From the Division of Cellular and Molecular Biology,
Dana-Farber Cancer Institute and Harvard Medical School, Boston,
Massachusetts 02115, the § Institute of Molecular Biology,
University of Vienna, A-1030 Vienna, Austria, the ** Department of
Biochemistry and Winship Cancer Center, Emory University School of
Medicine, Atlanta, Georgia 30322, and the
§§ Harvard Microchemistry Facility, Harvard
Biological Laboratories, Cambridge, Massachusetts 02138
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ABSTRACT |
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Carboxymethylation of proteins is a highly
conserved means of regulation in eukaryotic cells. The protein
phosphatase 2A (PP2A) catalytic (C) subunit is reversibly methylated at
its carboxyl terminus by specific methyltransferase and methylesterase
enzymes which have been purified, but not cloned. Carboxymethylation
affects PP2A activity and varies during the cell cycle. Here, we report that substitution of glutamine for either of two putative active site
histidines in the PP2A C subunit results in inactivation of PP2A and
formation of stable complexes between PP2A and several cellular
proteins. One of these cellular proteins, herein named protein
phosphatase methylesterase-1 (PME-1), was purified and microsequenced,
and its cDNA was cloned. PME-1 is conserved from yeast to human and
contains a motif found in lipases having a catalytic triad-activated
serine as their active site nucleophile. Bacterially expressed PME-1
demethylated PP2A C subunit in vitro, and okadaic acid, a
known inhibitor of the PP2A methylesterase, inhibited this reaction. To
our knowledge, PME-1 represents the first mammalian protein
methylesterase to be cloned. Several lines of evidence indicate that,
although there appears to be a role for C subunit carboxyl-terminal
amino acids in PME-1 binding, amino acids other than those at the
extreme carboxyl terminus of the C subunit also play an important role
in PME-1 binding to a catalytically inactive mutant.
Protein phosphatase 2A
(PP2A)1 is a highly conserved
serine/threonine phosphatase involved in the regulation of a wide
variety of enzymes, signal transduction pathways, and cellular events (1, 2). Consonant with its diverse roles, subpopulations of PP2A have
been found to localize to the nucleus, cytoplasm, cytoskeleton, and
membranes (3-6). The smallest functional unit of PP2A thought to exist
in vivo consists of a heterodimer between a catalytic 36-kDa
subunit, termed C, and a constant regulatory 63-kDa subunit, termed A
(7). This A/C heterodimer often further complexes with a member of one
of three additional cellular regulatory subunit families termed B (or
B55), B' (or B56), and B" (or PR72/120) (1). In cells stably
transformed by the middle tumor antigen (MT) of polyomavirus, MT
substitutes for the B subunit in a small portion (~10%)
(8)2 of PP2A complexes (9).
MT·PP2A complex formation is known to be important for MT-mediated
transformation (10-13), but the precise functional consequences of MT
association with PP2A are still being elucidated.
Efforts aimed at understanding PP2A regulation have uncovered a complex
set of noncovalent and covalent mechanisms. These include association
with different regulatory subunits (1), association with heat stable
inhibitors (14), action of a phosphotyrosyl activator protein (15),
lipid binding (16), phosphorylation (17), and methylation (18-22).
These mechanisms affect the catalytic activity, substrate specificity,
and cellular localization of PP2A. However, little is known about the
molecular bases of their effects, and even less about how these effects
might be coordinated and integrated to orchestrate PP2A functions
throughout the cell.
The carboxyl terminus of the PP2A C subunit seems to be a focal point
for regulation of PP2A. In addition to containing the amino acids
identified as the sites of tyrosine phosphorylation and methylation,
this region contains residues essential for stable binding of the B
regulatory subunit (23). It is possible that these three events may
influence one another. We have recently shown that substitution of
tyrosine 307, the site of tyrosine phosphorylation, with a negatively
charged amino acid abolishes both B subunit binding (23) and
methylation of the C
subunit.3 In contrast, MT
does not require these residues to form PP2A heterotrimers (23),
raising the possibility that different B-type subunits might be
differentially affected by, or differentially affect, covalent
modification at the carboxyl terminus.
The first indication that PP2A C subunit was methylated involved two
observations. Rundell (18) showed that a 36-kDa SV40 small tumor
antigen (ST)-associated cellular protein was a major acceptor of the
methyl group from radiolabeled S-adenosylmethionine added to
cell extracts. Subsequently, this ST-associated cellular protein was
reported to be the PP2A C subunit (9). More recently, several groups
showed that PP2A C subunit is indeed methylated and reported that this
methylation is reversible and occurs on the carboxyl-terminal amino
acid leucine 309, forming a methyl ester (19-22). In addition, the
PP2A methyltransferase and methylesterase enzymes have been purified
and an initial characterization performed (19, 24). However, cloning of
cDNAs for these enzymes has not been reported and their primary
sequences are unknown.
Although methyltransferase and methylesterase enzymes specific for PP2A
have been purified, the regulation and role(s) of PP2A methylation have
only begun to be elucidated. Based on differential antibody recognition
of methylated and non-methylated C subunits, PP2A has been reported to
undergo cell cycle dependent changes in methylation (6). This suggests
that PP2A methylation may participate in the regulation of, or be
regulated by, cell cycle progression. cAMP was found to stimulate PP2A
methylation in Xenopus egg lysates (25), suggesting that
this second messenger may be involved in the regulation of PP2A
methylation. The activity of PP2A toward phosphorylase a and
a phosphopeptide substrate was reported to increase approximately
2-fold upon methylation (21). Greater effects might be observed with
other substrates, given that the effects of some other mechanisms of
PP2A regulation, such as B-type subunit association and heat-stable
inhibitor proteins, have been shown to be highly substrate
dependent. Another possibility is that PP2A methylation might affect
B-type subunit association, or vice versa.
