Download Embryonic activation of the myoD gene is regulated

Development 121, 637-649 (1995)
Printed in Great Britain © The Company of Biologists Limited 1995
637
Embryonic activation of the myoD gene is regulated by a highly conserved
distal control element
David J. Goldhamer1,*, Brian P. Brunk2,†, Alexander Faerman3, Ayala King3, Moshe Shani3
and Charles P. Emerson, Jr2,†
1Department of Cell and Developmental Biology, University of Pennsylvania
2Institute for Cancer Research, Fox Chase Cancer Center, Philadelphia, PA
3Institute of Animal Science, The Volcani Center, Bet Dagan 50250, Israel
School of Medicine, Philadelphia, PA 19104, USA
19111, USA
*Author for correspondence: Department of Cell and Developmental Biology, 219 Anatomy-Chemistry Building, University of Pennsylvania School of Medicine,
Philadelphia, PA 19104, USA. E mail: [email protected]
†Present address: Department of Cell and Developmental Biology, University of Pennsylvania School of Medicine, Philadelphia, PA 19104, USA
SUMMARY
MyoD belongs to a small family of basic helix-loop-helix
transcription factors implicated in skeletal muscle lineage
determination and differentiation. Previously, we identified
a transcriptional enhancer that regulates the embryonic
expression of the human myoD gene. This enhancer had
been localized to a 4 kb fragment located 18 to 22 kb
upstream of the myoD transcriptional start site. We now
present a molecular characterization of this enhancer.
Transgenic and transfection analyses localize the myoD
enhancer to a core sequence of 258 bp. In transgenic mice,
this enhancer directs expression of a lacZ reporter gene to
skeletal muscle compartments in a spatiotemporal pattern
indistinguishable from the normal myoD expression
domain, and distinct from expression patterns reported for
the other myogenic factors. In contrast to the myoD
promoter, the myoD enhancer shows striking conservation
between humans and mice both in its sequence and its
distal position. Furthermore, a myoD enhancer/heterologous promoter construct exhibits muscle-specific
expression in transgenic mice, demonstrating that the
myoD promoter is dispensable for myoD activation. With
the exception of E-boxes, the myoD enhancer has no
apparent sequence similarity with regulatory regions of
other characterized muscle-specific structural or regulatory genes. Mutation of these E-boxes, however, does not
affect the pattern of lacZ transgene expression, suggesting
that myoD activation in the embryo is E-box-independent.
DNase I protection assays reveal multiple nuclear protein
binding sites in the core enhancer, although none are
strictly muscle-specific. Interestingly, extracts from
myoblasts and 10TG fibroblasts yield identical protection
profiles, indicating a similar complement of enhancerbinding factors in muscle and this non-muscle cell type.
However, a clear difference exists between myoblasts and
10TG cells (and other non-muscle cell types) in the
chromatin structure of the chromosomal myoD core
enhancer, suggesting that the myoD enhancer is repressed
by epigenetic mechanisms in 10TG cells. These data indicate
that myoD activation is regulated at multiple levels by
mechanisms that are distinct from those controlling other
characterized muscle-specific genes.
INTRODUCTION
motifs known as E-boxes, sequences present in the regulatory
regions of most muscle-specific genes. Transfection assays
indicate that myoD family members also directly or indirectly
positively regulate each other’s expression (Thayer et al., 1989;
Braun et al., 1989; Edmondson et al., 1991, 1992). These crossand auto-regulatory functions may amplify expression of the
myogenic factors and stabilize the muscle phenotype (Thayer
et al., 1989). Importantly, these genes can induce myogenesis
in a variety of non-muscle cell types when expressed from a
constitutive promoter (reviewed by Emerson, 1990; Olson,
1990; Weintraub et al., 1991), a finding consistent with a
function in determination of the skeletal muscle lineage.
In vertebrates, skeletal muscle progenitor cells are derived
predominantly from the somites, which are formed by seg-
An understanding of the molecular mechanisms that govern the
determination and differentiation of skeletal muscle lineages
has progressed rapidly since the discovery of the myoD family
of myogenic regulatory genes. These genes, myoD (Davis et
al., 1987), myogenin (Edmondson and Olson, 1989; Wright et
al., 1989), myf5 (Braun et al., 1989) and MRF4 (Rhodes and
Konieczny, 1989; also known as myf6 [Braun et al., 1990] and
herculin [Miner and Wold, 1990]), which are expressed exclusively in skeletal muscle, encode structurally related transcription factors of the basic helix-loop-helix (bHLH) class (Murre
et al., 1989). These myogenic factors activate muscle-specific
gene transcription in differentiating cells by binding to DNA
Key words: myoD, transcription, myogenesis, basic helix-loop-helix
factors, transgenic mice
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D. J. Goldhamer and others
mentation of the paraxial mesoderm in a rostral-to-caudal
sequence. The myotome gives rise to axial muscles, whereas
trunk and limb muscles are derived from myogenic progenitor
cells that migrate away from the ventrolateral edge of the
maturing somite (Wachtler and Christ, 1992; Ordahl and Le
Douarin, 1992). Myogenic cells of the branchial arches, which
form facial, jaw and throat musculature, originate both from
anterior somites and from cranial paraxial mesoderm (Noden,
1991; Couly et al., 1992).
Gene-targeting experiments in mice showed that the elaboration of the myogenic phenotype is dependent on myogenic
regulatory gene function. Mice homozygous null for both
myoD and myf5 completely lack differentiated skeletal muscle
by both histological and biochemical criteria (Rudnicki et al.,
1993). In these mice, muscle-forming regions of the embryo
are devoid of myoblasts, as evidenced by the absence of
desmin-positive cells (Rudnicki et al., 1993). Interestingly,
mice homozygous null for either myf5 or myoD alone exhibit
no muscle defects at birth, indicating that the functions of
myoD and myf5 in myogenesis are at least partially redundant
(Braun et al., 1992; Rudnicki et al., 1992, 1993). In contrast,
although myogenin knockout mice show severe muscle differentiation defects, myoblasts are present in approximately
normal numbers and positions in these embryos (Hasty et al.,
1993; Nabeshima et al., 1993). These data indicate that myoD
and myf5 serve upstream functions in the myogenic developmental program to determine, expand, or maintain myogenic
lineages, whereas myogenin, although not required for commitment of cells to the myogenic lineage, is required for
normal biochemical and morphological differentiation of
skeletal muscle (reviewed by Weintraub, 1993).
Each of the myogenic genes is expressed in a unique spatiotemporal pattern in developing skeletal muscle (Buckingham, 1992; Faerman and Shani, 1993), indicating that
members of the myoD family are regulated by distinct developmental signals. Given the formative role of myoD and myf5
in establishing the skeletal muscle lineage (Rudnicki et al.,
1993), a mechanistic understanding of lineage determination
will require detailed information of how these genes are transcriptionally controlled. Cis and trans analysis of the myoD
enhancer offers a powerful means to define upstream signaling
pathways and transcriptional events that govern myoD activation in the embryo. Previously, we showed by transgenic
analyses that the embryonic expression of the human myoD
gene is regulated by a distant enhancer localized to a 4 kb
fragment approximately 18 to 22 kb upstream from the start of
myoD transcription (Goldhamer et al., 1992). In the present
study, we define and characterize the distal enhancer that
regulates the embryonic activation of the human myoD gene.
These data indicate that myoD activation is regulated by a
highly conserved and complex regulatory system that is
distinct from mechanisms that regulate the other myogenic regulatory genes.
