From ETH Zürich, Institut für Molekularbiologie und Biophysik, ETH-Hönggerberg, CH-8093 Zürich, Switzerland
Received for publication, February 17, 2003, and in revised form, March 6, 2003
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Myocyte enhancer factor 2 (MEF2) proteins play a
pivotal role in the differentiation of cardiac and skeletal muscle
cells. MEF2 factors are regulated by histone deacetylase enzymes such as histone deacetylase 5 (HDAC5). HDAC5 in turn is responsive to
Ca2+ signaling mediated by the intracellular calcium
sensor calmodulin. Here a combination of proteolytic fragmentation,
matrix-assisted laser desorption ionization mass spectrometry, Edman
degradation, circular dichroism, gel filtration, and surface plasmon
resonance studies is utilized to define and characterize a stable core
domain of HDAC5 and to examine its interactions with MEF2a and
calmodulin. Results from real time binding experiments provide evidence
for direct interaction of Ca2+/calmodulin with HDAC5
inhibiting MEF2a association with this enzyme.
In eukaryotes, transcription occurs in the nucleus where the DNA
template is packaged in chromatin. DNA is wrapped tightly around
histone proteins in nucleosomes that are arranged in a chromatin higher
order structure (1-3). The organization of DNA in chromatin is thought
to act as a barrier to transcription causing gene repression. Two
alterations of chromatin have been implicated in transcription
regulation: remodeling by ATP-dependent factors such as
SWI/SNF, RSC, NRD, or NURF (4); and post-translational modification of
the histone N-terminal tails including phosphorylation, methylation,
ubiquitination, and acetylation (5). Among these, the acetylation
modification has been recognized as a major contributor to
transcription regulation (6) and is maintained by the dynamic interplay
of histone acetylase and deacetylase
(HDAC)1 enzyme antagonists.
Acetylation of histone tails by histone acetylases is thought to create
a chromatin structure accessible to the transcription machinery (1, 7),
and conversely, hypoacetylated chromatin, the product of HDACs, is
often associated with transcriptionally silent DNA (8).
Histone deacetylases have attracted considerable attention recently,
due to findings that compounds blocking these enzymes can reactivate
gene expression (9), inhibit growth and survival of tumor cells (10),
and increase the life span of Drosophila (11). To date, 17 HDAC isoforms were described in humans, which are divided into classes
based on sequence homology to yeast enzymes and association with
DNA-binding proteins. Class I HDACs 1-3 and 8 are similar to Rpd3 from
yeast (12-15). Class III enzymes (7 human genes), with yeast Sir2 as
prototype, appear to be unique in their dependence on NAD+
cofactor (16). Class II HDACs 4-7, 9, and 10 show similarity to yeast
Hda1 (17-22) and contain a conserved C-terminal catalytic domain, with
the exception of HDAC6 which has two functional deacetylase domains
arranged in tandem (23). Furthermore, HDACs 4, 5, 7, and 9 contain a
conserved N-terminal region that shows similarity to a co-repressor
protein first isolated from Xenopus laevis named MITR (MEF2 interacting
transcription repressor, see Ref. 24). These
class II members, like MITR, bind directly to myocyte enhancer factor 2 (MEF2) transcription factors and repress their transcriptional activity
(25-28).
MEF2 transcription factors have an essential role in the myogenesis and
morphogenesis of cardiac and skeletal muscle cells (29). MEF2 factors
specifically recognize the control regions of the majority of
muscle-specific genes, as well as nerve-specific and other unrelated
genes (30, 31). In vertebrates, the MEF2 family comprises the members
MEF2a-d, each having several splicing variants. Together, the factors
show more than 85% identity in an 86-amino acid core that is
sufficient for specific DNA binding and dimerization (32). MEF2
proteins belong to the MADS box superfamily of transcription factors
characterized by strong sequence homology in a 58-amino acid DNA
binding domain. This MADS domain is N-terminal in MEF2 proteins and
immediately followed by a conserved 28-amino acid MEF domain that
determines homo- and heterodimerization products of MEF2 family members
and excludes heterodimerization with other MADS box transcription
factors (33, 34). Recently, this laboratory reported the crystal
structure of MEF2a core encompassing amino acids 2-78 bound to cognate
DNA at 1.5 Å resolution (35). The structure revealed the DNA binding
interactions and showed that the MEF domain adopts a configuration
distinct from the respective regions in other MADS box proteins, thus
explaining its dimerization properties. An NMR study of a similar MEF2a
protein-DNA complex corroborated the findings (36).
Considerable evidence supports a role of MEF2 proteins as integrators
of calcium signaling (29) mediated by
calcium/ calmodulin-dependent protein kinase (CaMK), and CaMK
regulation of MEF2 activity appears to play a crucial role in processes
leading to myocardial hypertrophy (26, 37, 38). The responsiveness to
CaMK mediated activation mapped to the MADS/MEF domains of MEF2a, which
were unphosphorylated, however (36). Instead, the targets of CaMK
activity were HDAC4 and HDAC5. Phosphorylation of these proteins leads
to the disruption of the MEF2a-HDAC complexes and resulted in
activation of MEF2-controlled genes (26). Phosphorylated HDAC was found
to bind the chaperone protein 14-3-3, resulting in translocation of
phospho-HDAC into the cytosol (39-41). Consistent with a role in
regulation of MEF2, class II HDACs are expressed predominantly in
tissues where MEF2 levels are highest (heart, skeletal muscle, and
brain). CaMK function is triggered by calmodulin, which in turn is
activated by increased Ca2+ levels in the cell, thus
linking MEF2 function to Ca2+ signaling (29). Recently,
however, a more direct role of Ca2+/calmodulin in the
Ca2+-dependent regulation of MEF2-controlled
gene expression was suggested based on the observation that HDAC4
protein is retained on calmodulin-conjugated resin in the presence of
Ca2+ and that a putative calmodulin-binding motif exists in
the N-terminal region of HDAC4 (42).
In this report, we demonstrate that class II enzyme HDAC5 directly
associates with calmodulin in a Ca2+-dependent
manner. By using a combination of biochemical and biophysical techniques, we define a stable core of HDAC5 that binds to MEF2a and
calmodulin with high affinity. The dissociation constants of the
interaction with MEF2a on the one hand and calmodulin on the other hand
are determined by real time binding experiments. Experimental support
for overlapping MEF2a and calmodulin-binding sites in HDAC5 is
provided. By using purified proteins, we show for the first time that
direct interaction of Ca2+/calmodulin inhibits HDAC5
binding to MEF2a.
Protein Expression and Purification--
Human HDAC5 repressor
core polypeptides were expressed as fusion proteins containing a
His6 tag at the C terminus. Genes encoding for amino acids
140-308 (RprcL) and 140-227 (RprcS),
respectively, of human HDAC5 (17) with a starting methionine added were
cloned into a pET28a plasmid (Novagen) using the NcoI and
HindIII sites and expressed in Escherichia coli
BL21(DE3). Pellets were resuspended in ice-cold Buffer T (25 mM BisTris, 100 mM NaCl, 10 mM
imidazole, pH 7.0) and passed through a cell cracker. Cleared lysate
was loaded on Talon cobalt resin (Clontech) and
washed with Buffer T containing 600 mM NaCl. Bound protein
was eluted with an imidazole gradient to 200 mM,
concentrated into Buffer P (25 mM BisTris, 100 mM NaCl, 0.5 mM EDTA, 1 mM DTT, pH
6.0), and applied to a Poros HS column (Perseptive Biosystems).
RprcL as well as RprcS eluted in a single peak
around 400 mM NaCl. During purification of
RprcL, a major proteolytic breakdown product ("SmFr,"
see "Results") was eluted at 300 mM NaCl and could be
pooled separately. Repressor core proteins were finally passed through
a Superdex S200 HR column (Amersham Biosciences) and concentrated up to
14 mg/ml for storage. Protein concentrations were determined by UV
absorption at 280 nm assuming an extinction coefficient of
The gene encoding for full-length calmodulin (CaM) from X. laevis (44) was excised from the construct pTSNco12CaM (45) and ligated into NcoI/HindIII-digested pET28a
plasmid to yield pET28CaM. CaM was expressed in BL21(DE3) cells,
purified as published (45), lyophilized, and stored as a powder. Prior
to use, CaM was dissolved in water at concentrations up to 30 mg/ml.
