From the Program in Molecular Medicine, University of Massachusetts Medical School, Worcester, Massachusetts 01605
Received for publication, February 10, 2003, and in revised form, February 19, 2003
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ABSTRACT |
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Post-translational modifications of
histones influence both chromatin structure and the binding and
function of chromatin-associated proteins. A major limitation to
understanding these effects has been the inability to construct
nucleosomes in vitro that harbor homogeneous and
site-specific histone modifications. Here, we describe a native peptide
ligation strategy for generating nucleosomal arrays that can harbor a
wide range of desired histone modifications. As a first test of this
method, we engineered model nucleosomal arrays in which each histone H3
contains a phosphorylated serine at position 10 and performed kinetic
analyses of Gcn5-dependent histone acetyltransferase
activities. Recombinant Gcn5 shows increased histone acetyltransferase
activity on nucleosomal arrays harboring phosphorylated H3 serine 10 and is consistent with peptide studies. However, in contrast to
analyses using peptide substrates, we find that the histone
acetyltransferase activity of the Gcn5-containing SAGA complex is not
stimulated by H3 phosphorylation in the context of nucleosomal arrays.
This difference between peptide and array substrates suggests that the
ability to generate specifically modified nucleosomal arrays should
provide a powerful tool for understanding the effects of
post-translational histone modifications.
Histone proteins, the core protein components of
nucleosomes, play a central role in controlling a wide range of
DNA-dependent processes, including gene expression, DNA
repair, and DNA replication (1). How histones modulate these processes
appears to be dictated in part by their wide range of
post-translational modifications, including acetylation,
phosphorylation, methylation, ribosylation, and ubiquitination (1, 2).
Many of these modifications are found to occur simultaneously,
suggesting the potential for sequential or combinatorial interactions
that may act synergistically or antagonistically to regulate downstream
biological effects (2). For instance, previous studies have used small
peptide substrates to demonstrate that phosphorylation of serine 10 within histone H3 enhances the ability of yeast Gcn5p to acetylate
lysine residues within the same histone N-terminal domain (3, 4). This
dual histone mark appears to be key for transcriptional control
in vivo (3, 4).
A major technical limitation for testing various mechanistic aspects of
individual or combinations of histone modifications is the inability to
generate nucleosomes in vitro that harbor specific histone
modifications. Small peptides encompassing a histone N-terminal tail
that contain very specific and homogeneous modifications can be
synthesized, and such substrates have played a key role in
understanding the potential roles of histone modifications (3-5).
However, peptides lack the more complex determinants of nucleosomes,
such as DNA and multiple histone tails, which are likely to play an
important role in their biological functions. Similarly, nucleosomes
purified from extracts or treated enzymatically can provide more
physiologically relevant substrates, but such nucleosomes lack
homogeneous patterns of modifications.
Here we describe a native chemical ligation strategy that permits the
reconstitution of nucleosomal arrays that harbor a wide range of
individual or combinations of histone modifications, including serine
phosphorylation, lysine acetylation, and lysine methylation. In this
method, solid phase peptide synthesis is used to generate a histone
N-terminal tail domain that contains modified amino acids at any
desired location. Native ligation chemistry is used to assemble
full-length recombinant histone, which is reconstituted into histone
octamers and subsequently assembled into nucleosomal arrays. We
describe the first test of this method in which we have engineered
model nucleosomal arrays harboring a phosphorylated serine at position
10 of each histone H3 protein.
N-terminal Histone H3 Peptide Fragment Synthesis--
Peptides
were synthesized with an automated peptide synthesizer using
commercially available
Fmoc1-amino acids, standard
coupling reagents, and NovaSyn TGT resin preloaded with Fmoc-alanine.
The N-terminal alanine residue was incorporated as a
N-tert-butyloxycarbonyl-protected amino acid. The side
chain-protected, free C-terminal carboxylic acid peptide was cleaved
from the resin and worked up as described (6). This peptide was
dissolved to 1 mM in dimethylformamide and coupled to benzyl mercaptan (20 mM) at 35 °C for 20 h by
the addition of
2-(1-H-benzotriazole-1-yl)-1,1,3,3-tetramethyluronium
hexafluorophosphate (20 mM),
N,N-diisopropylethylamine (40 mM),
and benzyl mercaptan (20 mM) to generate the C-terminal
thioester moiety. Dimethylformamide was removed in vacuo.