DNA constructs
A 1.7 kb ApaI/PstI fragment derived from the 4 kb enhancer-containing fragment (fragment 3 in Goldhamer et al., 1992; see Fig. 2)
was subcloned into pBluescript KS+ (KS+; Stratagene) by blunt-end
ligation into the unique EcoRV site of the multiple cloning site. Bidirectional nested deletions were created by exonuclease III digestion
using the Erase-a-base kit (USB), or by partial digestion with PvuII
at nucleotide 95, to create ∆15 (3′-5′). ∆14KS+ (3′-5′) contains 258
base pairs (bp) of enhancer sequence, and is referred to as the core
enhancer throughout.
The reporter plasmid ptkCAT∆EH (derived from pBLCAT2
[Luckow and Schutz, 1987] by deletion of the NdeI/HindII fragment
of pUC 18) was the parental plasmid used for transfection experiments (Goldhamer et al., 1992). A 2.7 kb genomic fragment containing human myoD 5′ flanking sequences extending from an EcoRI site
at −2.5 kb to +198 relative to the TATA box (−37 relative to the translational initiation codon) was generated from a sequencing deletion
in KS+. −2.5CAT was produced by excising this promoter-containing fragment from KS+ by digestion with SacI and KpnI, followed by
blunt end-ligation into the XbaI and BglII sites of ptkCAT∆EH. (XbaI
and BglII digestion removes all thymidine kinase promoter
sequences.) myoD enhancer/promoter CAT constructs were generated
by digesting enhancer deletion constructs in KS+ with SalI and XbaI
and cloning into unique SalI and XbaI sites of −2.5CAT. (The XbaI
site was derived from KS+ during the construction of −2.5CAT.)
The lacZ vector pPD46.21 (kindly provided by Andrew Fire) was
used for transgene constructions. pPD46.21 is identical to pPD1.27
(Fire et al., 1990) except that it lacks the sup-7 gene. It contains an
initiation codon and SV40 T antigen nuclear localization signal just
upstream from the bacterial lacZ gene, and polyadenylation sequences
from the SV40 early region. Enhancer/myoD promoter lacZ constructs
were prepared by liberating enhancer/promoter inserts from the CAT
vector by digestion with SalI and XhoI, and cloning into the SalI site
of pPD46.21. Orientation was determined by restriction digests.
tklacZ was prepared by digesting ptkCAT∆EH with BamHI and BglII
and inserting the tk promoter fragment (from −105 to +51 of the
HSVtk gene) into the BamHI site of pPD46.21. Orientation was
determined by sequencing. To produce 258tklacZ, the BamHI/BglII
tk promoter fragment was cloned into the BamHI site of ∆14KS+.
After orientation was determined by sequencing, the 258tk fusion was
excised by digestion with HindIII and XbaI and cloned into the corresponding sites of pPD46.21. The E-box mutant construct
258(E1-E3)/−2.5lacZ was cloned by excising the mutagenized core
enhancer (see below) from ∆14KS+ with SalI and XbaI, and cloning
the fragment into the SalI and SpeI (derived from the KS+ multiple
cloning site during excision of the myoD promoter fragment) sites of
−2.5lacZ.
MATERIALS AND METHODS
Sequence analysis
The sequence of the human and mouse myoD enhancer was determined on both strands by dideoxy sequencing using the Sequenase kit
(version 2.0; USB). The UWGCG sequencing package was used for
sequence analysis, and transcription factor site searches utilized signal
scan software (Prestridge, 1991). The nucleotide sequence of the
Unless otherwise noted, all molecular biological techniques were
conducted using standard methods (Sambrook et al., 1989). All DNAs
were purified by double banding in cesium chloride equilibrium
gradients.
Cloning of the mouse myoD enhancer
Mouse sequences homologous to the human enhancer were detected
by Southern analysis of PstI-digested mouse genomic DNA. To create
a size-limited library, the hybridizing band was excised from an
agarose gel, ligated into PstI-digested KS+, and electroporated into
NM554 bacteria using the BioRad Gene Pulser. This library was
screened by standard methods using a probe from the human core
enhancer to isolate the corresponding mouse sequences. Linkage to
the mouse myoD gene was verified by hybridization of enhancer-containing cosmid clones (cosmid library was kindly provided by
Yoshimichi Nakatsu) with a mouse myoD cDNA.
Regulation of the myoD gene in mouse embryos
human and mouse myoD core enhancer has been submitted to the
GenBank database.
Mutagenesis
E-box mutations were created by the PCR-based overlap extension
method as previously described (Ho et al., 1989), using Vent polymerase, and ∆14KS+ as the template. By using mutant derivatives as
the template for subsequent rounds of mutagenesis, a construct was
obtained in which all three conserved E-boxes were mutated. All Eboxes were changed from the wild-type sequence CANNTG to
CTNNTA, which destroys the minimal sequence required for binding
of bHLH myogenic factors. Mutations were confirmed by sequencing
both strands of the core enhancer in ∆14KS+.
Transfections and CAT assays
Culturing of 23A2 myoblasts (Konieczny and Emerson, 1984), transfections, cell extract preparation and CAT assays were done as previously described (Goldhamer et al., 1992). A minimum of three independent experiments were conducted for each DNA construct.
Identical molar amounts (0.8 pmoles) of each plasmid were used
(between 2.2 µg and 6 µg, depending on the size of the plasmid),
which was adjusted to 25 µg with pUC 8 carrier plasmid DNA.
Protein concentrations in cell extracts were measured by the modified
Bradford assay (Bio-Rad) using bovine serum albumin as the
standard. 15 µg to 25 µg of protein was used in each CAT assay,
which yielded CAT activities within the linear range of the assay.
Nuclear extract preparation
C2C12 myoblasts, 10TG fibroblasts and JEG-3 choriocarcinoma cells
were purchased from the American Type Culture Collection. HMP8
and FC1010 cells (kindly provided by Mark Lovell and Jerome Freed,
respectively) are primary human myoblasts and foreskin fibroblasts,
respectively. C2C12, 10TG, and JEG-3 cells were grown in DMEM
(Gibco: with high glucose and sodium pyruvate) supplemented with
10% (10TG and JEG-3) or 15% (C2C12) fetal bovine serum (FBS;
Hyclone). HMP8 cells were grown in Ham’s F-10 supplemented with
20% FBS and 0.5% chick embryo extract. FC1010 cells were grown
in RPM-I supplemented with 10% FBS. All media was supplemented
with penicillin (100 units/ml) and streptomycin (100 µg/ml). Cells
were fed fresh medium every 3 days and harvested at about 50 to 80%
confluence. Typically, 20 to 40, 15 cm plates were used for each
nuclear extract preparation. Nuclear extracts were prepared according
to the method of Zaret (personal communication) as follows. After
rinsing cells twice with calcium- and magnesium-free phosphatebuffered saline at room temperature, 5 ml of ice-cold PSDP (0.15 M
NaCl, 20 mM sodium phosphate pH 7.4, 0.35 M sucrose, 0.5 mM
dithiothreitol [DTT], 1 mM phenylmethylsulfonyl fluoride [PMSF], 5
µg/ml leupeptin) was added per plate, and cells were scraped into 50
ml tubes on ice. Cells were pelleted at 2,000 g for 5 minutes at 4°C.