Mass spectrometric analysis and Edman degradation of the purified
protein revealed that the N-terminal methionine had been quantitatively removed. The resulting protein encompassing amino acids alanine 2 to
lysine 149 is termed full-length CaM throughout the text.
For biotinylation, purified CaM protein was incubated with
NHS-LC-biotin (Pierce) following a procedure described by Billingsley et al. (46) modified such that an equimolar ratio of
NHS-LC-biotin to protein was chosen rather than an excess of 14 to 1 (46). Thus, on average the attachment of one biotin to each CaM
molecule was achieved as verified by MALDI analysis.
MEF2a core protein (amino acids 2-86 of full-length MEF2a)
encompassing the MADS box and the MEF domain was expressed and purified
following published procedures (35). This core protein is termed MEF2a
throughout the text. All expressed proteins (RprcL, RprcS, RprcS(L187G), CaM, and MEF2a) were
purified to homogeneity as verified by Coomassie staining, N-terminal
sequencing, and mass spectrometric analysis.
Circular Dichroism Spectroscopy--
CD spectra were recorded at
20 °C using a J710 spectropolarimeter (Jasco) equipped with a
thermoelectric temperature controller and interfaced to a personal
computer. Stock solutions of protein samples were prepared in a 25 mM BisTris buffer, pH 6.0, containing 100 mM
KF. Protein solutions of 10 µM or less were used to
obtain the data. The CD spectra were measured at a bandwidth of 2 nm, with a step size of 0.5 nm with 4 s averaging time per point in a
0.1-cm cuvette. Spectra were signal averaged by adding three scans,
base-line corrected, and smoothed using the software provided by Jasco.
Tryptic Fragmentation Experiments--
Protease digestion of
HDAC5 repressor core proteins was performed by treating 1 mg/ml
solution of protein with trypsin from bovine pancreas (Roche Applied
Science) in Proteolysis Buffer (50 mM Tris-HCl, 150 mM NaCl, 2 mM DTT, pH 8.0) at a fixed protein to protease mass ratio at 24 °C in a 1-ml reaction volume. At serial
time points, 55-µl aliquots were taken from the reaction. A fraction
(5 µl) of these aliquots was immediately flash-frozen in liquid
nitrogen and stored for mass spectrometric analysis. The remaining 50 µl were transferred into prepared tubes containing reducing protein
gel loading dye and flash-frozen for SDS-PAGE analysis. Reactions were
repeated several times showing the reproducibility of the proteolytic
fragmentation pattern by using this sample freezing procedure. Digests
of calmodulin or purified calmodulin-repressor core complex,
respectively, were performed in a similar fashion, with the exception
that CaCl2 was added to the reaction buffer to a final
concentration of 2 mM.
Proteolytic fragments were separated by SDS-PAGE (18%) and visualized
by staining with Coomassie Brilliant Blue. Protein gels were blotted on
a polyvinylidene difluoride membrane, and prominent bands were
subjected to N-terminal sequencing by Edman degradation. Mass
spectrometry of the proteolytic mixtures was performed using a Vestec
Voyager Biospectrometry Workstation (Perspective Biosystems). Sinapinic
acid in 67% acetonitrile, 0.03% trifluoroacetic acid was used as a
matrix. Data were analyzed with the Data Explorer software package 4.0 (Applied Biosystems). In ambiguous cases, where Edman degradation
yielded several overlapping sequences for the N termini, further
purification of the proteolytic fragments was performed by reversed
phase chromatography on a Nucleosil 300-5 C8 column (Macherey-Nagel)
applying a trifluoroacetic acid/acetonitrile gradient. Fractions
containing enriched peptide species were lyophilized, resuspended in
30% acetonitrile containing 0.1% trifluoroacetic acid, and used
directly for N-terminal sequencing and MALDI analysis.
Tryptic fragments were assigned according to their experimental average
masses and taking into account the N-terminal sequences utilizing the
FindPept tool of the Expasy package (www.expasy.ch). Band intensities
of full-length calmodulin on SDS-PAGE gels displaying tryptic fragments
of CaM or CaM-RprcS complex, respectively, were recorded on
an Alpha Imager 2200 Documentation and Analysis System device (Alpha
Innotech Corp.), using the spot densitometry module of software package
Imager 2200 version 5.1, and normalized to the respective band
intensity of the undigested sample.
Electrophoretic Mobility Shift Assay (EMSA)--
A 60-mer DNA
oligonucleotide duplex matching the muscle creatine kinase (MCK)
promoter region (47) centered on the MEF2-responsive element with the
sequence
5'GCAGAGGAGACAGCAAAGCGCCCTCTAAAAATAACTCCTTTCCCGGCGACCGAACCCTC (core MEF2 element underlined) was synthesized on an Applied Biosystems DNA synthesizer. MEF2a protein was incubated in concentration ranges
from 1 × 10 Analytical Size Exclusion Chromatography of CaM-Rprc
Complexes--
RprcL or RprcS, respectively,
was mixed with an excess (less than 10%) of CaM (assuming
Real Time Binding Studies by Surface Plasmon Resonance (SPR)
Measurement Using BIAcore--
Binding experiments were performed on a
BIAcore 1000 biosensor system (Amersham Biosciences AB) at 20 °C.
MEF2a protein (15-25 µl of 1 µM polypeptide in 10 mM sodium acetate, pH 6.5) was coupled through its amino
groups to the sensor surface of a CM5 biosensor chip (Amersham
Biosciences) using
N-hydroxysuccinimide/N-ethyl-N'-(dimethylaminopropyl)carbodiimide chemistry as recommended by the manufacturer. After immobilizing 1200 resonance units of MEF2a, remaining activated groups on the sensor
surface were inactivated with 1 mM ethanolamine. For
control experiments, one sensor surface was treated as above in the
absence of MEF2a protein. Interaction experiments between MEF2a and MCK promoter DNA (as above) were carried out with DNA concentrations ranging from 0.5 to 100 nM at a constant flow rate of 10 µl per min using Buffer M (12.5 mM HEPES-Na, pH 7.2, 50 mM NaCl, 5 mM MgCl2, 2 mM DTT) as running buffer. Between injections, the sensor chip was regenerated with 50 µl of Buffer M containing 500 mM NaCl. Binding experiments of HDAC5 RprcL
protein to MEF2a were carried out in Buffer R1 (25 mM
BisTris, 100 mM NaCl, 0.5 mM EDTA, 1 mM DTT, pH 7.0) with protein concentrations ranging from 2 to 35 nM. RprcS has a lower theoretical pI
(8.0) as compared with RprcL (9.95), and its titration
curve shows a plateau between pH 7.0 and 9.0. Therefore, binding
experiments of RprcS to MEF2a were carried out at pH 6.2. For both RprcL and RprcS, it was found that at
the concentrations utilized, the best reproducibility of data was
achieved by diluting protein solutions from a 1 µM stock
with running buffer immediately prior to injection.
To measure the interaction between calmodulin and repressor core
protein, 1500 resonance units of biotinylated calmodulin were
immobilized on a streptavidin-coated SA5 chip (Amersham Biosciences), and an uncoated streptavidin surface was used as a control.
Measurements were performed in Buffer D (10 mM BisTris, 250 mM NaCl, 1 mM CaCl2, pH 6.2) at a
flow rate of 20 µl per min in a range of 2-50 nM RprcS. The sensor surface was regenerated with buffer
containing 500 mM NaCl and 5 mM EGTA instead of
CaCl2. RprcS(L187G) mutant protein was measured
similarly, however, in the concentration range of 50-400
nM.
Nonspecific binding of protein or DNA to the respective control surface
was not detectable under the conditions of our experiments. Data were
evaluated using the BIAevaluation software package provided (Amersham
Biosciences AB). Kinetic constants were obtained by fitting curves to a
single site binding model (A + B = AB).