Side chain deprotection of the peptide was performed according to
standard protocols. Crude, fully deprotected C-terminal thioester
peptide was purified by C18 reversed-phase high pressure
liquid chromatography (HPLC) (acetonitrile/0.1% trifluoroacetic
acid and water/0.1% trifluoroacetic acid mobile phase), dried,
and weighed. Phosphorylated peptide was additionally purified away from
unphosphorylated peptide by metal affinity chromatography as described
(7). Purity and identity of the peptides were confirmed by electrospray
mass spectroscopy.
C-terminal Histone H3 Protein Fragment Preparation--
A
bacterial expression plasmid containing the C-terminal histone H3
fragment was prepared using standard cloning techniques. Briefly, The
C-terminal portion of histone H3 from amino acids 33 to 135 was
PCR-amplified from a wild-type Xenopus histone H3 expression
plasmid (8) using an upstream primer containing a NdeI
restriction site, a start codon, codons for a minimal Factor Xa
cleavage site, and a cysteine codon
(5'-GCACTCGAGCCATATGATCGAAGGTCGTTGTGGCGGAGTCAAGAAACCTCACCGTTAC-3') and a downstream primer containing a BglII restriction site
(5'-AGCTCGCAATAGATCTAAGCCCTCTCGCCTCGGATTCT-3'). The resulting product
was digested and ligated into a pET11c expression vector. The identity
of the expression plasmid was confirmed by DNA sequencing.
The C-terminal histone H3 protein fragment was expressed and purified
as previously described (8) with several modifications: expression was
performed in Escherichia coli BL21 cells lacking a pLys
expression plasmid; at least 5 mM DTT was present
throughout the purification steps to prevent intra- and intermolecular
disulfide formation; and following the final dialysis step, the protein was used directly.
The N-terminal cysteine of the histone H3 protein fragment was exposed
with Factor Xa protease under the following conditions: 0.5 mg/ml
histone fragment protein, 1.0 µg/ml Factor Xa protease, 20.0 mM Tris-HCl, pH 8.0, 100.0 mM NaCl, 2.0 mM CaCl2, 1.25 mM DTT. Digestion
proceeded for 10 min at 25 °C and then quenched with
phenylmethylsulfonyl fluoride at a final concentration of 1.0 mM. Salts were removed by dialysis in water/0.1%
trifluoroacetic acid, and the resulting solution was dried by
lyophilization. Undigested and overdigested product was purified
from the desired protein product by preparative C4
reversed-phase HPLC using a mixed solvent system of acetonitrile/0.1%
trifluoroacetic acid and water/0.1% trifluoroacetic acid.
Histone H3 Ligation and Purification--
2 mg/ml C-terminal H3
protein fragment and 2.5 mg/ml N-terminal H3 peptide fragment were
ligated at 25 °C for 20 h in 3 M guanidine-HCl, 0.1 M potassium phosphate, pH 7.9, in the presence of 1%
benzyl mercaptan and 1% thiophenol, similar to Ref. 9. The crude
reaction mixture was dissolved into 25:75:0.1
acetonitrile/water/trifluoroacetic acid, diluted 5-fold into 200 mM NaCl SAU-200 buffer (8), loaded onto a Hi-Trap
sulfopropyl-Sepharose high performance ion exchange column, and eluted
with a linear 200-600 mM NaCl gradient. Salts were removed
by dialysis against 5 mM DTT. Ligated histone H3 was
quantified by comparison with a known quantity of wild-type Xenopus recombinant histone H3 on an 18% SDS-PAGE gel,
stained by Coomassie Blue.
Histone Octamer and Nucleosomal Array Preparation--
To
generate histone octamers, equivalent amounts of denatured ligated
histone H3 and denatured recombinantly expressed and purified
Xenopus H2A, H2B, and H4 were dialyzed into 2.0 M NaCl, purified by gel filtration chromatography, and then
quantified by absorbance (8). Nucleosomal arrays were generated with
208-11 DNA template and then analyzed by EcoRI
digestion/native gel analysis as described (10). Array concentration
was determined by DNA absorbance.
