a direct protein-protein interaction between NRF1
and MEF2A. These factors may cooperate in the recruitment of transcriptional
co-regulator(s) or the induction of chromatin remodeling to account for the
observed functional synergy. Paired MEF2 and NRF1 elements also exist in other
MEF2 gene regulatory regions, including the aforementioned
MEF2C promoter (supplemental Fig. S2) and the Drosophila
DMef2 II-E enhancer (supplemental Fig. S3). The DMef2
5′-region has various enhancers that govern the complex developmental
and spatial expression of the sole MEF2 gene in the fly
( 64). II-E is responsible for
transcriptional autoregulation
( 65) and for DMef2
expression near and after terminal differentiation of somatic muscle
( 64). We find that a canonical
EWG element in this region binds NRF1 and governs enhancer activity
(supplemental Fig. S3). Control of MEF2 transcription by NRF1 may
therefore be conserved among higher metazoans. The sea urchin NRF1 ortholog,
P3A2, directs territory-specific transcription of muscle genes during
embryonic development ( 66),
and EWG is known to regulate flight muscle development
( 31). Our work suggests a
potentially relevant mechanism and establishes a foundation for in
vivo functional analyses in various model systems to elucidate the
contributions of NRF1/EWG to the developmental and spatial expression of
MEF2 genes. We thank Grace Gill (Tufts University), Richard Scarpulla (Northwestern
University), and Stephen Tapscott (University of Washington) for reagents and
Shiguang Li and Pan Yin for technical assistance. 1. Scheffler, I. E. (1999. )
Mitochondria, pp.
141–245, Wiley-Liss, New York. 2. Scarpulla, R. C. (2006. ) J. Cell.
Biochem. 97
673–683 [PubMed] 3. Kelly, D. P., and Scarpulla, R. C. (2004. )
Genes Dev. 18
357–368 [PubMed] 4. Virbasius, C. A., Virbasius, J. V., and Scarpulla, R. C.
(1993. ) Genes Dev.
7
2431–2445 [PubMed] 5. Gugneja, S., Virbasius, J. V., and Scarpulla, R. C.
(1995. ) Mol. Cell. Biol.
15
102–111 [PubMed] 6. Virbasius, J. V., and Scarpulla, R. C. (1994. )
Proc. Natl. Acad. Sci. U. S. A.
91
1309–1313 [PubMed] 7. Gleyzer, N., Vercauteren, K., and Scarpulla, R. C.
(2005. ) Mol. Cell. Biol.
25
1354–1366 [PubMed] 8. Lenka, N., Vijayasarathy, C., Mullick, J., and Avadhani, N. G.
(1998. ) Prog. Nucleic Acid Res. Mol. Biol.
61
309–344 [PubMed] 9. Li, B., Holloszy, J. O., and Semenkovich, C. F. (1999. )
J. Biol. Chem. 274
17534–17540 [PubMed] 10. Chau, C. M., Evans, M. J., and Scarpulla, R. C. (1992. )
J. Biol. Chem. 267
6999–7006 [PubMed] 11. Huo, L., and Scarpulla, R. C. (2001. ) Mol.
Cell. Biol. 21
644–654 [PubMed] 12. Taanman, J. W., Hall, R. E., Tang, C., Marusich, M. F., Kennaway,
N. G., and Capaldi, R. A. (1993. ) Biochim. Biophys.
Acta 1225
95–100 [PubMed] 13. Tripoli, G., D'Elia, D., Barsanti, P., and Caggese, C.
(2005. ) Genome Biol.
6
R11.1–R11.7 [PubMed] 14. Anthony, G., Reimann, A., and Kadenbach, B. (1993. )
Proc. Natl. Acad. Sci. U. S. A.
90
1652–1656 [PubMed] 15. Yu, M., Jaradat, S. A., and Grossman, L. I. (2002. )
Biochim. Biophys. Acta
1574
345–353 [PubMed] 16. Wan, B., and Moreadith, R. W. (1995. ) J.
Biol. Chem. 270
26433–26440 [PubMed] 17. Zhu, B., and Gulick, T. (2004. ) Mol. Cell.
Biol. 24
8264–8275 [PubMed] 18. Zhu, B., Ramachandran, B., and Gulick, T. (2005. )
J. Biol. Chem. 280
28749–28760 [PubMed] 19. Ramachandran, B., Yu, G., Li, S., Zhu, B., and Gulick, T.
(2008. ) J. Biol. Chem.