Inhibition of HDAC5 repressor core binding to MEF2a by CaM was
investigated using a sensor surface with 2000 resonance units of MEF2a
immobilized. Buffer D was used as running buffer at a constant flow
rate of 20 µl/min. RprcS was diluted to 30 nM
from a 1 µM stock solution into Buffer D containing
calmodulin at concentrations ranging from 30 nM to 3 µM. Sample was injected after 10 min of incubation at
room temperature. Control spectra were recorded with sensor surface
activated and deactivated in the absence of MEF2a protein as described above.
HDAC5 Core Encompassing the Proposed MEF2a and CaM Binding
Regions--
HDAC5 contains a C-terminal catalytic domain and an
N-terminal extension that mediated binding to MEF2 proteins (17, 19, 23) (Fig. 1A). The N-terminal
part of the protein is highly homologous to the non-catalytic
transcriptional co-repressor MITR (24, 48) and is termed
"repressor" in this study. It has a glutamine-rich stretch close to
its N terminus (Q in Fig. 1A) followed by a
domain (A in Fig. 1A) that contains the 18-amino acid stretch identified as crucial for interaction with MEF2 (26) centered around the 100% conserved STEVK 5-amino acid motif (amino acids 180-184 in HDAC5). Furthermore, a region homologous to the putative CaM binding domain delineated for HDAC4 (41) is present in
A. The nuclear localization sequence (49) is contained in a
third domain (B in Fig. 1A). Segments
Q, A, and B are highly conserved in
HDACs 4, 5, 7, and 9 and also in MITR from mouse and X. laevis with more than 60% identity between the
proteins.
Assuming that the glutamine-rich stretch constitutes a region with
poorly defined geometry in the protein, we generated, based on homology
alignment, a repressor core construct termed RprcL spanning
amino acids 140-308 of the full-length protein. RprcL encompasses the conserved segments A and B (Fig.
1A). Repressor core boundaries were placed in regions with
low homology and having an amino acid composition indicating possible
loops. We observed that purification of RprcL (see
"Experimental Procedures") consistently yielded, besides the 21-kDa
full-length protein, a smaller product migrating at around 12 kDa on an
SDS gel accounting for between 30 and 50% of total protein. This small
fragment (termed "SmFr") could be separated from full-length
RprcL by ion exchange chromatography (Fig.
2, lanes 2 and 3).
N-terminal sequencing and MALDI mass spectrometry identified SmFr as a
proteolytic breakdown product of RprcL starting at
tryptophan 214 (Fig. 1B and Table
I) and extending to the C-terminal
vector-encoded His6 tag. Addition of phenylmethylsulfonyl
fluoride and EDTA did not reduce the amount of SmFr in the
preparations. The occurrence of this proteolytic fragment, resulting
from cleavage of RprcL by endogenous protease present in
the E. coli lysate between segments A and
B (Fig. 1A), indicated that the region between
the boundaries of these conserved segments is exposed.
We investigated the folding state of RprcL by CD
spectroscopy (Fig. 3). The molar
ellipticity of RprcL exhibits a shoulder at 222 nm,
consistent with
To verify this, we probed RprcL by performing limited
proteolytic fragmentation using trypsin from bovine pancreas. Limited proteolysis is a powerful tool for the identification of boundaries of
stable domains within proteins for structural (50, 51) or folding (52,
53) studies. The potential trypsin cleavage sites are clustered in the
N-terminal and C-terminal portion of RprcL (Fig.
1B); tryptic fragmentation could therefore be expected to
provide information about the relative stabilities of these regions of
the protein. The results of a proteolytic fragmentation of
RprcL at a 500 to 1 protein to trypsin mass ratio is shown in Fig. 2 (lanes 4-7) and depicted schematically in Fig.
1B, with corresponding masses of tryptic fragments as
determined by MALDI listed in Table I. RprcL was
trypsinized leaving no full-length protein detectable after 30 min.
Cleavage occurred at the onset of the reaction mainly in the C-terminal
region of RprcL at amino acids Arg-277, Arg-268, Arg-266,
and Lys-256, leading to tryptic fragments Tr0, Tr1a, Tr1b, and Tr2
(Fig. 1B and Fig. 2). Of these fragments, only Tr1a, Tr1b,
and Tr2 persisted until 30 min in the digest, and then also disappeared
completely. Cleavage occurred at Lys-212 as well, with fragment Tr3
appearing after 10 min in the digest. Further proteolysis occurred in
the central region of the protein, between amino acids Lys-184 and
Lys-196, leading to progressively shorter fragments (Tr5a,b and
Tr6a-c). The only tryptic fragment derived from the most C-terminal
part of the protein that was stable for a significant time corresponded
to a stretch encompassing amino acids Lys-256 to Arg-296 of
RprcL (Tr6b). Fragments Tr4a and 4b, which are derived from
Tr1b and Tr2 and cleaved after Lys-194, appeared at 10 min. At 60 min, they were only weakly detectable by MALDI due to further
degradation resulting in fragment Tr8b, which is resistant to cleavage
by trypsin as it lacks lysine or arginine amino acids (Table I).
At the N terminus, three tryptic sites were targeted by the protease,
leading to truncations after amino acids Arg-145, Arg-147, and finally
Arg-152 of the protein. At 120 min, the reaction mixture consisted
mainly of an assortment of N-terminal pieces of the protein ranging
from less than 3 (Tr8a and -8c) to 4 kDa (Tr7a and Tr7b) and Tr8b from
the central region of RprcL (Fig. 1 and Table I). A strong
Coomassie band from the 120-min sample migrates around 10 kDa in Fig. 2
(lane 7), which seemed to indicate that comparatively large
amounts of Tr3, Tr4a, or Tr4b are still present. However, only Tr3,
encompassing amino acids 140-212 of HDAC5, was unambiguously
identified in this sample by MALDI and Edman degradation of the
corresponding band. The C-terminal half of RprcL was more
rapidly proteolyzed than the N-terminal half, without any larger (>3
kDa) fragments derived from the C-terminal part of the protein detected
after 120 min (Fig. 1B and Table I).
Taken together, these results suggest that the C-terminal part of
RprcL containing segment B of HDAC5 (Fig. 1)
consists largely of random coil with limited tertiary folding. The
N-terminal part, containing segment A with the proposed MEF2
and CaM binding domains (Fig. 1A), on the other hand,
contains a better defined tertiary structure, as numerous potential
tryptic sites in this region were protected even after 120 min (Table
I). We therefore prepared a construct of HDAC5 devoid of the
proteolytically sensitive C-terminal portion. We extended the
protease-resistant fragment Tr3 (Glu-140 to Lys-212) toward the C
terminus by adding a high homology region including the amino acid
stretch PKCW with tryptophan 214. This 4-amino acid motif
PXXW is fully conserved in all class II HDACs containing an
MITR-like repressor domain. The resulting protein, RprcS,
encompasses amino acids 140-227 of HDAC5. The CD spectrum of purified
RprcS (Fig. 3) has a minimum at 222 nm characteristic for
MEF2a Binding Studies--
We analyzed the interaction of
RprcL and RprcS with MEF2a by real time binding
studies (Fig. 4). As a control for the
activity of purified MEF2a used in our experiments, we determined the
binding constant of MEF2a to a DNA oligonucleotide duplex by
electrophoretic mobility shift assay (Fig. 4A) and by
BIAcore with the protein immobilized on a dextran surface (Fig.
4B). The DNA sequence used was derived from the muscle
creatine kinase promoter containing a MEF2-responsive element (see
"Experimental Procedures"). EMSA and BIAcore yielded close to
identical dissociation constants (KD = 0.5-0.6
nM) for the MEF2a/DNA interaction demonstrating that
immobilized MEF2a had retained its activity. Next, the dissociation constants of RprcL and RprcS were determined by
binding the repressor core proteins to immobilized MEF2a over a range
of concentrations. The resulting sensorgrams are shown in Fig. 4,
C and D. Both polypeptides bound immobilized
MEF2a with comparable, high affinity (KD = 6-8
nM) in our experiments, corroborating that the region of HDAC5 important for high affinity interaction with MEF2a is indeed contained within RprcS. In contrast, the C-terminal
extension present in RprcL encompassing segment
B of HDAC5 does not contribute significantly to binding
MEF2a.