Purification of Chromatin Modifying Enzymes--
Recombinant
Gcn5p protein was overexpressed in E. coli BL21 cells and
purified as described (11). Gcn5p concentration was quantified by
Bradford assay in addition to 10% SDS-PAGE followed by Coomassie Blue
staining. Gcn5p-containing SAGA complex was purified from whole cell
yeast extracts (strain CY396) as described (11). SAGA concentration was
determined by comparative Western blotting against known amounts of
recombinant Gcn5p using anti-Gcn5p antibody (sc-9078; Santa Cruz).
SWI/SNF complex was purified from whole cell extracts (strain CY944) as
described (12).
SWI/SNF Remodeling--
Nucleosomal arrays (1 nM) were assayed for SWI/SNF (2-6 nM)
remodeling in buffer containing 20 mM Tris-HCl, pH 8.0, 50 mM NaCl, 5 mM MgCl2, 100 µg/ml
BSA, and 1 mM DTT as described previously (10).
HAT Assays--
Liquid nucleosomal HAT assays were performed
with recombinant Gcn5p protein or purified SAGA complex in HAT buffer
(50 mM Tris, pH 7.5, 5% glycerol, 0.125 mM
EDTA, 50 mM KCl, 1 mM DTT, 1 mM
PMSF, 10 mM sodium butyrate, 3.33 µM
3H-acetyl-CoA, (4.7 Ci/mmol), 6.66 µM
acetyl-CoA). Phosphatase inhibitors (1 mM
Na3VO4, 5 mM NaF) were added to all reactions.
Generating full-length histone proteins by native ligation
chemistry requires two components: (i) an N-terminal protein fragment that terminates with a C-terminal thioester, and (ii) a C-terminal protein fragment that begins with an N-terminal cysteine residue (Fig.
1) (13). Mixing these two components
initiates a two-step chemical reaction producing a full-length ligated
product in which the N-terminal fragment is linked to the C-terminal
fragment via a canonical peptide bond (13).
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS AND DISCUSSION
REFERENCES
EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS AND DISCUSSION
REFERENCES
RESULTS AND DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS AND DISCUSSION
REFERENCES
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Fig. 1.
Native chemical ligation strategy for
generating histone H3 proteins containing specifically modified
N-terminal residues. An N-terminal peptide fragment of histone H3
that contains specifically modified amino acid residues (in this
example a phosphoserine residue denoted by an encircled P)
and a C-terminal thioester moiety (COSR), is produced by standard solid
phase peptide synthesis on an acid-hypersensitive support
(left). A C-terminal protein fragment of histone H3
containing an N-terminal cysteine residue is generated by proteolytic
trimming of recombinant protein (right). Reaction of these
two fragments in the presence of thiol reagents produces native
full-length histone H3 containing the modifications of interest.
Synthesis of N-terminal Histone H3 Peptide Fragments--
Since
most histone H3 post-translational modifications occur within the first
28 amino acids (2), standard Fmoc-based, solid-phase peptide synthesis
can be used to create histone N-terminal domains (1-31) that contain a
wide range of modifications in this region. For these experiments we
chose to study serine or phosphoserine incorporated at position 10 of
H3. To install the desired C-terminal thioester residue, a strategy of
selectively exposing and modifying the peptide C terminus was utilized
(14) (Fig. 1). Initial solid-phase peptide synthesis on an
acid-hypersensitive resin allows cleavage of the peptides from the
resin with weak acid, selectively unmasking the C terminus, which can
then be selectively derivatized to generate a thioester. Side chain
deprotection with strong acid followed by HPLC was sufficient to
generate the desired unphosphorylated H3 peptide fragment in pure form
(Fig. 2A). Because some
dephosphorylation occurred during the synthetic process, an additional
metal affinity purification step was used to obtain homogeneous,
phosphorylated H3 peptide (7). The identity of the peptides was
confirmed by electrospray mass spectrometry, and their purity
demonstrated by analytical HPLC (data not shown).