The exact determinants on PP2A essential for functional recognition by
the PP2A methyltransferase and methylesterase enzymes are unknown,
although they must include more than just the highly conserved carboxyl
terminus itself. Xie and Clarke (22) showed that a synthetic
carboxyl-terminal PP2A C subunit octapeptide functions neither as a
PP2A methyltransferase substrate nor as an inhibitor, and Lee and
co-workers (24) demonstrated that a methylated synthetic
carboxyl-terminal tetrapeptide also functions neither as a PP2A
methylesterase substrate nor as an inhibitor. In the latter study, a
600-fold excess of unmethylated PP2A C subunit was found to inhibit the
PP2A methylesterase by 50%, while a 106-fold excess of a
carboxyl-terminal C subunit decapeptide did not inhibit the
methylesterase at all. Collectively, these results suggest that both
enzymes make essential contacts with C subunit residues that are not in
the carboxyl terminus. One hint as to where such contacts might be
located comes from the observation that the potent PP2A inhibitors
okadaic acid and microcystin-LR also inhibit the PP2A methyltransferase
and/or methylesterase enzymes (19, 20, 24). It has been suggested that
this inhibition may be due to these inhibitors binding in part to the
carboxyl-terminal region of the C subunit, interfering with the binding
of the methyltransferase and methylesterase. However, an equally
attractive possibility is that these two enzymes interact with residues
in or around the active site of PP2A.
In this study, we report that an individual substitution of either of
two PP2A C subunit active site histidines with glutamine results in a
catalytically inactive PP2A mutant that forms a stable complex with
several cellular proteins not bound stably by wt C subunit. The
formation of this stable complex enabled us to purify, microsequence,
and clone the first of these cellular proteins, which we have
designated PME-1. PME-1 was identified as a PP2A methylesterase by the
ability of the bacterially expressed and purified enzyme to demethylate
PP2A C subunit. To our knowledge, this is the first mammalian
methylesterase for which protein or cDNA sequences have been
reported. Data base searches reveal a single homolog in
Saccharomyces cerevisiae, as well as complete or partial
sequences, respectively, for homologs in Caenorhabditis elegans and zebrafish, indicating that this enzyme is conserved across eukaryotes. PME-1 contains a motif found in lipases that utilize
a catalytic triad-activated serine as their active site nucleophile,
and has other scattered homology with other lipases in which this motif
is conserved. Based on a number of results, we propose that the
specificity of PME-1 for PP2A may in part be determined by interaction
with residues or metals in or near the PP2A active site.
Plasmids and Mutagenesis--
Site-directed mutagenesis was
performed on a HA-tagged wt C subunit cDNA cloned in the pcDNA
I Amp vector (23) using the Muta-Gene Phagemid In Vitro
Mutagenesis Kit according to the manufacturer's instructions
(Bio-Rad). The entire cDNA of both H59Q and H118Q was sequenced to
confirm successful mutagenesis and to ensure that no additional
mutation occurred. Mutant C subunit cDNAs including the HA tag
coding sequence were cloned into the dexamethasone-inducible vector,
pGRE5-2 (26). The construction of a pGRE5-2 vector expressing HA-tagged
wt PP2A C subunit has been previously described (23).
To make a PME-1 construct (pPS.PME-1) to be used for sequencing and
in vitro transcription/translation, the PME-1 cDNA
product generated by RT-PCR (see below) was inserted via blunt end
ligation into an SrfI site in the PCR-ScriptTM
SK(+) vector using the PCR-ScriptTM SK(+) kit (Stratagene).
Cells and Cell Culture--
NIH3T3 lines expressing wt
polyomavirus MT and a geneticin resistance gene (27) were transfected
by the calcium phosphate precipitation method (28), and individual
clones and mixtures of clones expressing wt C subunit (wt C sub), H59Q,
H118Q, or empty vector (GREonly) were selected and maintained as
described previously (23). H118Q expressed at a level well below that of endogenous wt C subunit, while H59Q expressed at a level equal to or
greater than the wt level. Although the inducible vector, pGRE5-2, was
used to express these proteins, their levels were substantial in the
absence of dexamethasone; for this reason, GREonly cells were used as a
negative control in this study rather than uninduced wt or mutant C
subunit expressing cells. However, dexamethasone treatment was used
throughout to obtain maximal expression of the C subunits.
Radiolabeling of Cells--
For metabolic labeling of cells with
methionine, subconfluent dishes of cells were labeled for 5 h with
[35S]methionine (300 µCi/ml) in Dulbecco's modified
Eagle's medium minus methionine supplemented with 0.5% dialyzed fetal
bovine serum.
Preparation of Cell Lysates and Immunoprecipitation--
The
details of treating the cells with dexamethasone, preparation of cell
lysates, and immunoprecipitation of C subunits have been described
previously (23). For experiments quantitating PME-1 binding to
different mutants (Fig. 6B), immunoprecipitates were washed
twice with Nonidet P-40 lysis buffer, twice with phosphate-buffered saline, and once with ddH20. Washed immune complexes were used for
phosphatase assays or analyzed by one or two-dimensional gel electrophoresis.
One- and Two-dimensional Gel Electrophoresis and
Fluorography--
SDS-polyacrylamide gel electrophoresis (10%
acrylamide) was performed according to Laemmli (29). Two-dimensional
gel analysis was performed as described previously (30). Gels were
silver stained by the procedure of Wray et al. (31) except
that after electrophoresis the gels were sequentially incubated 10 min
in distilled water (200 ml), 10 min in 95% ethanol (200 ml), 1 h in 50% methanol (100 ml), and 30 min in distilled water (100 ml) prior
to staining.
Immunoblotting--
Immunoblotting (32) was performed with mouse
monoclonal anti-tag antibody (16B12; 1:5000 dilution of ascites;
BAbCO), rabbit anti-B subunit antibody (P16; 1:5000), affinity-purified
rabbit (R39; 1:5000) or mouse monoclonal (4G7; 1 µg/ml) anti-A
subunit antibodies, mouse monoclonal anti-MT antibody (F4; 0.25 µg/ml) (33), mouse monoclonal anti-C subunit antibody (1D6; 0.25 µg/ml), or rabbit anti-PME-1 antibodies (AR2 or E37; see below).
Immunoblots were developed with enhanced chemiluminescence (Amersham or
NEN Life Science Products Inc.).