283
10318–10329 [PubMed] 20. Fazio, I. K., Bolger, T. A., and Gill, G. (2001. )
J. Biol. Chem. 276
18710–18716 [PubMed] 21. Ausubel, F. M., Brent, R., Kingston, R. E., Moore, D. D., Seidman,
J. G., Smith, J. A., and Struhl, K. (1998. ) Current
Protocols in Molecular Biology, John Wiley & Sons, Inc., New
York. 22. Parsons, W. J., Williams, R. S., Shelton, J. M., Luo, Y., Kessler,
D. J., and Richardson, J. A. (1996. ) Am. J.
Physiol. 270
H567–H574 [PubMed] 23. Watanabe, T., Inoue, S., Hiroi, H., Orimo, A., Kawashima, H., and
Muramatsu, M. (1998. ) Mol. Cell. Biol.
18
442–449 [PubMed] 24. Bergstrom, D. A., Penn, B. H., Strand, A., Perry, R. L., Rudnicki,
M. A., and Tapscott, S. J. (2002. ) Mol.
Cell 9
587–600 [PubMed] 25. Bachman, N. J., Riggs, P. K., Siddiqui, N., Makris, G. J., Womack,
J. E., and Lomax, M. I. (1997. ) Genomics
42
146–151 [PubMed] 26. Black, B. L., and Olson, E. N. (1998. ) Annu.
Rev. Cell Dev. Biol. 14
167–196 [PubMed] 27. Shalizi, A. K., and Bonni, A. (2005. ) Curr.
Top. Dev. Biol. 69
239–266 [PubMed] 28. Baar, K., Song, Z., Semenkovich, C. F., Jones, T. E., Han, D. H.,
Nolte, L. A., Ojuka, E. O., Chen, M., and Holloszy, J. O. (2003. )
FASEB J. 17
1666–1673 [PubMed] 29. Zhao, M., New, L., Kravchenko, V. V., Kato, Y., Gram, H., di
Padova, F., Olson, E. N., Ulevitch, R. J., and Han, J. (1999. )
Mol. Cell. Biol. 19
21–30 [PubMed] 30. Herzig, R. P., Scacco, S., and Scarpulla, R. C. (2000. )
J. Biol. Chem. 275
13134–13141 [PubMed] 31. DeSimone, S., Coelho, C., Roy, S., VijayRaghavan, K., and White, K.
(1996. ) Development (Camb.)
122
31–39. 32. Wu, Z., Puigserver, P., Andersson, U., Zhang, C., Adelmant, G.,
Mootha, V., Troy, A., Cinti, S., Lowell, B., Scarpulla, R. C., and Spiegelman,
B. M. (1999. ) Cell
98
115–124 [PubMed] 33. Handschin, C., Rhee, J., Lin, J., Tarr, P. T., and Spiegelman, B.
M. (2003. ) Proc. Natl. Acad. Sci. U. S. A.
100
7111–7116 [PubMed] 34. Handschin, C., and Spiegelman, B. M. (2006. )
Endocr. Rev. 27
728–735 [PubMed] 35. Czubryt, M. P., McAnally, J., Fishman, G. I., and Olson, E. N.
(2003. ) Proc. Natl. Acad. Sci. U. S. A.
100
1711–1716 [PubMed] 36. Michael, L. F., Wu, Z., Cheatham, R. B., Puigserver, P., Adelmant,
G., Lehman, J. J., Kelly, D. P., and Spiegelman, B. M. (2001. )
Proc. Natl. Acad. Sci. U. S. A.
98
3820–3825 [PubMed] 37. Cam, H., Balciunaite, E., Blais, A., Spektor, A., Scarpulla, R. C.,
Young, R., Kluger, Y., and Dynlacht, B. D. (2004. ) Mol.
Cell 16
399–411 [PubMed] 38. Gugneja, S., and Scarpulla, R. C. (1997. ) J.
Biol. Chem. 272
18732–18739 [PubMed] 39. Gopalakrishnan, L., and Scarpulla, R. C. (1995. )
J. Biol. Chem. 270
18019–18025 [PubMed] 40. Wang, C., Li, Z., Lu, Y., Du, R., Katiyar, S., Yang, J., Fu, M.,
Leader, J. E., Quong, A., Novikoff, P. M., and Pestell, R. G.
(2006. ) Proc. Natl. Acad. Sci. U. S. A.
103
11567–11572 [PubMed] 41. Piantadosi, C. A., and Suliman, H. B. (2006. )
J. Biol. Chem. 281
324–333 [PubMed] 42. Naya, F. J., Black, B. L., Wu, H., Bassel-Duby, R., Richardson, J.
A., Hill, J. A., and Olson, E. N. (2002. ) Nat.
Med. 8
1303–1309 [PubMed] 43. Lin, Q., Schwarz, J., Bucana, C., and Olson, E. N.
(1997. ) Science
276
1404–1407 [PubMed] 44. Subramanian, S. V., and Nadal-Ginard, B. (1996. )
Mech. Dev. 57
103–112 [PubMed] 45. Edmondson, D. G., Lyons, G. E., Martin, J. F., and Olson, E. N.