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C-terminal H3 Protein Fragment-- Since the C-terminal fragment of histone H3 must contain an N-terminal cysteine residue, we used site-directed mutagenesis of a Xenopus histone H3 bacterial expression construct to replace the threonine codon at position 32 with a cysteine codon. Importantly, this T32C substitution is not expected to disrupt nucleosome structure since Thr-32 is located outside the structured domain of the histone octamer (15) and is not well conserved among eukaryotes. Because the N-terminal methionine of recombinantly expressed proteins is not readily removed, a minimal Factor Xa cleavage site was also introduced directly N-terminal to the cysteine at position 32. Consequently, cleavage of purified, recombinant T32C histone H3 with Factor Xa exposes the N-terminal cysteine. Reversed-phase HPLC was used to isolate the desired C-terminal H3 protein fragment (32-135). The identity and purity of this fragment was confirmed by SDS-PAGE (Fig. 2A) and MALDI-TOF mass spectral analysis (data not shown).
H3 Protein Ligation-- Full-length unphosphorylated (T32C-H3) and Ser-10 phosphorylated histone H3 (S10Phos-H3) polypeptides were generated by mixing the N-terminal peptide and C-terminal protein fragments under denaturing conditions in the presence of thiol reagents (9, 13). Note the S10Phos-H3 polypeptide also contains the T32C amino acid change. Successful chemical ligation was monitored by SDS-PAGE, which detected formation of a protein product that co-migrates with recombinant wild-type Xenopus histone H3 (Fig. 2A). Further analysis of the T32C-H3 reaction mixture by MALDI-TOF revealed that the mass of the largest molecular weight species corresponds to the expected mass for full-length T32C histone H3 (Fig. 2A). Because the ligation reactions were performed under conditions of excess peptide and because the reaction proceeds to roughly 50% completion, ion exchange chromatography was used to purify the ligated products to homogeneity (Fig. 2A).
Octamer and Nucleosome Assembly-- To generate histone octamer containing either the ligated T32C-H3 or S10Phos-H3, the ligated H3 proteins were denatured and mixed with denatured, recombinant Xenopus histones H4, H2A, and H2B in equal ratios, and the renatured histone octamers were isolated by gel filtration as described previously (8). No significant differences in the efficiency of octamer reconstitution were detected when ligated T32C-H3 or S10Phos-H3 histones were substituted for wild-type H3 (Fig. 2B). To make certain that S10Phos-H3 was not dephosphorylated during the course of octamer assembly or purification, MALDI-TOF mass spectral analysis was performed on the purified octamer (Fig. 2C). Each of the four different histone polypeptides was detected, and the largest molecular weight component corresponds to the expected mass of the singly phosphorylated histone H3. Moreover, no dephosphorylated H3 was detectable, confirming that this histone octamer is homogeneous for H3 serine 10 phosphorylation.
Chicken erythrocyte or recombinant Xenopus histone octamers
were used to assemble model nucleosomal arrays using a DNA template composed of eleven head to tail repeats of a 208-bp 5 S rRNA
gene from Lytechinus variegatus (the 208-11 template; see
Fig. 3A; (16)). Each 5 S
repeat can rotationally and translationally position a nucleosome after
in vitro salt dialysis reconstitution, yielding a positioned
array of nucleosomes (17). This 208-11 template also contains a unique
SalI restriction enzyme site in the central repeat of the
array (see Fig. 3A; (16)). To monitor nucleosome assembly,
we digested the reconstituted nucleosomal arrays with EcoRI.
Since each 5 S rDNA repeat in the 208-11 array template is bordered by
EcoRI restriction sites (Fig. 3A), cleavage releases either a 208-bp free DNA fragment or a mononucleosome that can
be identified due to its slower mobility after native gel
electrophoresis. Comparison of the resolved digestion products (Fig.
3B) indicates that octamers containing either ligated
T32C-H3 or S10Phos-H3 can be incorporated into the DNA template with
efficiencies similar to that of chicken erythrocyte or wild-type
(Thr-32) recombinant octamers (Fig. 3B and data not shown).