Phosphatase Assay--
Phosphatase activity present in anti-HA
tag immunoprecipitates from the different cell lines was assayed using
phosphorylase a and histone H1. Purification and Microsequencing of p44A(PME-1)--
To obtain
p44A(PME-1) protein for microsequencing, H59Q C subunit complexes
containing p44A(PME-1) were immunoaffinity purified. In total, 135 confluent 15-cm dishes of MT-transformed NIH3T3 cells expressing
HA-tagged H59Q were used. Forty-five separate immunoaffinity
purifications were performed on 3 dishes of lysate at a time, reusing
the same immunoaffinity matrix at least 15 times. To prepare the
immunoaffinity matrix, anti-HA tag antibody (12CA5; obtained from
BAbCO) was chemically cross-linked to protein A-Sepharose beads
(Pharmacia) by published methods (35). Cell lysates were incubated for
1 h at 4 °C while rocking with 500 µl of the cross-linked
antibody-bead complexes. Complexes were washed once with Nonidet P-40
lysis buffer, three times with Tris-buffered saline, and then twice
with distilled deionized H2O. Bound H59Q complexes
containing p44A(PME-1) were eluted by three sequential incubations with
300 µl of 20 mM triethylamine. Eluates were quickly frozen on dry ice and stored frozen until all batches of affinity purification had been completed. The antibody-bead complexes were then
washed twice with 20 mM triethylamine and twice with
Nonidet P-40 lysis buffer prior to reuse. After H59Q complexes had been purified from all 135 dishes of cells, eluates containing p44A(PME-1) were concentrated to dryness by vacuum centrifugation, and the residues
were suspended in phosphate-buffered saline and gel buffer and analyzed
on three separate SDS-polyacrylamide gels. One-dimensional gels were
chosen to avoid losses associated with two-dimensional gel analysis.
Because p44A(PME-1) migrates closely to actin, the separation of these
two proteins was maximized by the use of an 8% gel run for an extended
period of time.
Trypsin Digestion, HPLC Separation, and
Microsequencing--
After separation of p44A(PME-1) complexes by
SDS-PAGE, the proteins were electrotransferred to polyvinylidiene
difluoride membrane and stained with Ponceau S. Individual protein
bands were excised and subjected to in situ digestion with
trypsin (36, 37). The resulting peptide mixture was separated by
microbore high performance liquid chromatography using a Zorbax C18
2.1 × 150-mm reverse phase column on a Hewlett-Packard 1090 HPLC/1040 diode array detector. Optimum fractions from the chromatogram were chosen based on differential UV absorbance at 205, 277, and 292 nm, peak symmetry and resolution. Peaks were further screened for
length and homogeneity by matrix-assisted laser desorption time-of-flight mass spectrometry on a Finnigan Lasermat 2000 (Hemel United Kingdom), and selected fractions were submitted to automated Edman degradation on an Applied Biosystems 494A, 477A (Foster City, CA)
or Hewlett-Packard G1005A (Palo Alto, CA). Details of strategies for
the selection of peptide fractions and their microsequencing have been
previously described (37).
cDNA Cloning via PCR--
To obtain the missing 5' portion
of the PME-1 coding region, nested and semi-nested PCR were performed
using human B cell, human hippocampus, and human kidney plasmid
libraries. 5' primers corresponded to vector sequence that flanked
cDNA inserts in the library being used as template, while 3'
primers corresponded to known sequence (EST or newly derived 5' PME-1
sequence). Southern blotting using an end-labeled 20-base pair
oligonucleotide corresponding to known PME-1 sequence upstream of the
3' PCR primer was employed to identify authentic PME-1 products after
each reaction. PCR products containing 5' extensions of the PME-1
sequence were purified using a PCR product purification kit (Roche
Molecular Biochemicals), cloned, and sequenced. New primers were
designed for PCR and Southern blotting and then the above steps were
repeated until the sequence of the remainder of the PME-1 coding region
and a portion of the 5' UTR were obtained.
RT-PCR--
Total mRNA was purified from HeLa cells using
Trizol Reagent (Life Technologies) according to the manufacturer's
instructions. First strand synthesis was performed with avian
myeloblastosis virus reverse transcriptase (Roche Molecular
Biochemicals) by the manufacturer's protocol using a primer from the
PME-1 3' UTR (TGTTGAGGAGGGGTGGACAG). Using Pfu polymerase (Stratagene),
the product was used for PCR with the same 3' primer and a primer from
the PME-1 5' UTR (TGTATGGGGACCTTCCTCCT) to generate a cDNA containing the entire PME-1 coding region and much of the 5' UTR.
Computer Analyses--
The NCBI BLAST program (38) was used to
probe various data bases for p44A(PME-1) ESTs and related proteins. The
DNASTAR Lasergene software package was utilized for alignments and
identification of the PROSITE data base lipase motif found in
p44A(PME-1).
Northern Blot--
Adult Balb/c mice were sacrificed and organs
removed and flash-frozen in liquid nitrogen. Total RNA from the organs
was isolated using the RNeasy kit (QIAGEN), and analyzed on
formaldehyde, 1% agarose gels to check for RNA integrity and to
estimate the amount of the 18 S and 28 S RNAs. Based on these
estimates, similar amounts of RNA were separated on formaldehyde, 1%
agarose gels and transferred to GeneScreen nylon membranes. After UV
cross-linking, the membranes were stained with a 0.04% methylene blue
solution to visualize the RNA. Filters were then hybridized with a
32P-radiolabeled probe generated by random primer labeling
of a DNA fragment from the 3'-untranslated region of the mouse PME-1 cDNA. The probe, 395 base pairs in length, is an
EcoRI-NotI fragment of a PME-1 EST clone
(accession number W34856). The blots were used for autoradiography with
x-ray film and/or analyzed on a STORM PhosphorImager (Molecular Dynamics).
Production of Polyclonal Antibodies Recognizing PME-1--
Two
different antisera recognizing PME-1 were raised in rabbits. The first,
AR2, was raised against a 16-residue PME-1 peptide sequence
(RIELAKTEKYWDGWFR) found encoded in the PME-1 cDNA. The peptide was
conjugated to keyhole limpet hemocyanin via an added carboxyl-terminal
cysteine residue using a Pierce Imject conjugation kit, and the
conjugate was used as immunogen. The second antiserum, E37, was raised
against a mixture of two nickel agarose-purified, 6xHis-tagged,
bacterially expressed human PME-1 fragments that together represent the
carboxyl-terminal half of the protein. For each immunogen, a single
female New Zealand White rabbit was immunized and boosted multiple
times using Freund's adjuvant.
Demethylation Assays--
Assays utilizing methylation-sensitive
antibodies (6) were performed to evaluate PP2A demethylation.