Ca2+-dependent Calmodulin/Repressor Core
Interaction--
Youn and co-workers (42) had observed that the HDAC4
enzyme is readily retained on calmodulin-conjugated resin in the
presence of Ca2+. HDAC5 has a sequence homologous to the
CaM binding region proposed for HDAC4 (42). We therefore investigated
whether our HDAC5 repressor core constructs also interacted with CaM
activated by Ca2+. Fig. 5
shows the results of analytical gel permeation experiments of CaM
mixtures with RprcL (Fig. 5A) and
RprcS (Fig. 5B), respectively. In both cases,
equimolar addition of CaM shifted the elution peak of the repressor
core protein quantitatively to higher molecular weights in the presence
of 1 mM CaCl2. SDS-PAGE revealed that this peak
contained both CaM and the repressor core protein in a 1:1 ratio as
judged by the intensity of the bands. Fractions containing complex
could be rechromatographed resulting again in the same symmetric peak
profile. Replacing Ca2+ with 2 mM EGTA in
sample and running buffer resulted in dissociation of the complex with
the components eluting individually. The analytical gel filtration
experiments therefore confirm that class II deacetylases containing an
MITR-like N-terminal domain interact with CaM in a
Ca2+-dependent manner and that the
RprcS region of HDAC5 is sufficient for MEF2a and CaM
binding.
Trypsin Digest of Calmodulin-Repressor Core Complex--
Trypsin
digest of Ca2+-saturated CaM results in cleavage mainly at
amino acids Lys-78, Arg-75, and Lys-76, leading to a mixture of tryptic
fragments containing the N-terminal EF-hand motifs and the C-terminal
EF-hand motifs extended by the respective amino acids of the connecting
linker helix (54). Ca2+/CaM has a dumb-bell shape in
solution, with the linker helix connecting the EF-hand containing
domains being rather flexible. Upon binding to most cognate peptide
motifs, CaM collapses to a globular structure with the "hinge"
amino acids (Lys-78 to Ser-82) permitting this rearrangement. Cognate
peptide motifs are typically present as
With the aim of detecting tryptic sites on either calmodulin or HDAC5
repressor core that are protected on complex formation, we performed limited tryptic
fragmentation of RprcS, CaM, and purified
RprcS-CaM complex (Figs. 6 and
7). Tryptic fragments identified by Edman
degradation and MALDI mass spectrometry are listed in Table
II. The tryptic fragmentation of
RprcS alone followed essentially the same pattern of the
N-terminal portion of RprcL. The difference observed was
the presence of full-length RprcS up to 60 min in the
digest, hinting at increased stability of the shorter construct. Two
further fragments (Tr10a and Tr10b) were identified as
RprcS lacking 5 or 7 N-terminal amino acids (Fig.
6A and Table II). These fragments also persisted longer than
30 min in the digest. CaM fragmentation under the conditions used in
this study (see "Experimental Procedures") yielded predominantly three C-terminal fragments in the digest (Tr1C,
Tr2Ca, and Tr2Cc, compare with Ref. 54). At 120 min, about 50% of full-length CaM was
digested at the protease to protein ratio indicated (Fig. 6). The MALDI
spectrum of the reaction mixture at this time point is shown in the
region from 8,000 to 17,000 Da (Fig. 7A, right). The corresponding tryptic cleavage sites are depicted in a CaM model
(Fig. 7C, right), and the fragments are listed
(Table II). Four sites are located in the region of the flexible linker
helix that connects the EF-hand containing domains of CaM.
The tryptic fragmentation of the CaM-RprcS complex (Fig.
6B) showed a similar pattern as the RprcS digest
as far as the repressor core is concerned, with only minor differences
detected by SDS-PAGE, MALDI, and Edman degradation. In contrast, CaM
was apparently not digested at all, showing no proteolytic breakdown
product of CaM even after 120 min (Fig. 6C). MALDI mass
spectrometry of the 120-min digest sample (Fig. 7B, right)
resulted in only one peak above 8000 Da matching full-length CaM
(16,708 Da). The linker helix thus appears to be fully protected upon
complex formation with RprcS.
Trypsin digest of RprcS in complex with CaM showed a
virtually indistinguishable tryptic pattern for free or bound
RprcS on SDS-PAGE (Fig. 6). However, the stability of
full-length CaM in complex with RprcS allowed for a more
detailed analysis of the MALDI spectra of the digested complex, because
all protein mass signals with the exception of the full-length CaM
signal (16,706 Da) could be attributed to RprcS.
Representative MALDI spectra of RprcS and
CaM-RprcS complex taken at identical time points (60 min)
in the digest are shown (Fig. 7, A and B). Direct
comparison of the two spectra make it evident that the two peaks (4995 and 5910 Da) present in the digest of unbound RprcS are
absent in the digest of CaM/RprcS. These peaks correspond
to tryptic fragments Tr13c and Tr15b (Table II), which extend to the
tryptic site Arg-186 (Fig. 7C, left). Arginine 186 is
apparently not accessible during tryptic fragmentation of the complex,
whereas it is readily available if RprcS alone is digested.
Although MALDI-MS is not a quantitative tool for detection of
proteolytic fragments in a digest, the consistent absence of fragments
Tr13c and Tr15b in the MALDI spectra acquired for the purified
CaM-RprcS complex suggests that the region centered around
RprcS Leu-187 is inaccessible in the complex and that the binding sites for MEF2a and CaM are overlapping. Conversely, tryptic fragments Tr12 and Tr13b (Table II), which extend to Lys-194 (Fig. 7C), still appear in the MALDI spectrum, even though lysine
194 is also embedded in the postulated CaM binding domain. The reason may be found in the nature of the HDAC5 CaM-binding site that is
different from classical CaM-binding motifs. It does not adhere to
conventional 1-5-10 or 1-8-14 rules (55) concerning the positions of
key hydrophobic amino acids and is probably more extended (Table III). In fact, certain CaM-binding motifs
with extended recognition sequences (56, 57) were recently shown to
form hairpins or loop regions rather than a single Ca2+/CaM Competes with MEF2a for Repressor
Core Binding--
We investigated the interdependence of HDAC5
repressor core binding to MEF2a and CaM by SPR measurement with BIAcore
(Fig. 8). First, the interaction between
RprcS and CaM was quantified using biotinylated calmodulin
immobilized on a streptavidin-coated SA5 chip surface.
RprcS protein was passed over immobilized biotinylated calmodulin at the concentrations indicated, yielding a dissociation constant of 3 ± 0.8 nM for the CaM/RprcS
interaction (Fig. 8A). The evolutionary conservation of the
central leucine 187 in class II HDACs as well as MITR (Table III)
implies a central role of this hydrophobic amino acid for the
functionality of HDAC5. Leu-187 is in a region identified as crucial
for MEF2a interaction (Fig. 7C). Hydrophobic amino acids are
also important for binding Ca2+/CaM (Table III), and
mutating these amino acids in CaM-binding motifs to alanine or glycine
has been shown to markedly decrease (5-100-fold) CaM binding affinity
(58). We mutated leucine 187 in RprcS to glycine generating
the mutant protein RprcS(L187G). Real time binding
experiments yielded a KD of ~60 nM for
the interaction of the mutant protein with immobilized CaM (data not
shown), which represents a 20-fold decrease in affinity compared with
wild type.
Having obtained experimental support for overlapping MEF2a- and
CaM-binding sites in RprcS, we proceeded finally to
investigate the influence of HDAC5 repressor core binding to CaM on the
interaction with MEF2a. MEF2a was immobilized on a dextran surface, and
RprcS was passed over the MEF2a-coated surface at a
constant concentration (30 nM) with CaM added in a range
from 30 nM to 3 µM in the mobile phase (Fig.
8B). We found that CaM inhibits RprcS binding to
immobilized MEF2a with increasing concentration. At a molar ratio of
1:100 (RprcS to CaM), repressor core binding to MEF2a was
virtually abolished in this experiment.