Additionally, increasing ratios of ligated T32C-H3-containing octamer
to DNA template results in an expected increase in array saturation as
demonstrated by the increase of mononucleosome and oligonucleosome
products relative to free DNA. Together these results suggest that
octamers generated from ligated histones are competent to form
nucleosomal arrays.
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SWI/SNF-dependent Remodeling of S10Phos-H3 Nucleosomal Arrays-- Restriction enzyme accessibility assays were performed to further characterize the nucleosomal arrays containing ligated histones and to investigate whether ATP-dependent nucleosome remodeling by yeast SWI/SNF might be influenced by H3-serine 10 phosphorylation. Radiolabeled nucleosomal arrays containing chicken octamer, recombinant wild-type Xenopus H3, and ligated T32C-H3 and S10Phos-H3 were prepared and characterized by EcoRI digestion (data not shown). Arrays with a similar degree of nucleosome saturation were then subjected to digestion with SalI, which yields a quantitative measurement of the accessibility of DNA within the central nucleosome (Fig. 3A) (16). In the absence of ATP, nucleosomal arrays reconstituted with ligated histones exhibited nearly identical occlusion of the SalI site compared with arrays reconstituted with wild-type recombinant or chicken octamers (Fig. 3 and data not shown). Thus, the DNA wrapped onto ligated histone octamers does not appear to be inherently more accessible to restriction enzymes. Furthermore, when ATP was added to initiate SWI/SNF-dependent remodeling, we found that all of the arrays were excellent substrates for SWI/SNF action, showing similar kinetics over different enzyme and array concentrations (Fig. 3 and data not shown). For each array, SWI/SNF action enhanced SalI digestion kinetics by over 30-fold. Since this remodeling assay appears to measure the rate of nucleosome movements by ATP-dependent remodeling enzymes (18), these data suggest that H3 serine 10 phosphorylation does not grossly alter the ability of SWI/SNF to mobilize nucleosomes.
Histone Acetyltransferase Assays--
Previous studies with
peptide substrates demonstrated that H3 serine 10 phosphorylation
enhances the histone acetyltransferase activities of recombinant Gcn5p
and native Gcn5-containing HAT complexes, such as SAGA (3, 4). To
investigate whether Ser-10 phosphorylation within the context of
nucleosomal arrays also stimulates the HAT activity of Gcn5p, we first
performed radioactive HAT assays using recombinant Gcn5p and
nucleosomal arrays containing recombinant wild-type Xenopus
H3, T32C-H3, or S10Phos-H3. Importantly, our HAT assays contained 50 mM monovalent cation, which facilitates the nucleosomal HAT
activity of rGcn5p (19). The extent of 3H-acetate
incorporation into the nucleosomal arrays was plotted as a function of
time, and time points in the linear portion of the reaction were fit to
determine initial velocities (Fig.
4A). Initial velocities for
unphosphorylated nucleosomal array substrates containing recombinant
wild-type Xenopus H3 and ligated T32C-H3 were nearly
equivalent. In contrast, the initial velocity for the phosphorylated
nucleosomal array substrate was increased greater than 2-fold relative
to the unphosphorylated array, consistent with previous studies
demonstrating enhanced rGcn5 HAT activity with phosphorylated peptide
substrates.
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To determine whether this increased activity toward phosphorylated nucleosomal arrays extended to a native Gcn5p-containing protein complex, we performed nucleosomal HAT assays using the multi-subunit SAGA complex (Fig. 4B). Contrary to the increased HAT activity of Gcn5p on the phosphorylated array, the initial velocities for HAT activity of SAGA complex did not show a difference between the phosphorylated and unphosphorylated array substrates. Because this result was contrary to previous results that used SAGA and phosphorylated peptides (3), a more extensive kinetic characterization was performed. To ensure that the HAT assays were not performed under saturating substrate concentrations, where differences in binding affinity would be largely masked, assays were repeated over a wide range of nucleosomal array concentrations for all three substrates. Plots of initial velocities of HAT activity as a function of array concentration (Fig. 4C) show that SAGA exhibits saturation kinetics for all three nucleosomal array substrates. Strikingly, comparison of initial velocities of SAGA for phosphorylated and unphosphorylated arrays over a range of substrate concentrations confirms an absence of any significant difference in initial velocities. Furthermore, SDS-PAGE analysis of these HAT reactions demonstrates that the H3 subunit remains the preferred acetylation substrate (Fig. 4D) and that no detectable difference in the level of acetylation is detectable among the three substrate arrays.