All subsequent procedures were conducted in a cold room on ice. Cell
pellets were washed two times with PSDP and gently resuspended (2
ml for each original 50 ml of cell suspension) in an ice-cold hypotonic
buffer (buffer A; 10 mM KCl, 10 mM Hepes pH 7.9, 1.5 mM MgCl2,
0.5 mM DTT, 1 mM PMSF, and 5 µg/ml leupeptin). Cell suspensions
were combined, swelled on ice for 5 minutes, and centrifuged at 2,000
g for 5 minutes at 4°C. Cell pellets were resuspended in 3 ml of buffer
A containing 0.5% NP-40, incubated on ice for 5 minutes, and cells
lysed with a dounce homogenizer (10-15 strokes, pestle A). The cell
lysate was transferred to a 15 ml disposable tube, 6 ml of buffer B
(60 mM KCl, 15 mM NaCl, 15 mM Tris-HCl pH 7.4, 0.2 mM EDTA,
0.2 mM EGTA, 0.5 mM spermine, 0.15 mM spermidine, 1 mM DTT,
2 mM PMSF, and 5 µg/ml leupeptin) was added, and the cell lysate
was centrifuged at 1,600 g for 5 minutes at 4°C. After gently washing
the nuclear pellet with 2 ml of buffer B and centrifuging as above, the
nuclear pellet was gently resuspended in 2 ml of hypertonic buffer C
(0.42 M NaCl, 20 mM Hepes pH 7.9, 1.5 mM MgCl2, 0.2 mM EDTA,
25% glycerol, 1 mM DTT, 2 mM PMSF, 5 µg/ml leupeptin). The
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nuclear suspension was transferred to two microfuge tubes and
incubated on ice for 30 minutes, gently inverting the tubes every 5
minutes. Samples were microfuged at 3,000 revs/minute at 4°C for 5
minutes, and the supernatants dialyzed (6,000 to 8,000 Mr cutoff) with
1 liter of buffer D (60 mM KCl, 20 mM Hepes pH 7.9, 20% glycerol,
0.2 mM EDTA, 0.5 mM DTT, 1 mM PMSF) with two changes. After
dialysis, extracts were microfuged for 5 minutes at full speed at 4°C,
and small aliquots were quick-frozen in liquid nitrogen. Protein concentrations were determined by the modified Bradford assay (BioRad) using bovine serum albumin as the standard.
DNase I protection assays
The 258 bp core enhancer in ∆14 KS+ was 5′ end-labeled on the
forward or reverse strand with [γ-32P]ATP (6,000 Ci/mmole; NEN)
using unique SalI and XbaI restriction sites flanking the insert.
Labeled fragments were purifed on non-denaturing 5% acrylamide
gels, and after elution, were concentrated by ethanol precipitation. For
each 50 µl reaction, 20 µg of nuclear protein and 5,000 to 10,000
cts/minute (1.4 fmoles) of labeled fragment was incubated in binding
buffer (12 mM Tris pH 8.0, 50 mM KCl, 1 mM DTT, 1 mM MgCl2,
1 mM CaCl2, 5 mM NaCl, 100 µg/ml bovine serum albumin, 5%
glycerol, 2% polyvinyl alcohol, and 0.5 µg poly(dI/dC)) on ice for 1
hour. Reactions were equilibrated to room temperature, 50 µl of
binding buffer was added and, 1 minute later, DNase I (Worthington)
was added to a final concentration of 1 µg/ml (experimental lanes) or
6 or 12 ng/ml (control lanes containing bovine serum albumin in place
of extract). After exactly 2 minutes, stop buffer (50 mM EDTA, 0.2%
SDS, 100 µg/ml yeast tRNA) was added, proteinase K was added to
75 µg/ml and tubes were incubated at 37°C for 30 minutes. Samples
were extracted with an equal volume of phenol and the DNA was precipitated with ethanol for 5 minutes at room temperature. After centrifugation, pellets were washed two times in 70% ethanol, dried in a
Speed-Vac, and resuspended in 8 µl of loading buffer (40%
formamide, 1 mM EDTA). Samples were heated to 80°C and 3 µl of
each sample was resolved on a 6% denaturing polyacrylamide gel.
Dried gels were exposed to X-OMAT AR X-ray film (Kodak) with
intensifier screens for 2 to 4 days. Positions of footprints were determined by comparison with G+A sequencing ladders prepared by
standard Maxam and Gilbert chemical cleavages. At least two independently prepared extracts were tested for each cell type. DNase I
protection assays were conducted a minimum of 3 times with each
extract.
DNase I hypersensitivity blots
All cell types were from the American Type Culture Collection. The
NB41A3 neuroblastoma cell line was grown in F-10 medium supplemented with 15% horse serum and 2.5% FBS. All other cell types
were grown in DMEM with 10% FBS. Cells were grown in 10 cm
dishes and fed every 3 days to approximately 80% confluence. Cells
were harvested by scraping in calcium- and magnesium-free
phosphate-buffered saline, permeabilized in NP-40, and treated with
a range of DNase I concentrations (from 40-120 µg/ml) for 3 minutes
on ice by the method of Rigaud et al. (1991). DNA was isolated by
standard methods. Restriction enzymes and probes used are shown in
Fig. 6. Hybridization and washing was done as described (Church and
Gilbert, 1984).
Whole-mount in situs
Whole-mount in situs utilized a digoxigenin-labeled probe as
described (Conlon and Rossant, 1992). The probe was from
nucleotides 751 to 1785 of the mouse myoD cDNA (Sassoon et al.,
1989; Faerman and Shani, 1993).
Transgenic mice
lacZ fusions were liberated from vector sequences and fragments
purified as previously described (Goldhamer et al., 1992). Transgenic
mice were produced by pronuclear injection of FVB/N 1-cell-stage
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D. J. Goldhamer and others
embryos (Hogan et al., 1986; Shani, 1986). To produce stable transgenic lines, DNA-positive male mice, or male offspring of female
transgenic mice were mated to FVB/N mice. For analysis of injected,
Fo mice, at least four independently derived lacZ-positive embryos
were analyzed for each construct. DNA-positive mice were detected
by Southern blot analysis of tail (stable lines) or placental (Fo mice)
DNA as described (Shani, 1986), using a probe specific to lacZ
sequences.
RESULTS
A distal 258 bp core enhancer controls the
embryonic activation of myoD
We previously identified a muscle-specific enhancer contained
in a 4 kb fragment (fragment 3 in Goldhamer et al., 1992)
located 18 to 22 kb upstream of the human myoD gene. In combination with the myoD promoter, this enhancer drives
expression of a bacterial lacZ gene in all skeletal muscle compartments of mouse embryos, conferring the endogenous myoD
pattern of expression in 11.5 days post-coitum (d.p.c.). transgenic mice (Goldhamer et al., 1992). To analyze further the
temporal and spatial pattern of activation of the transgene, we
created stable transgenic lines that harbor the 4 kb enhancercontaining fragment juxtaposed to the myoD promoter
(contained in 2.5 kb of myoD 5′ flanking sequence) cloned
upstream of the lacZ gene. The spatiotemporal pattern of lacZ
expression throughout embryogenesis coincides with that of
the endogenous mouse myoD gene (Fig. 1; Faerman et al.,
unpublished data). Several interesting features of transgene
expression patterns in somites are described below.
In most somites anterior to the level of the forelimb bud of
all three independently derived transgenic lines, lacZ-positive
cells are scattered throughout the dorsal-ventral myotomal axis
(Fig. 1A-C) and transgene expression appears to be coordinately activated along the dorsal-ventral axis beginning at 1010.5 d.p.c. (Fig. 1B). The most intense staining in these
anterior somites is in the dorsal region of the myotome (Fig.