(1994. ) Development (Camb.)
120
1251–1263 [PubMed] 46. Breitbart, R. E., Liang, C. S., Smoot, L. B., Laheru, D. A.,
Mahdavi, V., and Nadal-Ginard, B. (1993. ) Development
(Camb.) 118
1095–1106. 47. Wu, H., Naya, F. J., McKinsey, T. A., Mercer, B., Shelton, J. M.,
Chin, E. R., Simard, A. R., Michel, R. N., Bassel-Duby, R., Olson, E. N., and
Williams, R. S. (2000. ) EMBO J.
19
1963–1973 [PubMed] 48. Molkentin, J. D., Black, B. L., Martin, J. F., and Olson, E. N.
(1995. ) Cell
83
1125–1136 [PubMed] 49. Liu, M. L., Olson, A. L., Edgington, N. P., Moye-Rowley, W. S., and
Pessin, J. E. (1994. ) J. Biol. Chem.
269
28514–28521 [PubMed] 50. Mora, S., and Pessin, J. E. (2000. ) J. Biol.
Chem. 275
16323–16328 [PubMed] 51. Thai, M. V., Guruswamy, S., Cao, K. T., Pessin, J. E., and Olson,
A. L. (1998. ) J. Biol. Chem.
273
14285–14292 [PubMed] 52. Lin, J., Handschin, C., and Spiegelman, B. M. (2005. )
Cell Metab. 1
361–370 [PubMed] 53. Lin, J., Wu, H., Tarr, P. T., Zhang, C. Y., Wu, Z., Boss, O.,
Michael, L. F., Puigserver, P., Isotani, E., Olson, E. N., Lowell, B. B.,
Bassel-Duby, R., and Spiegelman, B. M. (2002. )
Nature 418
797–801 [PubMed] 54. Huo, L., and Scarpulla, R. C. (1999. )
Gene 233
213–224 [PubMed] 55. Finck, B. N., Bernal-Mizrachi, C., Han, D. H., Coleman, T.,
Sambandam, N., LaRiviere, L. L., Holloszy, J. O., Semenkovich, C. F., and
Kelly, D. P. (2005. ) Cell Metab.
1
133–144 [PubMed] 56. Zong, H., Ren, J. M., Young, L. H., Pypaert, M., Mu, J., Birnbaum,
M. J., and Shulman, G. I. (2002. ) Proc. Natl. Acad.
Sci. U. S. A. 99
15983–15987 [PubMed] 57. Cuthbertson, D. J., Babraj, J. A., Mustard, K. J., Towler, M. C.,
Green, K. A., Wackerhage, H., Leese, G. P., Baar, K., Thomason-Hughes, M.,
Sutherland, C., Hardie, D. G., and Rennie, M. J. (2007. )
Diabetes 56
2078–2084 [PubMed] 58. Towler, M. C., and Hardie, D. G. (2007. )
Circ. Res. 100
328–341 [PubMed] 59. Bergeron, R., Ren, J. M., Cadman, K. S., Moore, I. K., Perret, P.,
Pypaert, M., Young, L. H., Semenkovich, C. F., and Shulman, G. I.
(2001. ) Am. J. Physiol.
281
E1340–E1346. 60. Holmes, B. F., Sparling, D. P., Olson, A. L., Winder, W. W., and
Dohm, G. L. (2005. ) Am. J. Physiol.
289
E1071–E1076. 61. Berdeaux, R., Goebel, N., Banaszynski, L., Takemori, H., Wandless,
T., Shelton, G. D., and Montminy, M. (2007. ) Nat.
Med. 13
597–603 [PubMed] 62. van der Linden, A. M., Nolan, K. M., and Sengupta, P.
(2007. ) EMBO J.
26
358–370 [PubMed] 63. Patti, M. E., Butte, A. J., Crunkhorn, S., Cusi, K., Berria, R.,
Kashyap, S., Miyazaki, Y., Kohane, I., Costello, M., Saccone, R., Landaker, E.
J., Goldfine, A. B., Mun, E., DeFronzo, R., Finlayson, J., Kahn, C. R., and
Mandarino, L. J. (2003. ) Proc. Natl. Acad. Sci. U. S.
A. 100
8466–8471 [PubMed] 64. Nguyen, H. T., and Xu, X. (1998. ) Dev.
Biol. 204
550–566 [PubMed] 65. Cripps, R. M., Lovato, T. L., and Olson, E. N. (2004. )
Dev. Biol. 267
536–547 [PubMed] 66. Calzone, F. J., Hoog, C., Teplow, D. B., Cutting, A. E., Zeller, R.
W., Britten, R. J., and Davidson, E. H. (1991. )
Development (Camb.) 112
335–350.
|