Logarithmically growing wt C subunit-expressing MT-transformed NIH3T3
cells were lysed and C subunit immunoprecipitated as described
previously (23). The C subunit immunoprecipitate was divided into equal
aliquots for use as substrate in demethylation reactions. C subunit
immune complexes from one 10-cm dish of cells could support 8 demethylation reactions. To each aliquot of substrate, 38.75 µl of
reaction buffer containing 55 mM Tris, pH 8.0, 55 mM NaCl, 1 mM dithiothreitol, 1.0 mM CaCl2, 1.0 mM MgCl2,
0.55% Nonidet P-40, and 0.2 mg/ml bovine serum albumin was added.
Then, 0.5 µl of inhibitor (okadaic acid or PMSF) dissolved in
dimethyl sulfoxide or dimethyl sulfoxide without inhibitor was added to
the appropriate tubes. After 3 min, 0.75 µl of lysate from bacteria
expressing or not expressing PME-1 was added to the appropriate tubes
(to obtain bacterial lysate, bacteria were lysed by sonication in 25 mM Tris, pH 8.0, containing 140 mM NaCl, and
lysates were cleared by centrifugation at 13,000 × g
for 5 min). Demethylation reactions were carried out at 32 °C with
shaking for 60 min. Then the reactions were combined with SDS-PAGE
sample buffer and boiled. Following SDS-PAGE and electrophoretic
transfer of proteins to nitrocellulose, the membrane was immunoblotted
as described in the legend to Fig. 5.
Generation and Characterization of Catalytically Inactive PP2A C
Subunits--
In order to create catalytically inactive PP2A C subunit
mutants that retained the maximum amount of structural integrity, single residues likely to be important for catalysis were mutated. To
identify such residues, an alignment of PP2A and various related phosphatases was performed to identify highly conserved residues (data
not shown). A small number of residues were found that are identical in
PP2A, PP1, PPX, PP2B, and PP
After construction of stable lines, the C subunit mutants were
characterized with respect to two properties: 1) competence to form
complexes containing the A and B subunits or MT and 2) catalytic
activity. To examine complex formation in vivo,
immunoprecipitates of epitope-tagged wt and mutant C subunits were
probed by immunoblotting for the presence of additional subunits and MT
(Fig. 1). Both mutants bind substantial A
subunit. H118Q also binds a small amount of B subunit, while H59Q binds
almost none of this subunit. Although a small amount of MT was found in
control immunoprecipitates from cells which do not express any
epitope-tagged C subunit, levels of MT well above this were readily
detected in the mutant immunoprecipitates, indicating that A·C·MT
trimeric complexes had been formed by these proteins. A portion of the
MT coimmunoprecipitated with H59Q is shifted relative to the MT
associated with wt C subunit; this result is reproducible and will be
described in more detail
elsewhere.4 The fact that
both mutants bind additional subunits indicates that these mutants have
substantial native structure in vivo.
To test for catalytic activity, phosphatase assays were performed on
anti-tag immunoprecipitates from the various cell lines. Using both
phosphorylase a and histone H1 as substrates, only wt C
subunit immunoprecipitates were found to have increased activity compared with control immunoprecipitates prepared from a cell line
containing only empty vector (Table I).
Immunoprecipitates of the two mutants showed no activity above the
control level toward either substrate. This finding is consistent with
the results of others who found that mutation of the corresponding
residues in related phosphatases also completely inactivated those
enzymes (41-43).
Two 44-kDa Cellular Proteins Associate with the Inactive C-subunit
Mutants, H59Q and H118Q--
To determine if novel cellular proteins
associate with one or both catalytically inactive C subunit mutants,
anti-tag immunoprecipitates were prepared from 35S-labeled
cells and analyzed on two-dimensional gels. Several proteins were seen,
including one prominently labeled protein of 44 kDa with a pI near 7 (p44B), that specifically associate with H59Q and H118Q but not with
control immunoprecipitates prepared from cells containing empty vector
or cells expressing wt C subunit (data not shown).
To determine if sufficient p44B could be obtained to allow
microsequencing, scaled up immunoprecipitates from vector only control
cells (GREonly) and from cells expressing H118Q were analyzed on
two-dimensional gels and silver-stained (Fig.
2). P44B was not readily visible in these
gels (see brackets); however, another 44-kDa protein was seen that also
specifically coimmunoprecipitates with H118Q. This protein was present
in almost a 1:1 stoichiometry with the A and C subunits and was
initially designated p44A because its pI, approximately 6, was more
acidic than that of p44B. A similar p44A spot was found in
silver-stained immunoprecipitates of H59Q (data not shown). An
35S-labeled spot corresponding to p44A was absent in the
two-dimensional gels of 35S-labeled immunoprecipitates
described above, suggesting that p44A may be synthesized at a very low
rate and may have a much longer half-life than the PP2A C or A subunits
or p44B. Alternatively, there may be a delay before newly synthesized
p44A forms a complex with the inactive mutants.
Purification, Microsequencing, and cDNA Cloning of p44A Reveals
That It Is a Novel Protein (PME-1) Highly Conserved from Yeast to
Human--
To facilitate the identification or cloning of p44A,
sufficient protein for microsequencing was obtained by purifying
HA-tagged H59Q complexes on an anti-tag immunoaffinity column as
described under "Experimental Procedures." After analysis on
one-dimensional gels, both actin and a clearly separated p44A band
migrating just above it were excised for further processing.
Microsequencing of two tryptic peptides from the lower band confirmed
that it was indeed actin (data not shown). Nine microsequences obtained from the p44A band matched no known protein in GenBank, indicating that
it was a novel protein.
Initial searches of the expressed sequence tag (EST) data base also
revealed no match for the microsequences from p44A; however, in time, a
human EST sequence (accession number H12112) was deposited that encoded
three of those microsequences. Because a human EST sequence matched
first, we decided to clone the human version of this protein, which we
have renamed PME-1 for functional reasons that will become obvious
below. Additional DNA sequencing of EST H12112 revealed coding
information for two more PME-1 microsequences, and it was determined
that this EST encoded most of the carboxyl-terminal half of PME-1 (162 amino acids).
To obtain the missing 5' portion of the PME-1 coding region, nested and
semi-nested PCR were performed as described under "Experimental
Procedures." The sequence of the remainder of the coding region and a
portion of the 5'-UTR were obtained. Because errors may have occurred
during the multiple PCR reactions that were necessary to obtain the
complete cDNA sequence, RT-PCR was then performed with HeLa cell
mRNA as template to provide a reliable PME-1 coding sequence.