1A,C; see also Fig. 3A). In somites posterior to the forelimb
bud, however, lacZ-expressing cells initially accumulate in the
ventral portion of the myotome (Fig. 1B), followed by
expression in dorsal myotomal cells, including cells of the
dorsal medial lip of the dermamyotome (Fig. 1A,D). In
contrast to more rostral somites, the most intense staining in
these somites is in the ventral region of the myotome (Fig.
1A,B,D), which is likely due to a greater density of β-galactosidase (β-gal)-positive cells (Fig. 1D). Interestingly, a population of myotomal cells between these ventral and dorsal populations does not express the transgene (Fig. 1A,D) throughout
myotomal stages, a result confirmed by analysis of serial
sections. Whole-mount in situ analysis of endogenous mouse
myoD mRNA yielded a similar pattern of myotomal expression
that is dependent on axial position; message accumulation is
highest dorsally in the myotome of the most anterior somites
and ventrally in the myotome of more posterior somites (Fig.
1E). These data demonstrate the sufficiency of the identified
regulatory elements in recapitulating the normal spatial pattern
of myoD expression in somites.
To further localize enhancer activity, overlapping restriction
fragments and nested deletions were tested in transient transfection assays for their ability to enhance transcription from
the myoD promoter in 23A2 myoblasts. The myoD promoter
alone has a very low basal level of CAT reporter gene activity,
only about 5-fold above a promoterless CAT construct.
Fragment 3 typically increases this basal level of transcription
10- to 20-fold in 23A2 myoblasts (Goldhamer et al., 1992). An
internal 1.7 kb ApaI/PstI fragment was identified that yields
approximately 50% of the CAT activity of fragment 3 (Fig.
2A). No other subfragment residing entirely outside of this 1.7
kb fragment exhibits enhancer activity, although all of the
activity of fragment 3 could be reconstituted by the additive
activities of two fragments that overlap the 1.7 kb fragment;
the 2.1 kb KpnI fragment (see Fig. 2B), which includes the 5′
half of the 1.7 kb fragment, exhibits approximately 70% of the
activity of fragment 3, and a 1.4 kb KpnI/EcoRI, which
includes the 3′ half of the 1.7 kb fragment constitutes the
remaining activity (data not shown). Importantly, the 1.7 kb
fragment, and subfragments therein, yield the appropriate
pattern of muscle-specific expression in transgenic mice (see
below). Therefore, sequences outside of the 1.7 kb fragment
were excluded from further analysis.
Transfection analysis of bi-directional nested deletions of
the 1.7 kb fragment identified two non-contiguous DNA
elements with modest enhancer activity (Fig. 2A). Enhancer 1,
defined by the 3′ to 5′ deletion 14, was localized to a 258 bp
sequence at the 5′ end of the 1.7 kb fragment (Fig. 2A, bottom).
Further deletion to nucleotide +95 abolishes enhancer activity
(Fig. 2A; deletion 15). Approximately 1 kb downstream, a
second enhancer was identified whose 5′ end falls between the
endpoints of the 5′ to 3′ deletions, 7 and 8. The 3′ limit of this
enhancer lies between the endpoint of the 3′ to 5′ deletion 1,
and the 3′ end of the 1.7 kb fragment. Precise definition of the
enhancers’ boundaries is difficult because of their inherently
low activity in transfection assays. No activity was detected in
sequences between enhancers 1 and 2 (data not shown). As
with fragment 3 (Goldhamer et al., 1992), the 1.7 kb fragment,
as well as subfragments containing enhancer 1 or 2, are active
when transfected into both muscle and non-muscle cells in
culture.
Deletion constructs containing either enhancer 1 or enhancer
2 were tested transgenically for their ability to direct
expression of a lacZ reporter gene in the typical myoD
expression pattern (Fig. 2B, 3). Transgenes driven by enhancer
2 and the myoD promoter are only ectopically expressed with
no consistent pattern, which is typical of results observed with
promoter sequences alone (Goldhamer et al., 1992). In
contrast, both subfragments containing enhancer 1 (Fig. 2B)
are expressed specifically in skeletal muscle in transgenic
mice; all five Fo transgenic mice and a stable line containing
the minimal 258 bp enhancer 1 and the myoD promoter exhibit
a pattern of expression indistinguishable from the larger 1.7 kb
fragment or the 4 kb fragment 3 (Fig. 3A). Because enhancer
1 directs the myoD pattern of expression in transgenic mice,
this 258 bp sequence will hereafter be referred to as the myoD
core enhancer.
The myoD core enhancer was tested in combination with the
heterologous herpes simplex virus thymidine kinase (tk)
promoter to assess the relative contributions of the enhancer
and myoD promoter to the regulation of myoD expression. As
shown previously, the tk promoter alone exhibits only ectopic
expression due to integration site position effects (Allen et al.,
1988; data not shown). In contrast, all seven DNA-positive
Regulation of the myoD gene in mouse embryos
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Fig. 1. Spatial domains of myoD expression in somites. (A-D) β-gal expression in stable F3/−2.5 lacZ transgenic embryos. (A) Whole-mount
11.5 d.p.c. embryo showing the overall pattern of β-gal expression. Transgene expression is predominantly in the dorsal region of the myotome
of the most anterior somites (white arrows) and in the ventral region of myotomes (black arrows) posterior to the forelimb bud (fb). Staining is
also seen in forelimb and hindlimb buds (hb) and in branchial arches (ba). Ectopic staining in the neural tube is likely due to position effects.
The nasal epithelium (asterisk) usually shows transgene expression in independent lines, for unknown reasons. (B) 10.5 d.p.c. whole-mount
embryo from same transgenic line shown in A. Transgene expression is most prominent in the ventral-most portion of thoracic somites (black
arrows). Note staining along the entire dorsal-ventral axis of the most anterior somites. Weak staining in the dorsal-most region of the
myotomes of thoracic somites is obscured by ectopic neural tube staining. (C) Transverse section anterior to the forelimb bud of an 11.5 d.p.c.
embryo from an independent transgenic line. Nuclear localized β-gal expression is observed throughout the dorsal-ventral axis of the myotome
with the most intense staining dorsally (white arrow). asterisk, hypoglossal premuscle mass, corresponding to the more dorsal strip of stained
cells ventral to anterior somites in A (see Faerman and Shani, 1993). Ectopic expression is restricted to dorsal root ganglia (drg) in this line. nt,
neural tube. (D) Transverse section posterior to the forelimb bud from the same embryo shown in C. β-gal expression in the myotome of this
thoracic somite is most prominent ventrally (black arrow). Note the population of β-gal negative cells (bracket) under the dorsal-most region of
the myotome (white arrow). (E)Whole-mount in situ localization of endogenous mouse myoD mRNA in a 10.5 d.p.c. embryo. The spatial
pattern of myoD message accumulation closely matches the pattern of transgene expression. black arrows, localization of myoD mRNA in the
ventral region of myotomes of thoracic somites. white arrows, myoD mRNA is most abundant in the dorsal region of anterior somites. Low
level expression in the forelimb bud and branchial arches is not apparent in this preparation. Staining of the otic vesicle (ov) is non-specific
background.
embryos injected with a construct containing the core enhancer
in combination with the tk promoter show clear musclespecific expression at 11.5 d.p.c., exhibiting a rostrocaudal
gradient of expression in myotomes, as well as expression in
branchial arches, limb buds, and other myogenic centers (Fig.