In the present study, a stable core domain from the N-terminal
repressor part of human HDAC5 was defined, and the interaction of this
core domain with MEF2a on the one hand and Ca2+ activated
CaM on the other hand was analyzed. Both interactions were found to
have dissociation constants in the nanomolar range. Experimental
support for overlapping MEF2a- and CaM-binding sites was derived from
proteolysis in conjunction with MALDI mass spectrometry and mutant
data. Furthermore, direct interaction of Ca2+/CaM was
demonstrated to inhibit HDAC5 repressor core binding to MEF2a.
MEF2-controlled myogenic genes that are active in early myocyte
development need to be efficiently repressed in the adult muscle cell.
The importance of the underlying molecular mechanisms is exemplified by
the fact that deregulation of the tight control over expression of
myogenic genes or their faulty reactivation, for example, under
pathological challenge can lead to severe diseases such as myocardial
hypertrophy. The repression functionality has been attributed to class
II histone deacetylases such as HDAC5, an enzyme that binds MEF2a
transcription factor with low nanomolar affinity. Ca2+
signaling in the cell in turn regulates class II HDACs. It is well
established that kinases activated by the cellular calcium effector CaM
target class II histone deacetylases by phosphorylation of key serine
amino acids. This leads to removal of these HDACs from the nucleus by
14-3-3 chaperone-mediated translocation into the cytosol. Our results,
and in particular the demonstration that Ca2+ activated CaM
inhibits HDAC5 binding to MEF2a by direct interaction, argue
for a second functionality of CaM in addition to kinase activation
within the framework of myogenic control. This CaM dual action is
illustrated in Fig. 9. In this scenario,
dissociation of HDAC5 from MEF2a is aided by CaM binding directly to
the enzyme in the presence of Ca2+, thus inhibiting the
HDAC5/MEF2a interaction.
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
280 = 8250 M
1
cm
1 (RprcL) and
280 = 5690 M
1 cm
1 (RprcS),
respectively (43). The mutant protein RprcS(L187G) was
generated from the RprcS gene by site-directed mutagenesis (QuickChange, Stratagene) with primers
5'GCACTGAGGTAAAGCTGAGGGGCCAGGAATTCC3' and
5'GGAATTCCTGGCCCCTCAGCTTTACCTCAGTGC3'. RprcS(L187G) protein was expressed and purified exactly as described for RprcS
wild type.
9 to 7 × 10
9
M with 4 × 10
10 M of
32P-radiolabeled MCK promoter DNA in Incubation Buffer
(12.5 mM HEPES-Na, pH 7.2, 50 mM NaCl, 5 mM MgCl2, 2 mM DTT, 250 µg/ml bovine serum albumin, 5% glycerol) and loaded on a 6% polyacrylamide gel with 0.5× Tris borate/EDTA (TBE) as running buffer. 400 V were
applied for 10 min immediately after sample loading followed by 2-3 h
at 200 V at 4 °C. Gels were dried and exposed on BAS-IP MP 2040S
imaging plates (Fuji Film Inc.). Band intensities were recorded with a
Fuji Film BAS-2500 PhosphorImager and quantified with software Advanced
Image Data Analyzer AIDA/2D version 3.11 (Raytest
Isotopenmessgeräte GmbH). The ratio of protein-DNA complex [PD]
to free DNA [D] was obtained from the band intensity data and
plotted against the concentration of free protein [P]. Linear regression yielded the reciprocal of the dissociation constant KD at the chosen conditions following Equation 1,
(Eq. 1)
280 = 2560 M
1
cm
1 for calmodulin) and incubated on ice (1 h) in a 1-ml
volume at a total protein concentration of 2 mg/ml in Buffer C (25 mM BisTris, 100 mM NaCl, 1 mM DTT,
1 mM CaCl2, pH 6.0). The sample was passed through an S200 HR size exclusion column pre-equilibrated in Buffer C
at a flow rate of 0.25 ml/min with fractions collected at 1-min intervals. Peaks containing CaM-Rprc complex were
identified by 18% SDS-PAGE and pooled separate from fractions
containing excess CaM. Pooled complex was rechromatographed to yield a
single symmetric peak in the A280 trace. The
complex was concentrated to 0.5 ml, dialyzed extensively against Buffer
E (10 mM BisTris, 100 mM NaCl, 1 mM
DTT, 2 mM EGTA, pH 6.0), and passed through the S200 HR
column pre-equilibrated in Buffer E. The split peak profiles were
analyzed for protein content by SDS-PAGE. Unbound calmodulin and HDAC5 repressor core proteins were passed through the S200 HR column in
Buffer C under the same conditions as above for comparison.
RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
View larger version (29K):
[in a new window]
Fig. 1.
Human class II HDAC5. A,
domain structure of histone deacetylase 5 in a schematic
representation. An N-terminal PVELR CtBP-binding motif (C)
precedes a glutamine-rich stretch (Q) that is followed by
segment A containing the region implicated in binding to the
MEF2a binding domain and the proposed calmodulin binding domain. The
highly conserved segment B also encompasses the nuclear
localization sequence (NLS). The domain bound by
heterochromatin-binding protein HP1 (H) is followed by a
linker region (L) connecting the repressor half of the
protein to the deacetylase domain (black box). A nuclear
export signal (NES) is located at the extreme C terminus of
the protein. Domain assignments are based on Ref. 47 as well as
homology alignment of human HDAC5 to class II HDACs 4 (NCBI accession
number P56524), 7a (XP_027198), 7b (NP_478057), and 9 (NP_055522) from
human, HDAC5 (Q9Z2V6) and MITR (XP_126877) from mouse, and MITR
(CAB10167) from X. laevis. Numbers
above HDAC5 are residue positions. Boundaries are amino acids 140 and 308 for RprcL and 140-227 for RprcS.
Tryptophan 214 is marked. The N and C termini are indicated.
B, protease cleavage map of HDAC5 repressor core
RprcL. Proteolytic fragments produced by endogenous
protease(s) or trypsin are drawn as gray bars.
RprcL is also shown schematically with black
bands indicating all potential tryptic sites clustered in the N-
and C-terminal portions of the protein. Arrows show actual
cleavage by endogenous protease (gray arrows) and trypsin
(white arrows). The size of the arrows
corresponds to relative accessibility of the sites over time.
Purification of RprcL yielded two minor (End1 and End3) and
a major breakdown product (End2 or "Small Fragment") besides intact
protein. Peptides produced by tryptic digest of full-length
RprcL are shown as shaded bars labeled Tr0 to
Tr8c, with residues present at each end marked by numbers.
Corresponding SDS-PAGE patterns are shown in Fig. 2; fragment sizes as
derived by MALDI mass spectrometric analyses in connection with Edman
degradation are listed in Table I. The illustrations are
proportional.
View larger version (33K):
[in a new window]
Fig. 2.
Proteolytic fragmentation of HDAC5 repressor
core RprcL. Purified RprcL protein is
shown in lane 1. Protein content of the 400 and 300 mM NaCl fractions (lanes 2 and 3)
eluting from the Poros HS column are depicted with FL
indicating full-length RprcL. Protease to protein mass
ratio for the tryptic digest shown was 1:500. Sample time points are
indicated. Tryptic fragments, assigned according to N-terminal
sequencing and MALDI results, are marked with arrows and
denoted according to Fig. 1B and Table I.
Mass spectroscopic analysis of RprcL proteolytic
fragments
-helical secondary structure content. A pronounced
minimum at 203 nm is observed, indicative of a significant proportion
of random coil in the protein. Interestingly, the spectrum of purified
SmFr, which encompasses the C-terminal part of RprcL, is
also that of a random coil (Fig. 3). This implies that the region
contained within SmFr may also be unstructured in the context of
RprcL, explaining the comparatively large random coil
content of RprcL. If so, the secondary structure elements
accounting for the CD signal amplitude at 222 nm would mainly be
located in the N-terminal portion of the protein.
View larger version (25K):
[in a new window]
Fig. 3.
Circular dichroism of purified proteins.
Spectra of HDAC5 repressor core polypeptides RprcL
(···) and RprcS ( ) were subtracted to
yield the difference spectrum marked
L,S (- - -). The
CD spectrum of a purified breakdown product (SmFr, see under
"Results") of RprcL encompassing its C-terminal half is
superimposed (-·-·-). The CD signal (
) of calmodulin was
recorded with 2 mM Ca2+ added to the sample.