Sequencing of this cDNA resolved a few questionable nucleotides in
the coding region, and confirmed the presence of a stop codon just
upstream of the predicted start codon (Fig.
3A). The stop codon upstream
of the predicted start ATG is in-frame with the PME-1 coding sequence,
indicating that the PME-1 cDNA contains the complete PME-1 ORF.
A schematic of a PME-1 cDNA that includes the end of the 3'-UTR
deduced from overlapping ESTs is shown in Fig. 3A. The
entire sequence is approximately 2.5 kilobase pairs in size, with an open reading frame (ORF) of 1164 base pairs including two tandem stop
codons. When a probe specific for mouse PME-1 was used to detect
transcripts from different mouse organs, a single transcript of
2.6 ± 0.2 kilobase pairs was detected in all tissues (Fig. 3B). The level of this transcript was highest in brain and
testis. To date, multiple EST sequences which encode PME-1 have been
deposited in the DBEST data base. The sequenced portions of those ESTs
cover the entire 3'- and 5'-UTRs, but not the entire coding region. Information from the NCBI Cancer Genome Anatomy Project (CGAP) indicates that PME-1 ESTs have been mapped to human chromosome 11, interval D11S916-D11S911 (80-84 centimorgan). It is not known at this
time whether PME-1 is mutated in any of the diseases with defects
mapped to this general region of chromosome 11.
The 386-amino acid PME-1 (p44A) protein product encoded by the human
PME-1 cDNA ORF is shown in Fig. 3C. It has a predicted molecular size of just over 42 kDa, close to that predicted from its
migration on SDS-PAGE. In addition, it has a pI of 5.8, consistent with
its migration in the isoelectric focusing dimension on two-dimensional gels like the one shown in Fig. 2. All nine mouse PME-1 tryptic peptide
sequences (underlined in Fig. 3C) were accounted
for in the human sequence with differences present only at a few
positions, indicating that PME-1 is well conserved between these two
species. Because no tryptic peptide microsequences potentially
corresponding to p44B were found, efforts to identify this protein are
still ongoing.
Using the NCBI BLAST program, highly homologous sequences probably
corresponding to PME-1 homologs were found for zebrafish, C. elegans, and S. cerevisiae. A hypothetical 88.4-kDa
C. elegans protein in chromosome 3, B0464.7, contains some
of the C. elegans sequence homologous to PME-1, but lacks
other highly homologous sequences, suggesting that it may not represent
an accurate prediction of exon combinations. A more likely combination
of exons that includes all B0464 cosmid exons homologous to PME-1
generates a protein of 364 amino acids and approximately 40 kDa (Fig.
3D). S. cerevisiae PME-1 (Fig. 3D)
appears to be a single hypothetical 44.9-kDa protein (PIR accession
number S46814) of unknown function encoded by an ORF on chromosome 8R
(YHN5; GenBank accession number U10556). Recently, based on a single
partially homologous nonapeptide sequence, YHN5 was proposed to be a
mitochondrial ribosome small subunit protein and named YmS2 (44). Human
PME-1 has approximately 40 and 26%, respectively, identity with the
C. elegans and yeast sequences (Fig. 3D). A
highly charged stretch of amino acids is present in human PME-1 but
absent in PME-1s from C. elegans and S. cerevisiae. This stretch of amino acids does not represent a
cloning artifact, because 35S-labeled PME-1 in
vitro transcription/translation product generated using the human
PME-1 cDNA comigrates on gels with PME-1 from HeLa cell lysates
(data not shown).
In order to facilitate further experiments characterizing PME-1, an
anti-PME-1 peptide antibody was raised to a 16-amino acid peptide
sequence encoded by the PME-1 cDNA. This peptide antibody detected
a 44-kDa protein present in H59Q immunoprecipitates, but absent from
immunoprecipitates of wild-type C subunit (Fig. 4). Thus, PME-1, like p44B, associates
stably with the catalytically inactive mutant C subunits, but not with
wt C subunit. Because B subunit, but not MT, requires the C subunit
carboxyl terminus for association with the PP2A A/C heterodimer, we
wanted to determine if MT expression might increase the amount of PME-1
bound to H59Q. Similar levels of PME-1 were coimmunoprecipitated from
untransformed NIH3T3 cells and polyomavirus MT-transformed NIH3T3 cells
(Fig. 4), indicating that MT expression does not greatly affect the level of H59Q·PME-1 complex formation in the cell.
PME-1 Is a PMSF-resistant, Okadaic Acid-sensitive PP2A
Methylesterase That Probably Uses Serine as an Active Site
Nucleophile--
When the human, C. elegans, and S. cerevisiae PME-1 protein sequences were analyzed for motifs found
in the Prosite data base using DNASTAR Lasergene software, a consensus
sequence
((L/I/V)X(L/I/V/F/Y)(L/I/V/S/T)G(H/Y/W/V)S-XG(G/S/T/A/C)) for lipases utilizing an active site serine was found to be conserved. The invariant serine in this motif, corresponding to serine 156 in
human PME-1, is the active site serine in these enzymes. In addition,
scattered similarities can be seen between other regions of the PME-1
sequence and some of the lipases that have this motif. Therefore, PME-1
is probably a lipase whose active site serine is serine 156.
The various lipases that share this motif are found in both prokaryotes
and eukaryotes and include, among others, two Drosophila melanogaster carboxylesterases. In addition, CheB, a bacterial glutamate methylesterase, has a similar, but not identical, sequence surrounding its active site serine (45) (Table
II). CheB (46) and other lipases
utilizing an active site serine (e.g. Refs. 47 and 48) have
a catalytic triad in their primary sequence in the order Ser-Asp(or
Glu)-His. Of the conserved histidines in human PME-1, His-349 is a
likely candidate for a putative catalytic triad histidine (Fig.
3D). Identification of a putative PME-1 catalytic triad
acidic residue by sequence comparison is more problematic because there
are multiple acidic residues conserved between species. However, of
these, two aspartates in human PME-1, Asp-181 and Asp-324, show
conservation in position with putative catalytic triad aspartates in
other lipases, and therefore may be more likely possibilities.