3B). As with previous constructs, the most anterior somites and
thoracic somites could be distinguished by their prominent
dorsal and ventral staining, respectively. Interestingly,
however, most of these transgenic mice exhibit lacZ-positive
cells scattered throughout the dorsal-ventral myotomal axis of
all somites, regardless of their axial position (Fig. 3B). This
indicates a contribution either of sequences in fragment 3
outside of the core enhancer, or of myoD promoter sequences,
in restricting the spatial expression of myoD within the somite
myotome.
The myoD core enhancer is highly conserved
between humans and mice
As a means of identifying critical regulatory motifs within the
enhancer, we compared myoD enhancer sequences between
humans and mice, reasoning that important regulatory
sequences would be most highly conserved. For this analysis,
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D. J. Goldhamer and others
Fig. 2. (A) Localization of myoD
enhancer activities by transient
transfection assays in 23A2
myoblasts. The 1.7 kb ApaI/PstI
fragment within fragment 3 (see
B) was the parental molecule
used to create nested deletions.
All deletion constructs were
assayed for enhancer activity in
combination with the myoD
promoter. Numbers are relative to
the CAT activity of the promoter
alone, which was arbitrarily set to
a value of 1. Values (± s.e.m.) are
the average of at least three
independent experiments. Arrows
denote the maximum limits of the
two enhancers. (B) Summary of
transgenic data, localizing the
muscle-specific regulatory region
to a 258 bp core enhancer. All
fragments shown were assayed in
combination with the myoD
promoter. The approximate
positions of enhancer activities
defined by transient transfection
assays are shown (boxes).
the mouse myoD enhancer was cloned from a size-selected
plasmid library using the human core enhancer as a probe.
Restriction mapping and Southern blot analyses of enhancercontaining cosmid clones demonstrated linkage with the mouse
myoD gene, and revealed that the mouse myoD enhancer, like
the human enhancer, is located approximately 20 kb 5′ of the
start of transcription (data not shown). The human and mouse
core enhancer show extensive sequence similarity (89%
identity), particularly within the first 160 bp, in which the
enhancer in the two species is 94% identical (Fig. 4). Sequence
similarity drops dramatically outside of the core enhancer,
although small regions of homology exist throughout the 1.7
kb ApaI/PstI fragment as well as 5′ of the core enhancer
(unpublished observations).
The human core enhancer sequence contains four E-boxes,
three of which (E-1, E-2 and E-3) are conserved in sequence
and position in mice (Fig. 4). Both central and flanking
nucleotides, which strongly influence the affinity of bHLH
protein binding (Blackwell and Weintraub, 1990; Sun and
Baltimore, 1991; Wright et al., 1991), are conserved between
species in E-boxes 2 and 3. E-boxes 2 and 3 represent potential
high affinity binding sites for MyoD/E12-E47 and MyoD
homodimers, respectively (Blackwell and Weintraub, 1990;
Sun and Baltimore, 1991). With the exception of E-boxes, the
myoD core enhancer is distinguished by the lack of binding
sites for factors known to regulate the expression of other
muscle-specific genes (Fig. 4). Motifs that are absent include
MEF-2 (Gossett et al., 1989; Cserjesi and Olson, 1991), CArG
(Minty and Kedes, 1986), MHOX (Cserjesi et al., 1992), and
M-CAT (Mar and Ordahl, 1990) sites. The A-T rich sequences
Regulation of the myoD gene in mouse embryos
from nucleotides 16 to 25, and from nucleotides 31 to 41 (Fig.
4) resemble consensus CArG and MEF-2 motifs, but differ at
nucleotides required for serum response factor and MEF-2
binding (Treisman, 1986; Pollock and Treisman, 1990, 1991;
Cserjesi and Olson, 1991). A search of the transcription factor
database revealed consensus sequence binding sites for several
widely expressed transcription factors, including the Ets
protein, PEA3 (Xin et al., 1992); nts 28-33, reverse strand),
AP-1 (nts 143-149), NF-1 (nts 176 to 187), and H4TF-1
(Dailey et al., 1988; nts 165-173), a regulator of the histone H4
gene (Fig. 4).
Mutation of the enhancer E-boxes does not affect
enhancer activation in the embryo
To test the functional significance of the conserved E-boxes,
an enhancer construct in which all three conserved E-boxes (E1 through E-3; Fig. 4) were mutated, was tested in transgenic
mice. In this construct, the canonical E-box motif CANNTG
was changed to CTNNTA, thereby destroying the minimal
sequence required for bHLH factor binding. This construct
(258(E1-E3)/−2.5lacZ) contains the mutant core enhancer cloned
upstream of the myoD promoter in the lacZ vector used above.
Fo mouse embryos were analyzed at 11.5 d.p.c., which is about
1 to 1.5 days after the myotomal activation of both the endogenous myoD gene (Sassoon et al., 1989; Faerman and Shani,
1993), and myoD enhancer lacZ fusions. Defects in the activation function of the enhancer would result in a delay or loss
of lacZ expression, as observed with E-box mutations in the
myogenin promoter (Cheng et al., 1993; Yee and Rigby, 1993).
Analysis of four lacZ-positive embryos, however, revealed no
apparent difference between the pattern of expression of the
mutant construct and the wild-type constructs (Fig. 3C). Also,
there appears to be no delay in activation, since the caudal
extent of expression along the rostrocaudal expression gradient
is similar to that of wild-type constructs at 11.5 d.p.c. (compare
to Figs 1A, 3A). In addition, expression of the mutant
transgene was detected in the hindlimb bud at 11.5 d.p.c., a
time coincident with the initial detection of wild-type enhancer
constructs. An enhancer construct entirely lacking E-boxes (the
three conserved E-boxes and the fourth, non-conserved E-box;
see Fig. 4), also is expressed normally at 11.5 d.p.c (unpublished observations). We conclude that E-boxes in the myoD
core enhancer are not required for myoD gene activation.
Nuclear trans factors interact with multiple
sequence elements in the core enhancer
DNase I protection assays were used to identify enhancer
sequences that interact in vitro with nuclear factors from
human and mouse cells. Mouse cell types analyzed were
C3H10TG (10TG) fibroblasts and C2C12 myoblasts. Human
cells used were primary fibroblasts (FC1010), primary
myoblasts (HMP8) and the choriocarcinoma cell line, JEG-3.
A representative experiment is shown in Fig. 5. Extracts from
HMP8 myoblasts, C2C12 myoblasts and 10TG fibroblasts show
identical DNase I footprinting profiles (protected regions and
hypersensitive sites). Five protected regions and many hypersensitive sites were detected, nearly spanning the core
enhancer (Figs 4, 5). The hypersensitive sites not associated
with protected regions likely reflect lower affinity
DNA/protein interactions, or binding of lower abundance
proteins. Protected regions 4 and 5, which encompass
643
consensus binding sites for AP-1 (Lee et al., 1987) and H4TF1 (Dailey et al., 1988), respectively, are produced with nuclear
extracts from all cell types tested, including 23A2 azamyoblasts, primary chicken liver cells and BNL liver cells
(Fig. 5; data not shown). These sites represent the highest
affinity sites and/or are bound by the most abundant factors
because they typically exhibit complete protection, even under
more stringent conditions (10 µg protein, 4 µg poly(dI/dC);
non-specific competitor; data not shown). Protection of Eboxes was not observed, however E-box binding is difficult to
detect using standard DNase I protection assays (Buskin and
Hauschka, 1989).