The ability of Gcn5p and SAGA to efficiently utilize nucleosomes composed of either wild-type or ligated histones provides further support for the integrity of nucleosomal arrays reconstituted with histones generated by native chemical ligation. Moreover, the ability to generate homogeneous, phosphorylated nucleosomal arrays has allowed us to determine that the nucleosomal HAT activity of recombinant Gcn5p is enhanced by H3 phosphorylation and that the nucleosomal HAT activity of the native SAGA complex is insensitive to H3 phosphorylation. This latter result was particularly surprising since previous studies that used peptide substrates demonstrated stimulation of SAGA activity by H3 phosphorylation (3). In the case of rGcn5p it is known that the phosphorylation-dependent stimulation of HAT activity is due to an increased affinity for the peptide substrate (6- to 10-fold; (3, 4)). In the case of SAGA complex, the failure of H3 phosphorylation to stimulate nucleosomal HAT activity may be due to other subunits (e.g. Ada2p) contributing to a different mode of histone tail binding. In addition, SAGA may make additional contacts with nucleosomes that are simply not possible with small peptide substrates; for instance SAGA may interact with multiple histone tails, or the ability of SAGA to bind with high affinity to DNA (20) may also contribute to a different mode of nucleosomal substrate binding. We note that our biochemical studies are consistent with several recent reports that suggest that histone H3 phosphorylation is not always coupled to enhanced histone acetylation in vivo (21, 22).
Concluding Remarks--
Although our studies have used native
peptide ligation to generate homogeneous, phosphorylated nucleosomal
arrays, this strategy has the potential for generating and analyzing
nucleosomal arrays that harbor a wide range of histone
post-translational modification. Importantly, the ligation chemistry is
compatible with peptides that harbor specific acetylated or methylated
lysine residues. Furthermore, nucleosomal arrays could be generated
with monomethylated, dimethylated, or trimethylated lysine at position
9 of histone H3. The H3 dimethylated Lys-9 arrays could then
serve as substrates for reconstitution of heterochromatin-like
structures by incorporation of heterochromatin protein 1 (23).
Likewise, nucleosomal arrays harboring methylated lysine at position 4 of histone H3 could be reconstituted to investigate how this
modification controls the functions of ATP-dependent
remodeling enzymes, such as the mammalian NuRD complex (5). In essence,
the utility of this native peptide ligation strategy provides a unique
tool for dissecting how the complexities of histone modifications
control the structure and function of the chromatin fiber.
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ACKNOWLEDGEMENTS |
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We thank Bob Carraway and the University of Massachusetts Medical School peptide synthesis facility for enhanced access to the facility, Biliang Zhang for the use of his HPLC, and Peter Horn and Corey Smith for helpful comments on the manuscript.
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FOOTNOTES |
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* This work was supported by National Institutes of Health Grants GM49650 (to C. L. P.) and F32 AI10611 (to M. S.-K.), NCI National Institutes of Health Grant CA82834 (to C. L. P.), and a postdoctoral fellowship from the Leukemia and Lymphoma Society of America (to C. J. F.).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.
To whom correspondence should be addressed: Program in
Molecular Medicine, University of Massachusetts Medical School, 373 Plantation St., Biotech 2, Suite 210, Worcester, MA 01605. Tel.: 508-856-5858; Fax: 508-856-5011; E-mail:
craig.peterson@umassmed.edu.
Published, JBC Papers in Press, February 20, 2003, DOI 10.1074/jbc.M301445200
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ABBREVIATIONS |
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The abbreviations used are: Fmoc, N-(9-fluorenyl)methoxycarbonyl; HPLC, high pressure liquid chromatography; DTT, dithiothreitol; MALDI-TOF, matrix-assisted laser desorption ionization-time of flight.
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