A PP2A C subunit carboxyl methylesterase of 46 kDa has recently been
purified (24), but no sequence information was reported. To test the
possibility that PME-1 might be a PP2A methylesterase, PME-1 was
expressed in bacteria and bacterial lysates were tested for
methylesterase activity toward PP2A C subunit as described under
"Experimental Procedures." The results shown in Fig.
5 demonstrate that lysates of bacteria
expressing PME-1 contain a PP2A methylesterase activity not found in
bacterial lysates lacking PME-1. Similar results were obtained with
purified recombinant PME-1 (Fig. 5). These results indicate that PME-1
is indeed a PP2A methylesterase. Because its specificity toward other
methylated phosphatases (such as PPX) has not been characterized, it
was generically named protein phosphatase
methylesterase-1 (PME-1).
The 46-kDa PP2A methylesterase reported by Lee and co-workers (24) was
inhibited by okadaic acid, a potent PP2A inhibitor, but not by PMSF, a
covalent inhibitor of certain serine esterases. To determine if PME-1
displays similar sensitivities to these inhibitors, the above
demethylation assay was also conducted in the presence of okadaic acid
and PMSF (Fig. 5). The methylesterase activity of bacterially expressed
PME-1 was inhibited by 0.1 or 1 µM okadaic acid but not
by 1 or 5 mM PMSF, similar to the methylesterase purified
by Lee et al. (24).
Okadaic Acid and Other PP2A Inhibitors Decrease the Association of
PME-1 with H59Q--
Because single amino acid changes in the C
subunit active site were capable of inducing stable complex formation
of C subunit with PME-1, it was of interest to determine if PP2A
inhibitors could antagonize the H59Q·PME-1 complex. To assay for this
possibility, NIH3T3 cells expressing epitope-tagged H59Q C subunit were
lysed in the presence of various phosphatase inhibitors and H59Q was immunoprecipitated via its epitope tag. The amount of endogenous, untagged PME-1 coimmunoprecipitating in each case was assayed by
blotting with anti-PME-1 antibody (Fig.
6A). Inhibitors to which PP2A
is highly sensitive (okadaic acid, sodium fluoride, and sodium
pyrophosphate), but not those to which PP2A is less sensitive or
insensitive (vanadate and phenylarsine oxide, respectively), decreased
the amount of PME-1 bound to H59Q.
The H59Q Carboxyl Terminus Is Important, but Not Essential, for
Complex Formation with PME-1--
A PP2A methylesterase might be
expected to make important contacts with carboxyl-terminal residues.
However, Lee and co-workers (24) found that PP2A carboxyl-terminal
peptides functioned neither as inhibitors nor as substrates for their
46-kDa PP2A methylesterase, suggesting that, at a minimum, contacts
with other parts of the C subunit are essential. To investigate the
importance of the H59Q C subunit carboxyl terminus for stable
interaction with PME-1, a double mutant, H59Q/301Stop, was created.
This mutant combines the H59Q mutation, which induces stable binding of
PME-1, with a deletion of the nine C subunit carboxyl-terminal acids,
301-309. Fig. 6B, shows the results of an
immunoprecipitation assay measuring the relative abilities of H59Q and
H59Q/301Stop to bind A subunit and PME-1. Deletion of residues 301-309
from wt C subunit has previously been found to decrease the amount of A
subunit bound (23). Fig. 6B shows that deletion of these
residues from H59Q also reduces the binding of the PP2A A subunit to
H59Q. In addition, although similar amounts of H59Q and H59Q/301Stop
were immunoprecipitated in this experiment, the double mutant bound
less PME-1 than did H59Q, indicating that one or more of the deleted
carboxyl-terminal residues is important for H59Q·PME-1 complex
formation. PME-1 binding was not completely abolished, however,
demonstrating that interactions also exist between PME-1 and other
residues in the C subunit.
To address the same question via a different approach, we assayed via
immunoprecipitation whether antibodies directed against the C subunit
carboxyl terminus would compete with PME-1 for binding to H59Q. If an
antibody competes with PME-1 for binding to residues on H59Q that are
important for PME-1 association, that antibody would be expected to
coimmunoprecipitate reduced amounts of PME-1 with H59Q when compared
with an antibody that does not compete with PME-1. The
carboxyl-terminal C subunit monoclonal antibodies used for this
experiment, 1D6, 4B7, and 4E1, were recently generated against a
15-residue unmethylated carboxyl-terminal peptide.3 These
antibodies are unable to efficiently recognize a C subunit mutant
lacking the carboxyl-terminal leucine, indicating that they bind, at
least in part, at the very carboxyl
terminus.5 A positive control
monoclonal antibody, 12CA5, immunoprecipitates H59Q via its
amino-terminal epitope tag and should not interfere with interactions
at the C subunit carboxyl terminus (23). Comparison of the relative
ratios of the PME-1 and H59Q bands in Fig. 6C reveals that,
relative to 12CA5, 1D6 and 4B7 immunoprecipitate less PME-1 for the
same amount of H59Q C subunit (the band of endogenous, wt C subunit
immunoprecipitated by the carboxyl-terminal antibodies can be ignored
as wt C subunit does not associate stably with PME-1). Furthermore,
although 4E1 immunoprecipitated a substantial amount of H59Q C subunit
(within approximately 2-fold of 12CA5), no PME-1 could be detected even
on long exposures. These results thus further substantiate the
conclusions made from Fig. 6B. In addition, the fact that
1D6 and 4B7 coimmunoprecipitate similar amounts of PME-1, but
dramatically different amounts of A subunit indicates that PME-1
binding does not appear to be dependent on A subunit binding.
In this study, we report the identification of the first of a
number of cellular proteins that stably associate with catalytically inactive PP2A C subunit mutants, but not with wt C subunit. Two proteins of 44 kDa that differ in their isoelectric points, p44A and
p44B, uniquely associated with two different catalytically inactive C
subunit mutants substituted individually at two different active site
histidine residues. P44A was affinity purified and a cDNA encoding
it was cloned. This protein was identified as a PP2A methylesterase
(PME-1) by several criteria including: 1) molecular size, 2) the
presence of a motif found in lipases that use serine as their
nucleophilic catalytic residue, 3) activity assays performed in
vitro with bacterially expressed protein, and 4) the ability of
okadaic acid, a known inhibitor of both PP2A and the PP2A
methylesterase, to inhibit its activity and decrease its association
with the catalytically inactive C subunit mutant, H59Q.