Both primary human fibroblasts (FC1010) and JEG-3 cell
extracts yield cell-specific qualitative and quantitative differences in their footprinting profiles. Extracts from JEG-3 cells,
a cell line in which the transfected enhancer is inactive
(Goldhamer et al., 1992), does not show detectable DNase I
protection over region 2, and only partial protection over
region 1. Note also the absence of a hypersensitive site between
regions 1 and 2, and just 5′ of region 4 in JEG-3 cell extracts
(Fig. 5, forward strand). Also JEG-3 extracts yield partial protection of sequences between regions 2 and 3, and produced a
hypersensitive site between regions 3 and 4 (Fig. 5, forward
strand). Finally, protected region 3 extends slightly further 5′
with JEG-3 extracts than with the other extracts. FC1010
extracts exhibit partial or complete protection over all five
regions, although DNase I hypersensitivity was reduced or
absent at most sites denoted in Figs 4 and 5. In addition, protection over region 4 extends slightly further 3′ with FC1010
extracts (Fig. 5), suggesting a qualitative difference in
DNA/protein interactions over this site.
The endogenous myoD enhancer exhibits musclespecific DNase I hypersensitivity
The identical DNase I protection profiles produced using
nuclear extracts from non-myogenic 10TG cells (in which the
endogenous gene is inactive), C2C12 myoblasts and HMP8
myoblasts suggest that the constellation of enhancer binding
proteins are highly similar in these cell types. In addition, the
myoD enhancer is active in 10TG cells when introduced by
transfection (Goldhamer et al., 1992). These data raise the possibility that repression of the endogenous myoD gene in 10TG
cells is mediated by epigenetic mechanisms, such as packaging
into inactive chromatin, that could restrict accessibility of
critical cis-acting enhancer sequences to positive trans factors.
We used DNase I hypersensitivity as an indicator of chromatin
structure to assess whether chromatin structural differences
exist between myogenic and non-myogenic cells. For this
analysis, the endogenous mouse myoD enhancer was assayed
in C2C12 and 23A2 myogenic cells in addition to several nonmyogenic cell lines, while the endogenous human myoD
enhancer was assayed in primary human fibroblasts (FC1010)
and myoblasts (HMP8). Permeabilized cells were treated with
increasing concentrations of DNase I, and the presence and
position of hypersensitive sites determined by Southern
blotting using indirect end-labeled probes. Three distinct
hypersensitive sites are present in chromatin derived from
C2C12 and 23A2 myoblasts, two of which map to the core
enhancer, with the third mapping just 5′ of the enhancer (Fig.
6). In contrast, no hypersensitive sites were detected in
chromatin from non-myogenic mouse cells, including 10TG
644
D. J. Goldhamer and others
cells, BNL liver cells, or NB41A3 neuroblastoma cells (Fig.
6). Similarly, chromatin from primary human myoblasts
exhibits DNase I hypersensitivity within the core enhancer,
whereas chromatin from primary human fibroblasts is 5- to 10fold more resistant to DNase I digestion, although a very weak
signal was detected (data not shown). These data provide
evidence that the chromatin structure of the core enhancer is
altered in myogenic cells, perhaps reflecting greater accessibility to trans factors.
DISCUSSION
A distal core enhancer governs myoD gene
activation in the embryo
We have identified and characterized regulatory sequences
responsible for myoD expression to begin to define upstream
signaling pathways and transcriptional events that govern
myoD activation in the embryo. Cis-acting sequences that
regulate the embryonic expression of the human myoD gene
were localized to a 258 bp core enhancer, which is located
approximately 20 kb upstream from the start of myoD transcription (Goldhamer et al., 1992). A lacZ reporter gene, under
the control of the myoD enhancer and promoter, is expressed
in a muscle-specific and spatiotemporal pattern that coincides
with the endogenous mouse myoD expression domain, as
revealed by in situ hybridization (present study; Buckingham,
1992; Faerman and Shani, 1993; Faerman et al., in preparation). Transfection assays revealed a second, weak enhancer
approximately 1 kb 3′ of the core enhancer, which may contribute to quantitative levels of myoD expression in vivo. This
second region, however, in combination with the myoD
promoter, was neither necessary nor sufficient for musclespecific expression of the lacZ transgene. We conclude that the
258 bp core enhancer mediates myoD activation in all
embryonic skeletal muscle compartments, including somitic
myotomes, limb buds and branchial arches.
The relative contributions of the myoD core enhancer and
promoter in directing myoD expression was tested by replacing
the myoD promoter with the heterologous tk promoter. We
showed that the myoD promoter is dispensable for activation
of the myoD gene in the embryo, as this heterologous construct
also yields muscle-specific expression of a lacZ reporter gene
that closely resembles the endogenous pattern of myoD transcript accumulation. This experiment also establishes that
myoD mRNA accumulation is dictated primarily by transcriptional control mechanisms rather than by mRNA turnover,
because no transcribed human sequences were present in the
heterologous promoter construct. Consistent with these functional data, the core enhancer is highly conserved in sequence
Fig. 3. The 258 bp core enhancer directs the appropriate, myoD
pattern of expression in transgenic mouse embryos. (A) β-gal
expression of 11.5 d.p.c. whole-mount mouse embryo derived from a
stable line harboring the construct, 258/−2.5lacZ. β-gal expression is
detected in all myogenic compartments, including somites, limb
buds, and branchial arches. The spatial pattern of lacZ-expressing
cells is similar to that of larger enhancer constructs (see Fig. 1).
(B) β-gal expression of 11.5 d.p.c. Fo whole-mount mouse embryo
injected with the construct, 258/tklacZ. Expression with this
heterologous promoter construct is observed in all myogenic
compartments. Expression is most prominent in the dorsal portion of
the most anterior somites and in the ventral portion of thoracic
somites, similar to constructs with the myoD promoter. Somites
posterior to the forelimb bud, however, show continuous lacZ
expression along the dorsal-ventral myotomal axis, normally
observed only in the most anterior somites. Ectopic, position effect
staining in the brain and dorsal root ganglia is observed in this
embryo. (C) β-gal expression of 11.5 d.p.c. Fo whole-mount mouse
embryo injected with the E-box mutant construct, 258(E1-E3)/−
2.5lacZ. The pattern of β-gal expression is the same as that of wildtype constructs. Ectopic staining in the neural tube is observed in this
embryo. (A-C) Arrows; β-gal expression in branchial arches. hb;
hindlimb bud.
Regulation of the myoD gene in mouse embryos
645
Fig. 4. Sequence alignment of the human myoD
core enhancer and the corresponding sequence of
the mouse. Sequence similarity is 89% overall,
and 94% in the first 160 bp. Data from DNase I
protection assays (Fig. 5) is also summarized.
The positions of DNase I protection (bars below
sequence) and hypersensitive sites (asterisks
above sequence; the JEG-3-specific
hypersensitive site is denoted by a ‘+’) are
shown. Not all of the hypersensitive sites shown
are produced with FC1010 and JEG-3 nuclear
extracts (see Fig. 5). Protected region 2 is not
produced with JEG-3 nuclear extracts.
Consensus sequence binding sites are shown for
known transcription factors that reside within
regions of protection, or that have been shown or
postulated to function in muscle gene regulation.