The spectrum of RprcS(L187G) mutant protein is identical to
that of RprcS wild type (not shown).
-helices. The second minimum is shifted to 207 nm and is smaller
than in the case of RprcL, clearly indicating a lower random coil content and a more compact structure for the shorter polypeptide. The difference spectrum
L,S (Fig. 3) of
RprcL with RprcS is virtually superimposable on
the spectrum of SmFr, confirming that the C-terminal part of
RprcL largely accounts for its random coil content.
View larger version (50K):
[in a new window]
Fig. 4.
MEF2a interactions. A, EMSA
of MEF2a protein and MCK promoter DNA containing a MEF2-responsive
element (left). Migration of free DNA is shown (lane
f). MEF2a was added in increasing amounts from 1 × 10 9 M (lane 1) to 7 × 10
9 M (lane 8) to a constant
amount (4 × 10
10 M) of
32P-radiolabeled MCK promoter DNA. Migration of the
MEF2a-DNA complex is indicated. Higher aggregates appearing at
increasing MEF2a concentrations are marked with an asterisk.
The ratio of protein-DNA complex [PD] divided by free DNA [D] is
plotted against the concentration of free protein [P]
(right) with linear regression yielding the inverse of the
dissociation constant KD. The KD
value as calculated from four independent EMSA experiments is given.
Sample purity of MEF2a protein used is shown (inset) by a
Coomassie-stained polyacrylamide gel section. B,
determination of the dissociation constant KD for
the MEF2a/MCK promoter DNA complex by SPR measurement. MEF2a was
immobilized to the sensor surface, and MCK promoter DNA was injected at
concentrations from 0.5 × 10
9 to 1 × 10
7 M. Representative sections of the
sensorgrams from one of three experiments are shown. The close to
saturated curve at 100 nM DNA concentration was omitted
from the calculation of KD (top right).
C, the dissociation constant of the MEF2a-RprcL
complex was determined similarly utilizing the same sensor surface as
above with repressor core protein injected in a concentration range
from 2 × 10
9 to 3.5 × 10
8
M. The curve at 35 nM was not included in the
KD (top right) calculation. D,
RprcS was injected at concentrations between 2 × 10
9 and 3.5 × 10
8 M onto
the surface with immobilized MEF2a protein. Errors are standard
deviations from three sets of measurements.
View larger version (48K):
[in a new window]
Fig. 5.
Size exclusion chromatography of
calmodulin-repressor core complexes. A, the
A280 trace of the CaM-RprcL complex
is shown ( ) superimposed on the traces of the individual proteins
shown as dashed lines marked CaM and RprcL. The
protein content of the complex peak in the presence of
CaCl2 is shown below (18% SDS-PAGE). Peak
fractions were pooled, concentrated, and rechromatographed in the
presence of EGTA showing dissociation of the complex
(bottom). B, corresponding
A280 traces (top), SDS gel of the
complex peak with Ca2+ (below), and SDS gel of
the eluted fractions in the presence of EGTA (bottom) for
the CaM-RprcS complex are shown.
-helices in these complexes,
and the primary sequence of the bound peptide evidently dictates both
the extent of unwinding and expansion of the central linker helix of
CaM.
View larger version (43K):
[in a new window]
Fig. 6.
Calmodulin/repressor core interaction,
tryptic fragmentation. A, tryptic fragmentation of
RprcS and CaM. Lanes 1 (RprcS) and
7 (CaM) show the purified proteins used. Full-length
proteins are marked (FL). Protease to protein mass ratio for
tryptic digests (lanes 3-6 and 8-11) was 1:500.
Sample time points are indicated. Tryptic fragments, assigned according
to N-terminal sequencing and MALDI results, are marked and denoted
according to Table II. B, tryptic fragmentation of
CaM-RprcS complex. Lane 1 shows the purified
protein complex used as input. Protease to complex mass ratio was 1:500
in the digest (lanes 2-6). C, the presence of
full-length CaM in the tryptic digest with and without
RprcS bound was quantified by densitometry. Band
intensities are normalized against undigested sample and plotted
against time (right).
View larger version (32K):
[in a new window]
Fig. 7.
Calmodulin/repressor core interaction,
MALDI-MS. A, section of the MALDI spectrum showing the
area between 4200 and 6100 Da of the tryptic digest sample (60 min) of
uncomplexed RprcS with masses corresponding to fragments
Tr13c and Tr15b (see Table II) boxed (left). On
the right, a section of the MALDI spectrum between 8000 and
17,000 Da of the tryptic digest of uncomplexed CaM at 120 min is shown.
Full-length CaM has a molecular mass of 16,706.3 Da. Tryptic peptides
of CaM (see Table II) are boxed (right).
B, corresponding spectra from the tryptic digest of purified
CaM-RprcS complex. Mass peaks corresponding to those
boxed in A are not observed. C,
RprcS is shown as a gray bar with possible
tryptic cleavage sites shown as black bands and actual
cleavage sites marked with arrows. The gray bars
below represent tryptic fragments Tr13c and Tr15b (Table
II). The proposed MEF2 binding domain core (Glu-175 to Leu-192, Ref.
26) is boxed. The putative amphipathic -helix implicated
in CaM binding (41) is underscored. The hydrophobic leucine
187 following a potential tryptic site (Arg-186) is highlighted in
boldface (left). This tryptic site is apparently
protected upon complex formation with CaM (white arrow). The
arrow with an asterisk marks a tryptic site
(Lys-194) within the proposed CaM binding region that is still
accessible in the complex, corresponding to a tryptic fragment (Tr13b)
marked with an asterisk in the MALDI spectra above. The
structure of Ca2+ bound calmodulin based on crystal
coordinates (Protein Data Bank code 1EXR) is shown in a
schematic representation on the right. Tryptic cleavage
sites that are rendered inaccessible upon complex formation with
RprcS are marked with white arrows.
Mass spectroscopic analysis of RprcS and CaM tryptic fragments
-helix upon
complex formation (Table III). Such loop regions could be
cleavage sites in the fragmentation experiments.
Ca2+-dependent CaM-binding motifs (key hydrophobic
amino acids in bold)
View larger version (41K):
[in a new window]
Fig. 8.
Real time calmodulin binding studies.
A, determination of the dissociation constant of CaM and
RprcS in the presence of Ca2+. Repressor core
solutions ranging from 2 × 10 9 to 5 × 10
8 M (boxed on the
right) were passed over biotinylated calmodulin
(bCaM) immobilized on a streptavidin-coated sensor surface.
All curves were used for calculating the KD value
(top right). Curves from one of three experiments are shown.
B, CaM inhibits RprcS binding to MEF2a.
Solutions containing 30 nM repressor core and increasing
amounts of CaM (30 nM to 3 µM) were incubated
(10 min at room temperature) and passed over immobilized MEF2a as shown
schematically on the left. Representative sensorgrams
(right) show progressive reduction of binding with
increasing CaM concentration.
DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
View larger version (49K):
[in a new window]
Fig. 9.
Ca2+/calmodulin dual action
activation model. A, binding of HDAC5 to MEF2 factors
leads to repression of MEF2-dependent genes through
deacetylase activity resulting in chromatin containing hypoacetylated
histone proteins (adapted from Ref. 29). B, increased
Ca2+ levels lead to Ca2+/calmodulin-mediated
activation of calmodulin-dependent kinase
(CaMK), which phosphorylates HDAC5 at specific serine
residues. In addition, direct interaction of
Ca2+/calmodulin with the repressor core releases HDAC5 from
MEF2 proteins. Transcriptional adapter factors such as p300 can now
bind to MEF2 and acetylate histone protein tails. Binding of the 14-3-3 chaperone protein to phosphorylated deacetylase masks the nuclear
localization sequence and exposes the nuclear export signal (40). HDAC5
is subsequently sequestered in the cytosol.