The cloning of PME-1, to our knowledge, provides the first sequence of
a mammalian protein methylesterase. Like the specific bacterial
methylesterase CheB (49, 50), which participates in the chemotactic
response, PME-1 appears to be a serine esterase which is resistant to
PMSF. The molecular basis for resistance to PMSF is presently unknown,
but it has been proposed that a unique arrangement of the active site
residues in CheB might contribute (46). It will be interesting to see
if PME-1 has a similar arrangement of its catalytic residues. It seems
likely that PME-1, like CheB (46) and other lipases utilizing an active
site serine (e.g. Refs. 47 and 48), will be found to have a
catalytic triad of serine, histidine, and aspartate (or glutamate) in
its active site. We are presently attempting to obtain structural
information for PME-1 to determine which residues are in the active
site and to gain insight into the molecular basis of PME-1 recognition of PP2A C subunit and into potential regulation of this methylesterase.
Based on its molecular size, sensitivity to okadaic acid, and the lack
of effect of PMSF on PME-1 activity, PME-1 is likely to be equivalent
to the 46-kDa PP2A methylesterase whose purification and initial
characterization was recently reported by Lee and colleagues (24). Its
insensitivity to PMSF indicates that it is not the PMSF-sensitive
serine esterase/protease activity reported by Xie and Clarke (51),
which also could remove PP2A carboxymethyl groups. Lee and co-workers
(24) reported that their purified PP2A methylesterase eluted as two
different peaks from an anion exchange column, consistent with either
differential modification or the existence of two closely related
isoforms of the enzyme. The amounts of these two species were within
severalfold of each other. Two pieces of evidence from our studies
support the idea that those two forms probably represent differentially
modified forms of the enzyme. First, probing of the GenBank EST data
base with the PME-1 cDNA sequence provides no evidence for a
closely related PME-1 isoform, even though numerous ESTs are found
which correspond precisely to the PME-1 cDNA sequence. Second,
Northern blot analysis yielded a single band in multiple organs. In
addition, we have found via immunoblotting that mammalian PME-1 in cell lysates migrates on two-dimensional gels as two spots differing in
their isoelectric point in a manner consistent with a single charge
difference.6
The molecular basis of the cell cycle-dependent regulation
of PP2A C subunit methylation is unknown. The poor metabolic labeling of PME-1 in an asynchronous population of cells relative to a number of other proteins suggests that this protein is quite stable. This result argues against the possibility that cell cycle PP2A methylation is regulated by modulating the amount of the PP2A methylesterase. Whether PME-1 activity is regulated is unknown. In the
case of the bacterial chemotactic response, the CheB methylesterase is
regulated by phosphorylation (52, 53) while the methyltransferase is
thought to be constitutively active. Lee and co-workers (24) found no
difference in the activity of their two purified forms of PP2A
methylesterase, suggesting that the differential modification likely
responsible for generating these two forms might not be involved in
regulation of activity of this enzyme. It is possible, however, that
effects might be seen under other conditions, or that an additional
protein(s) may be necessary for an effect to be manifested. In
addition, it is possible that more than one modification occurs. More
definitive evidence should be obtained from a genetic analysis of the
importance of the site(s) of modification, once identified.
The intriguing possibility exists that PP2A methyltransferase and
methylesterase enzymes might achieve their specificity in part by
interacting with or near the active site of the PP2A C subunit. This
hypothesis is consistent with a number of experimental findings. First,
it was reported previously that neither the PP2A methyltransferase nor
the PP2A methylesterase can recognize short peptide substrates
corresponding to the C subunit carboxyl terminus. Thus, functional
recognition by both these enzymes requires additional C subunit
structure. Second, as demonstrated in this study, perturbation of the C
subunit active site by either of two different mutations can stabilize
the interaction with the PME-1 methylesterase. Third, PP2A inhibitors
have a destabilizing effect on the PME-1-H59Q interaction. Fourth, the
methyltransferase is inhibited by the PP2A inhibitors, okadaic acid and
microcystin-LR, and the methylesterase is inhibited by okadaic acid
(testing for inhibition of the methylesterase by microcystin has not
been reported). Although it has been proposed that this inhibition may
be due to the interaction of these inhibitors with carboxyl-terminal C
subunit residues, this would not explain the ability of the PP2A
inhibitors, sodium fluoride or sodium pyrophosphate, to partially or
fully disrupt PME-1·H59Q complexes. The latter effect is more
consistent with a role in binding the PME-1 methylesterase for active
site residues and/or metals, or nearby residues sensitive to effects on
the active site. Yet another experimental finding compatible with the
above hypothesis is that four separate catalytically inactive PP2A
active site point mutants, including the two described in this study,
are methylated at less than 3% of the wild-type level in
vivo and in vitro.3 Although all these
findings are consistent with our hypothesis of an interaction with
residues and/or metals in or near the active site, another equally
viable possibility is that mutation of active site residues and/or
binding of inhibitors may have more distant effects on the C subunit
conformation critical for stable complex formation with PME-1.
Contact between the C subunit and PME-1 could theoretically be with
PME-1 residues and/or with a phosphorylation site on PME-1. Because
H59Q and H118Q are virtually unmethylated,3
PME-1 apparently can remain bound to these mutants in the absence of a
methylated carboxyl terminus. At least with H59Q, PME-1's contacts
other than on the C subunit carboxyl terminus are strong enough to
result in substantial complex formation in the absence of the nine
carboxyl-terminal C subunit residues. This conclusion is further
supported by the finding that two C subunit carboxyl-terminal peptide
antibodies, known to require Leu-309 for efficient binding, could
immunoprecipitate H59Q·PME-1 complexes. However, the amount of PME-1
coimmunoprecipitated by these antibodies was less than that
coimmunoprecipitated by an antibody recognizing an amino-terminal epitope tag on the C subunit. The latter result and the fact that a
third carboxyl-terminal C subunit antibody could not immunoprecipitate H59Q·PME-1 complexes at all suggest that PME-1 is proximal to the C
subunit carboxyl terminus in the H59Q·PME-1 complex. Moreover, the
reduced amounts of PME-1 in complex with the H59Q·301Stop double
mutant indicate that carboxyl-terminal residues play a role in binding
of H59Q to PME-1. The contribution of these residues to the interaction
of wild type C subunit with PME-1 might be even more important in the
absence of the complex-stabilizing H59Q mutation.