(89% overall; 94% in the first 160 nucleotides) and position
between humans and mice, whereas 5′ flanking sequences are
only 66% similar within 250 bp of the start of transcription,
with no extended regions of sequence similarity present
upstream of the TATA box (unpublished observations). When
the tk promoter was used, however, the spatial patterning of
transgene expression was partially disrupted; the transgene was
expressed along the entire dorsal-ventral myotomal axis in all
somites, regardless of axial position (see below). Thus, the
myoD promoter may function to refine spatial domains of
myoD expression within the somite myotome. Also, it is
possible that the promoter regulates aspects of postnatal myoD
expression; a mouse genomic clone consisting of the myoD
promoter and a regulatory element at −5 kb (Tapscott et al.,
1992) that shares no obvious sequence similarity with the core
enhancer, appropriately confers transgene expression preferentially in fast glycolytic fibers of adult muscle (Hughes et al.,
1993). Our results, however, clearly establish that neither the
promoter nor the more proximal control element at −5 kb is
required for muscle-specific activation of myoD in embryos.
myoD is subject to complex transcriptional
regulation
The extensive sequence similarity between the human and
mouse myoD enhancer and the complex patterns of
DNA/protein interactions observed in vitro indicate that myoD
expression in the embryo is subject to complex regulation.
DNase I protection assays identified five protected regions and
eight regions with one or more hypersensitive sites (Figs 4, 5).
Several hypersensitive sites are not associated with detectable
protection, probably reflecting interactions with proteins of
lower abundance or affinity. Only protected regions 4 and 5
encompass consensus binding sites for known factors; region
Fig. 5. DNase I protection assay of the human myoD core enhancer
using nuclear extracts from myogenic (C2C12, HMP8) and nonmyogenic (10TG, FC1010, JEG-3) cell types. Five protected regions
(boxes) and multiple hypersensitive sites (asterisks; the JEG-3specific hypersensitive site is denoted by a ‘+’) were detected. JEG-3
extract does not protect region 2 and yields partial protection of
region 1 (most apparent on forward strand). FC1010 and JEG-3
extracts exhibit substantially different patterns of DNase I
hypersensitive sites compared to the other cell types. 10TG nuclear
extract yields the same footprinting profile as C2C12 and HMP8
myoblast nuclear extracts. The forward strand corresponds to the
sequence shown in Fig. 4. Protected region 1 on the reverse strand is
not very apparent due to band compression near the top of the gel.
BSA, control lanes in which bovine serum albumin replaced nuclear
extract. G+A, purine-specific sequence ladder.
646
D. J. Goldhamer and others
Fig. 6. Southern blot of DNase I treated chromatin, revealing
muscle-specific DNase I hypersensitive sites in the endogenous
myoD core enhancer. Chromatin from mouse myogenic cells (C2C12
and 23A2) exhibits three DNase I hypersensitive sites; two map to
the core enhancer (arrows) and one maps just 5′ of the core enhancer
(arrowhead). Non-myogenic mouse 10TG cells, NB41A3
neuroblastoma cells, and BNL liver cells do not exhibit DNase I
hypersensitivity. The degree of DNAse I digestion of total DNA is
comparable in all cell types, as determined by ethidium bromide
staining of agarose gels. The relative position of the enhancer, and
the restriction enzymes and probe used are shown diagrammatically.
4 includes a consensus AP-1 site, representing a potential
target for fos/jun complexes (Curran and Franza, 1988),
whereas region 5 contains a consensus sequence important in
histone H4 gene expression (Dailey et al., 1988). While we do
not know the molecular species bound to these sites, linkersubstitution mutants encompassing these regions adversely
affect enhancer activity in transfection assays (Lukitsch and
Goldhamer, unpublished observations).
With the exception of 10TG cells (see below), extracts from
non-myogenic cells yield footprinting profiles substantially
different from those of myogenic cells. These differences
observed with human primary fibroblasts (FC1010) and human
choriocarcinoma cells (JEG-3) do not simply reflect human and
mouse species differences because extracts from human
primary myoblasts (HMP8) exhibit a footprinting profile
similar to mouse C2C12 (and 23A2) myoblasts. Numerous
qualitative and quantitative differences in hypersensitive sites
exist with FC1010 and JEG-3 extracts, as detailed in the
results. In addition, extracts from primary human fibroblasts
yield a unique pattern of protection and hypersensitivity
encompassing and adjacent to the consensus AP-1 site (Figs 4,
5). Thus, the protein species bound to this site may be unique
to FC1010 cells. Recently, a hematopoietic-specific factor was
shown to regulate globin gene expression by binding to a
‘ubiquitous’ AP-1 site within the β-globin locus control region
(Andrews et al., 1993). The most dramatic difference between
myogenic and non-myogenic binding activities, however, was
the lack of protection over region 2 and only partial protection
over region 1 with nuclear extracts from ectodermally derived
JEG-3 cells (Figs 4, 5). As the transfected enhancer is inactive
in JEG-3 cells (Goldhamer et al., 1992), region 2 may interact
with tissue-restricted positive trans factors required for
enhancer activity; whether the protein(s) is restricted to mesodermal cells will require further analysis.
Footprinting experiments and previous transfection data
suggest that cis-acting epigenetic mechanisms repress myoD
expression in 10TG cells. Inspection of footprinting gels reveals
no differences in protection or hypersensitivity profiles
produced from 10TG extracts and C2C12, HMP8 and 23A2
myoblast extracts (Fig. 5; data not shown). Assuming that less
stable muscle-specific interactions were not missed in this
assay, these data suggest that the complement of regulatory
factors that bind the myoD enhancer are similar, if not
identical, in 10TG cells and myogenic cells. In addition, the
transfected enhancer is active in 10TG cells (Goldhamer et al.,
1992), indicating that repression of the endogenous myoD gene
is not a consequence of inactivation of these trans factors by
post-translational modification or heterodimerization with
negative regulators. We found, however, that the chromatin
within and immediately surrounding the myoD core enhancer
exhibits DNase I hypersensitivity in myogenic cells, but not in
10TG or other non-myogenic cells. We propose, therefore, that
the myoD core enhancer is packaged in inactive chromatin in
10TG cells, rendering it inaccessible to positive trans factors
required for gene activation. In theory, regulated, musclespecific changes in accessibility of trans factors to myoD
control elements, mediated by chromatin structural changes or
other epigenetic modifications such as DNA methylation,
could be a critical prerequisite for myoD activation, with such
changes being stably passed on to progeny cells (see Groudine
and Weintraub, 1982; Razin and Cedar, 1991). In this regard,
demethylation of specific CpGs in the mouse myoD enhancer
correlates with myoD expression both in cell culture and in
mouse embryos (Brunk et al., unpublished data). Analysis of
the temporal relationship between DNA demethylation,
changes in chromatin structure, and the expression of myoD
will address whether these epigenetic modifications are a cause
or an effect of myoD gene activation.
E-boxes in the core enhancer are not required for
myoD activation
The presence of E-boxes in the myoD core enhancer raised the
possibility that myoD activation is regulated by direct transactivation by other bHLH myogenic factors. Auto- and cross-regulatory interactions between the myogenic factors have been
well-documented in cell culture systems (reviewed by
Emerson, 1990; Olson, 1990). In addition, myogenin promoter
function in embryos is E-box dependent, suggesting that
myogenic factors regulate myogenin expression in vivo (Cheng
et al., 1993; Yee and Rigby, 1993). We found, however, that
enhancer constructs lacking the three conserved E-boxes
(present study) or lacking all four E-boxes (see Fig. 4; unpublished observations), exhibit the wild-type pattern of
expression at 11.5 d.p.c. In addition, both the wild-type and
mutant myoD enhancers are activated in somites to the same
caudal extent, indicating no delay in transgene activation.