CaM interaction with transcription regulators provides a direct mechanism by which transcription can be regulated in a Ca2+-dependent manner. For example, Ca2+/CaM was demonstrated to modulate selectively the activity of transcription factors of the basic helix-loop-helix family both in vitro and in vivo by directly masking their DNA binding domain (59, 60). More recently, a novel family of transcriptional activators was described for metazoans that also bind CaM with low nanomolar affinity in the presence of Ca2+ (61). In this context, it is notable that prolonged Ca2+ spike-evoked pulses were shown to be integrated into a sustained elevation of the nucleoplasmic CaM concentration in a number of cell lines including smooth muscle (62, 63). Upon Ca2+ signaling, Ca2+/CaM is thus readily available in the myocyte nucleus at elevated levels to inhibit class II HDAC association with MEF2 factors by direct interaction. The measured micromolar concentration of intracellular free CaM (62) and the nanomolar affinity of CaM for RprcS determined in this study suggest that a Ca2+/CaM-HDAC5 complex exists in myocytes and would effectively compete with a MEF2-HDAC5 complex. The high affinity of the CaM/HDAC5 interaction and the conservation of the CaM-binding motif in all class II HDACs known to be involved in myogenic control support a physiological role of CaM binding to these enzymes.
Calmodulin orchestrates cellular regulatory events via interaction with
a host of proteins, being unique in its ability to specifically
recognize a large variety of protein targets. Structural analysis of
CaM with and without Ca2+ and in complex with various CaM
binding domains has provided a wealth of information at the atomic
level and led to the delineation of recurrent motifs for peptide
domains bound by Ca2+-activated CaM. These motifs contain
key hydrophobic amino acids that are arranged in defined register
(Table III). Typically, hydrophobic amino acids spaced 10 or 14 amino
acids apart each form key interactions with either the N- or C-terminal
lobe of CaM depending on the orientation of calmodulin (parallel or
anti-parallel) with respect to its binding site. It appears that
CaM-binding motifs found in class II HDACs do not conform to these
conventional patterns. Our analysis of the HDAC5 repressor core mutant
RprcS(L187G) shows that the mutant protein binds CaM with
markedly reduced affinity compared with wild type, thus corroborating
the key importance of this leucine amino acid in CaM interaction. This
fully conserved leucine is evidently the central hydrophobic amino acid
clamped by one of the two EF-hand containing lobes present in CaM. It is not obvious from the alignment (Table III), however, where the interaction site of the other lobe is located within the HDAC5 primary
sequence. Clearly, high resolution structural studies are required to
resolve the atomic details of HDAC5 binding to CaM. Complementing these
studies with the elucidation of the three-dimensional structure of a
MEF2a-HDAC5 repressor core complex will allow us to identify further
key amino acids that are important for CaM binding but at the same time
do not compromise MEF2a interaction. Mutation of these residues in
full-length HDAC5 may largely reduce or abolish CaM binding by this
protein. Such a structure-based HDAC5 mutant can then be used to verify
the functional role of the direct interaction between class II histone
deacetylase and calmodulin in living myocytes.
![]() |
ACKNOWLEDGEMENTS |
---|
We thank Joachim Krebs for the pTSNco12CaM plasmid and for numerous discussions; Christina M. Grozinger and Stuart L. Schreiber for plasmid containing the HDAC5 gene; Uwe Schlattner and Rudi Fasan for assistance and advice; Rene Brunisholz for Edman sequencing, and Elisabeth Ehler for helpful comments on the manuscript.
![]() |
FOOTNOTES |
---|
* The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
Both authors contributed equally to this work.
§ Recipient of a Liebig fellowship from the Fonds der Chemischen Industrie (Germany).
¶ Supported by the Roche Research Foundation (Switzerland).
Present address: The Burnham Institute, 10901 North Torrey
Pines Rd., La Jolla, CA 92037.
** To whom correspondence should be addressed. Tel.: 41 1 633 2470; Fax: 41 1 633 1150; E-mail: richmond@mol.biol.ethz.ch.
Published, JBC Papers in Press, March 6, 2003, DOI 10.1074/jbc.M301646200
![]() |
ABBREVIATIONS |
---|
The abbreviations used are: HDAC, histone deacetylase; MEF2, myocyte enhancer factor 2; DTT, dithiothreitol; BisTris, 2-[bis(2-hydroxyethyl)amino]-2-(hydroxymethyl)propane-1,3-diol; CaM, calmodulin; CaMK, calcium/calmodulin-dependent protein kinase; MALDI, matrix-assisted laser desorption ionization; NHS, N-hydroxysuccinimide; EMSA, electrophoretic mobility shift assay; SPR, surface plasmon resonance; MCK, muscle creatine kinase.
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1. | Luger, K., Mader, A. W., Richmond, R. K., Sargent, D. F., and Richmond, T. J. (1997) Nature 389, 251-260[CrossRef][Medline] [Order article via Infotrieve] |
2. | Davey, C. A., Sargent, D. F., Luger, K., Maeder, A. W., and Richmond, T. J. (2002) J. Mol. Biol. 319, 1097-1113[CrossRef][Medline] [Order article via Infotrieve] |
3. | Widom, J. (1998) Annu. Rev. Biophys. Biomol. Struct. 27, 285-302[CrossRef][Medline] [Order article via Infotrieve] |
4. | Tsukiyama, T. (2002) Nat. Rev. Mol. Cell. Biol. 3, 422-429[CrossRef][Medline] [Order article via Infotrieve] |
5. |
Jenuwein, T.,
and Allis, C. D.
(2001)
Science
293,
1074-1080 |
6. | Strahl, B. D., and Allis, C. D. (2000) Nature 403, 41-45[CrossRef][Medline] [Order article via Infotrieve] |
7. |
Eberharter, A.,
and Becker, P. B.
(2002)
EMBO Rep.
3,
224-229 |
8. | Grunstein, M. (1997) Nature 389, 349-352[CrossRef][Medline] [Order article via Infotrieve] |
9. |
Wade, P. A.
(2001)
Hum. Mol. Genet.
10,
693-698 |
10. | Johnstone, R. W. (2002) Nat. Rev. Drug Discov. 1, 287-299[CrossRef][Medline] [Order article via Infotrieve] |
11. | Chang, K. T., and Min, K. T. (2002) Ageing Res. Rev. 1, 313-326[CrossRef][Medline] [Order article via Infotrieve] |
12. |
Yang, W. M.,
Inouye, C.,
Zeng, Y.,
Bearss, D.,
and Seto, E.
(1996)
Proc. Natl. Acad. Sci. U. S. A.
93,
12845-12850 |
13. | Taunton, J., Hassig, C. A., and Schreiber, S. L. (1996) Science 272, 408-411[Abstract] |
14. | Yang, W. M., Yao, Y.-L., Sun, J.-M., Davie, J. R., and Seto, E. (1997) J. Biol. Chem. 44, 28001-28007[CrossRef] |
15. | Gray, S. G., and Ekstrom, T. J. (2001) Exp. Cell Res. 262, 75-83[CrossRef][Medline] [Order article via Infotrieve] |
16. |
Guarente, L.
(2000)
Genes Dev.
14,
1021-1026 |
17. |
Grozinger, C. M.,
Hassig, C. A.,
and Schreiber, S. L.
(1999)
Proc. Natl. Acad. Sci. U. S. A.
96,
4868-4873 |
18. | Bertos, R., Wang, A. H., and Yang, X.-J. (2001) Biochem. Cell Biol. 79, 243-252[CrossRef][Medline] [Order article via Infotrieve] |
19. | Fischle, W., Kiermer, V., Dequiedt, F., and Verdin, E. (2001) Biochem. Cell Biol. 79, 337-348[CrossRef][Medline] [Order article via Infotrieve] |
20. |
Zhou, X.,
Marks, P. A.,
Rifkind, R. A.,
and Richon, V. M.
(2001)
Proc. Natl. Acad. Sci. U. S. A.
98,
10572-10577 |
21. |
Fischer, D. D.,
Cai, R.,
Bhatia, U.,
Asselbergs, F. A.,
Song, C.,
Terry, R.,
Trogani, N.,
Widmer, R.,
Atadja, P.,
and Cohen, D.
(2002)
J. Biol. Chem.
277,
6656-6666 |
22. |
Kao, H.-Y.,
Lee, C.-H.,
Komarov, A.,
Han, C. C.,
and Evans, R. M.
(2002)
J. Biol. Chem.
277,
187-193 |
23. |
Verdel, A.,
and Khochbin, S.
(1999)
J. Biol. Chem.
274,
2440-2445 |
24. |
Sparrow, D. B.,
Miska, E. A.,
Langley, E.,
Reynaud-Deonauth, S.,
Kotecha, S.,
Towers, N.,
Spohr, G.,
Kouzarides, T.,
and Mohun, T. J.
(1999)
EMBO J.
18,
5085-5098 |
25. |
Miska, E. A.,
Karlsson, C.,
Langley, E.,
Nielsen, S. J.,
Pines, J.,
and Kouzarides, T.
(1999)
EMBO J.
18,
5099-5107 |
26. |
Lu, J.,
McKinsey, T. A.,
Nicol, R. L.,
and Olson, E. N.
(2000)
Proc. Natl. Acad. Sci. U. S. A.
97,
4070-4075 |
27. |
Wang, A. H.,
Bertos, N. R.,
Vezmar, M.,
Pelletier, N.,
Crosato, M.,
Heng, H. H.,
Th'ng, J.,
Han, J.,
and Yang, X. J.
(1999)
Mol. Cell. Biol.
19,
7816-7827 |
28. |
Lemercier, C.,
Verdel, A.,
Galloo, B.,
Curtet, S.,
Brocard, M.-P.,
and Khochbin, S.
(2000)
J. Biol. Chem.
275,
15594-15599 |
29. | McKinsey, T. A., Zhang, C. L., and Olson, E. N. (2002) Trends Biochem. Sci. 27, 40-47[CrossRef][Medline] [Order article via Infotrieve] |
30. |
Mao, Z.,
Bonni, A.,
Xia, F.,
Nadal-Vicens, M.,
and Greenberg, M. E.
(1999)
Science
286,
785-790 |
31. |
Youn, H. D.,
Sun, L.,
Prywes, R.,
and Liu, J. O.
(1999)
Science
286,
790-793 |
32. | Shore, P., and Sharrocks, A. D. (1995) Eur. J. Biochem. 229, 1-13[Abstract] |
33. | Pollock, R., and Treisman, R. (1991) Genes Dev. 5, 2327-2341[Abstract] |
34. | Molkentin, J. D., Black, B. L., Martin, J. F., and Olson, E. N. (1996) Mol. Cell. Biol. 16, 2627-2636[Abstract] |
35. | Santelli, E., and Richmond, T. J. (2000) J. Mol. Biol. 297, 437-449[CrossRef][Medline] [Order article via Infotrieve] |
36. |
Huang, K.,
Louis, J. M.,
Donaldson, L.,
Lim, F.-L.,
Sharrocks, A. D.,
and Clore, G. M.
(2000)
EMBO J.
19,
2615-2628 |
37. |
Passier, R.,
Zeng, H.,
Frey, N.,
Naya, F. J.,
Nicol, R. L.,
McKinsey, T. A.,
Overbeek, P.,
Richardson, J. A.,
Grant, S. R.,
and Olson, E. N.
(2000)
J. Clin. Invest.
105,
1395-1406 |
38. | Zhang, C. L., McKinsey, T. A., Chang, S., Antos, C. L., Hill, J. A., and Olson, E. N. (2002) Cell 110, 479-488[Medline] [Order article via Infotrieve] |
39. |
Grozinger, C. M.,
and Schreiber, S. L.
(2000)
Proc. Natl. Acad. Sci. U. S. A.
97,
7835-7840 |
40. |
Wang, A. H.,
Kruhlak, M. J.,
Wu, J.,
Bertos, N. R.,
Vezmar, M.,
Posner, B. I.,
Bazett-Jones, D. P.,
and Yang, X. J.
(2000)
Mol. Cell. Biol.
20,
6904-6912 |
41. |
McKinsey, T. A.,
Zhang, C. L.,
and Olson, E. N.
(2001)
Mol. Cell. Biol.
21,
6312-6321 |
42. |
Youn, H.-D.,
Grozinger, C. M.,
and Liu, J. O.
(2000)
J. Biol. Chem.
275,
22563-22567 |
43. | Gill, S. C., and von Hippel, P. H. (1989) Anal. Biochem. 182, 319-425[Medline] [Order article via Infotrieve] |
44. | Chien, Y. H., and Dawid, I. B. (1984) Mol. Cell. Biol. 4, 507-513[Medline] [Order article via Infotrieve] |
45. | Shatzman, A. R., and Rosenberg, M. (1987) Ann. N. Y. Acad. Sci. 478, 233-248 |
46. | Billingsley, M. L., Pennypacker, K. R., Hoover, C. G., Brigati, D. J., and Kincaid, R. L. (1989) Proc. Natl. Acad. Sci. U. S. A. 82, 7585-7589 |
47. | Gossett, L. A., Kelvin, D. J., Sternberg, E. A., and Olson, E. N. (1989) Mol. Cell. Biol. 9, 5022-5033[Medline] [Order article via Infotrieve] |
48. |
Zhang, C. L.,
McKinsey, T. A.,
and Olson, E. N.
(2002)
Mol. Cell. Biol.
22,
7302-7312 |
49. | Hicks, G. R., and Raikhel, N. V. (1995) Annu. Rev. Cell Dev. Biol. 11, 155-188[CrossRef][Medline] [Order article via Infotrieve] |
50. | Gaiser, F., Tan, S., and Richmond, T. J. (2000) J. Mol. Biol. 302, 1119-1127[CrossRef][Medline] [Order article via Infotrieve] |
51. | Zhang, F., Kartner, N., and Lukacs, G. L. (1998) Nat. Struct. Biol. 5, 180-183[Medline] [Order article via Infotrieve] |
52. | Hlodan, R., Tempst, P., and Hartl, F. U. (1995) Nat. Struct. Biol. 2, 587-595[Medline] [Order article via Infotrieve] |
53. | Fontana, A., Polverino de Laureto, P., De Filippis, V., Scaramella, E., and Zambonin, M. (1999) in Proteolytic Enzymes: Tools and Targets (Sterchi, E. E. , and Stocker, W., eds) , pp. 257-284, Springer-Verlag, Heidelberg, Germany |
54. | Thulin, E., Anderron, A., Drakenberg, T., Forsen, S., and Vogel, H. J. E. (1984) Biochemistry 23, 1862-1870[Medline] [Order article via Infotrieve] |
55. |
Rhoads, A. R.,
and Friedberg, F.
(1997)
FASEB J.
11,
331-340 |
56. | Osawa, M., Tokomitsu, H., Swindells, M. B., Kurihara, H., Orita, M., Shibanuma, T., Furuya, T., and Ikura, M. (1999) Nat. Struct. Biol. 6, 819-824[CrossRef][Medline] [Order article via Infotrieve] |
57. | Schuhmacher, M. A., Rivard, A. F., Baechinger, H. P., and Adelmann, J. P. (2001) Nature 410, 1120-1124[CrossRef][Medline] [Order article via Infotrieve] |
58. |
Penheiter, A. R.,
Caride, A. J.,
Enyedi, A.,
and Penniston, J. T.
(2002)
J. Biol. Chem.
277,
17728-17732 |
59. | Corneliussen, B., Holm, M., Waltersson, Y., Onions, J., Hallberg, B., Thornell, A., and Grundstrom, T. (1994) Nature 368, 760-764[CrossRef][Medline] [Order article via Infotrieve] |
60. | Onions, J., Hermann, S., and Grundstrom, T. (2000) Biochemistry 39, 4366-4374[CrossRef][Medline] [Order article via Infotrieve] |
61. |
Bouche, N.,
Scharlat, A.,
Snedden, W.,
Bouchez, D.,
and Fromm, H.
(2002)
J. Biol. Chem.
277,
21851-21861 |
62. | Luby-Phelps, K., Hori, M., Phelps, J. M., and Won, D. (1995) J. Biol. Chem. 37, 21532-21538[CrossRef] |
63. | Deisseroth, K., Heist, K. E., and Tsien, R. W. (1998) Nature 392, 198-202[CrossRef][Medline] [Order article via Infotrieve] |