Although, we cannot formally rule out the possibility that Eboxes in promoter sequences functionally substitute for the
enhancer E-boxes in this construct, this is unlikely because the
myoD promoter shows no muscle specificity in transgenic mice
when assayed alone (Goldhamer et al., 1992) or in combination with enhancer 2 (Fig. 2B), and is dispensable for musclespecific transgene expression. Thus, unlike many musclespecific genes (see below), activation of myoD is not likely to
be mediated by E-boxes. Importantly, the present experiments
Regulation of the myoD gene in mouse embryos
do not address the possible function of E-boxes in the maintenance of myoD expression, once activated; because β-gal is a
relatively stable protein (see Paterson et al., 1991), E-box
mutant constructs will need to be investigated at later stages of
development further removed from the initial activation of
myoD.
myoD expression is controlled by regulatory
mechanisms distinct from other characterized
muscle-specific genes
myoD exhibits a distinct temporal and spatial pattern of
expression (reviewed by Buckingham, 1992; Faerman and
Shani, 1993; present results), indicating that regulatory mechanisms controlling myoD expression are unique. In the most
anterior somites of the mouse, for example, the myogenic
factors are activated sequentially over a 2.5 day period from 8
d.p.c. to 10.5 d.p.c. in the order; myf5, myogenin, MRF4 and
myoD (Buckingham, 1992). Also, while myf5 (Tajbakhsh and
Buckingham, 1994) and myogenin (Cheng et al., 1992; Yee
and Rigby, 1993) follow a strict rostral to caudal sequence of
activation in somites, myoD transcripts and myoD transgene
expression are first detected in thoracic somites at the level of
the forelimb bud, followed approximately a half day later by
expression in more anterior somites (Faerman et al., in preparation). In addition, spatial domains of myoD expression within
somites exhibit a distinctive axial position-dependent pattern
of expression. In somites anterior to the forelimb bud, myoD
transgene expression is most prominent in the dorsal region of
the myotome, whereas in somites posterior to the forelimb bud,
transgene expression is most prominent ventrally. This axial
position-dependent patterning of expression is also observed
for endogenous myoD transcripts. Finally, lacZ-positive cells
are found scattered throughout the dorsoventral myotomal axis
of the most anterior somites, whereas in somites posterior to
the forelimb bud, dorsal and ventral populations of lacZpositive cells are separated by a spatially restricted population
of myoD-negative myotomal cells. In contrast, myogenin and
myf5 regulatory elements drive expression of lacZ along the
entire dorsal-ventral myotomal axis regardless of axial position
(Cheng et al., 1992; Yee and Rigby, 1993; Tajbakhsh and
Buckingham, 1994). The embryological signals that dictate
these unique expression patterns remain to be defined.
Clear differences also are emerging in the organization and
regulation of cis sequences that control expression of the
myogenic genes, consistent with their distinct patterns of
expression. While myoD is controlled by a distal enhancer 20
kb upstream of the gene, which is highly conserved between
humans and mice, myogenin is regulated by highly conserved
proximal promoter elements (compare sequences in Salminen
et al., 1991 and Edmondson et al., 1992) within 200 bp of the
start of transcription (Salminen et al., 1991; Cheng et al., 1992;
Yee and Rigby, 1993). myf5 and myf6 are also regulated, at
least in part, by 5′ flanking sequences (within 6 kb) as transgenes containing these sequences recapitulate some aspects of
the expression pattern of the corresponding endogenous genes
(Patapoutian et al., 1993). In addition, the myoD core enhancer
exhibits no extended regions of sequence identity with the
myogenin promoter or other muscle-specific enhancers or
promoters (specific sequences controlling the embryonic
expression of myf5 and MRF4 genes have not yet been
defined). Furthermore, among the specific DNA motifs known
647
to regulate other muscle genes, including E-boxes, MEF-2
sites, CArG boxes, and MCAT sites, only E-boxes are present
in the core enhancer. The E-boxes, however, are not required
for myoD activation. In striking contrast, the myogenin
promoter contains an E-box and a MEF-2 site, both of which
are critical for normal myogenin expression in embryos (Cheng
et al., 1993; Yee and Rigby, 1993). Together with possible epigenetic mechanisms regulating the myoD enhancer by control
of chromatin structure and methylation status noted above,
these data indicate that myoD is regulated by a control system
distinct from the other characterized muscle-specific genes.
Analysis of the regulation and expression of the myogenic
factors has revealed heterogeneity among myotomal cell populations. Mutational studies of myogenin promoter function
revealed MEF-2-dependent and -independent populations of
cells (Cheng et al., 1993; Yee and Rigby, 1993). In the present
study, we document the existence of spatially restricted populations of myoD-positive and myoD-negative myotomal cells.
In addition, previous immunocytochemical studies identified
myosin-positive cells that express either MyoD or Myogenin,
neither protein, or both proteins (Cusella-De Angelis et al.,
1992). Finally, Myf5 protein appears to be expressed by more
cells than MyoD in cultures derived from somites (Smith et al.,
1993), although individual cells have not been assayed for both
markers. Whether these differential patterns of myogenic gene
expression reflect different fates or developmental potentials
among these cell populations is presently unclear. Given the
formative but functionally redundant roles of myf5 and myoD
in skeletal muscle formation, it will be important to determine
the degree of concordance of myoD and myf5 expression in
individual myogenic cells. This will help distinguish whether
functional redundancy arises because myf5 and myoD have
similar biochemical properties within the same cell or whether
distinct myf5-positive and myoD-positive myogenic populations can regulate their size and compensate for the loss of the
other (see Emerson, 1993; Weintraub, 1993). Using myoD
enhancer-lacZ transgenes to track myoD expression domains
in myf5 knockout mice will also address cellular compensatory
mechanisms.
Activation of myogenic gene expression is initiated very
early in presumptive myotomal cells (Ott et al., 1991; Pownall
and Emerson, 1992), whereas presumably committed
myogenic cells migrate from the lateral somite (Ordahl and Le
Douarin, 1992) into the developing limb before expression of
the myogenic genes is detected (Sassoon et al., 1989; Cheng
et al., 1992; Yee and Rigby, 1993; Tajbakhsh and Buckingham, 1994). This raises the questions of whether these distinct
skeletal muscle lineages are established and maintained by
similar mechanisms, and whether myogenic cell fate in presumptive limb muscle is governed by additional unknown
factors. As the myoD core enhancer functions as a molecular
target to integrate ‘upstream’ embryonic signals and activate
myoD in all skeletal muscle lineages, functional dissection of
the myoD enhancer will serve as a directed molecular approach
to investigate upstream regulatory processes, including the
mechanisms of skeletal muscle lineage determination in the
embryo.
We thank Yoshimichi Nakatsu for providing the mouse cosmid
library, and Marisa Bartolomei and Kristen Lukitsch for critical
comments on the manuscript. This work was supported by grants from
648
D. J. Goldhamer and others
the American Cancer Society (IRG-135N) and the National Institutes
of Health (NIH; AR-42644) to D. J. G., by a CORE grant (CA-06927)
from the NIH and an appropriation from the Commonwealth of Pennsylvania, by research grants from the Muscular Dystrophy Association and the NIH (HD-07796) to C. P. E., and by a grant from the
United States-Israel Binational Agricultural Research and Development Fund (BARD) to C. P. E. and M. S.
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(Accepted 30 November 1994)