The significance of the decreased B subunit binding observed with these
mutants is unclear, but an attractive hypothesis is that it might be
due indirectly to lack of methylation at the carboxyl terminus of these
mutants. The fact that H59Q and H118Q bind the structural PP2A A
subunit and polyomavirus MT suggests that they are not grossly altered
in their structure. Two other catalytically inactive point mutants that
bind A subunit and polyomavirus MT, but are highly deficient in
methylation,3 are also deficient in B subunit
binding.4 Given that the B subunit requires the C subunit
carboxyl terminus for stable complex formation with the A/C
heterodimer, the B subunit might require a methylated carboxyl terminus
for efficient binding to C subunit. An alternate, but not mutually
exclusive, possibility is that the carboxyl terminus and the active
site are proximal in the three-dimensional structure of the C subunit.
This model would provide an explanation for how events occurring at the
carboxyl terminus (B subunit binding, methylation, phosphorylation,
etc.) can affect the active site (activity, specificity), and vice
versa. In addition, at least for H59Q and H118Q, PME-1 and B subunit binding might be mutually exclusive, although this remains to be tested.
These catalytically inactive C subunit mutants should be useful for
identifying other proteins involved in PP2A signaling. H59Q and H118Q
bind multiple proteins not bound stably by wt C subunit. These include,
in addition to PME-1, p44B, and several other proteins. Interestingly,
initial experiments suggest that p44B binding to H59Q is even more
sensitive to phosphatase inhibitors than is PME-1
binding.7 These proteins
could be PP2A substrates or other proteins whose binding is sensitive
to the state of the C subunit active site. One of these proteins is the
same molecular size as the PP2A methyltransferase reported by Lee and
Stock (19). Catalytically inactive mutants of dual specificity and
tyrosine phosphatases (54, 55) have been previously used successfully
to identify novel substrates, but unlike PP2A, their catalytic
mechanisms involve the formation of covalent intermediates with
substrates. It will be very interesting to determine whether any of the
remaining H59Q/H118Q associated proteins are indeed PP2A substrates.
PME-1 and p44B differ in several characteristics, suggesting that these
two proteins are not simply modified forms of one another. They are
separated from each other on two-dimensional gels by approximately 1 pH
unit, which is unlikely to be accounted for by modification; PME-1
forms sharp spots on these gels while p44B migrates as a smear. In
addition, in vitro translation of PME-1 yields no product
migrating at the position of p44B and we have been unable to detect
p44B with antibodies raised against PME-1
sequences.6
Finally, because of the high conservation of PP2A with other
phosphatases such as PP1, PPX, PPV, etc., it will be of interest to see
if similar or different cellular proteins bind stably to these
phosphatases when the residues corresponding to PP2A His-59 and His-118
are mutated to glutamine. One question of special interest is whether
the corresponding catalytically inactive mutants of PPX,
which has the same last four carboxyl-terminal amino acids as PP2A and
is also methylated at its carboxyl-terminal leucine, will trap its
methylesterase. Given the lack of close relatives to PME-1 in the
various data bases, it would not be surprising if that methylesterase
turns out to be PME-1.
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
-32P-Labeled
phosphorylase a substrate was prepared from phosphorylase b according to the manufacturer's (Life Technologies, Inc.)
instructions. Histone H1 was phosphorylated by mitotic p34cdc2
purified from Nocodazole arrested HeLa cells as described previously (34). Lysates used for immunoprecipitation were equilibrated according
to HA-tagged C subunit expression levels. Assays were performed at a
linear range and with subsaturating amounts of each substrate.
RESULTS
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ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
. Of those, two histidines (H) at
positions 59 and 118 were chosen as having catalytic potential, and
were individually mutated to glutamine (Q), yielding the mutants H59Q
and H118Q. Subsequent to the construction of these mutants, the crystal
structures of PP1 and PP2B (39, 40) and a mutational analysis of PP
(41) were reported, the results of which suggested that these two
histidines would be involved in PP2A substrate binding and catalysis.
As described under "Experimental Procedures," each C subunit mutant
or wt cDNA was constructed with the hemagluttinin (HA) tag at its
amino terminus to allow for immunoprecipitation analysis (23).
Individual mutants, wild-type C subunit, or vector only were expressed
stably in NIH3T3 cell lines with and without coexpression of MT. In the
MT expressing cells, most PP2A complexes still contain B subunit
because MT is produced at a low level relative to endogenous PP2A
(8).2
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Fig. 1.
The catalytically inactive mutant PP2A C
subunits can form complexes with A subunit and MT in
vivo. Lysates from cells containing only control vector
(GREonly) or HA-tagged wt (wt C sub) or mutant C
subunits (H59Q and H118Q) were immunoprecipitated
with anti-HA tag antibody (12CA5) and analyzed by SDS-PAGE and
immunoblotting. The blot was probed first with anti-MT antibody, and
then sequentially with antibodies recognizing the A, C (via the HA
tag), and B PP2A subunits. Because a lower level of expression was
consistently seen with H118Q, the immunoprecipitate of this mutant was
prepared from more cells; to properly control for this, the control
immunoprecipitate was prepared from an equivalent amount of cells
expressing only the vector. Under these conditions, a small amount of
MT can be seen sticking nonspecifically to the immunoprecipitate in the
GREonly lane.
H59Q and H118Q are catalytically inactive
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Fig. 2.
p44A appears to bind stoichiometrically to
H118Q. Silver-stained two-dimensional gels of HA tag
immunoprecipitates prepared from unlabeled cells expressing vector only
(GREonly) or the C subunit mutant, H118Q, are shown. Only
the portion of each gel containing the relevant proteins is shown. The
A and C subunits, p44A, and anti-HA tag antibody heavy chain
(Ab) are indicated by labeled brackets and
arrowheads. Unlabeled arrowheads indicate the
corresponding positions in the GREonly control panel. For reference,
actin is indicated in both panels by a small unlabeled
arrow. The approximate position where p44B would be located on
these gels is indicated by the unlabeled brackets.
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Fig. 3.
PME-1 cDNA schematic, mRNA tissue
distribution, and predicted protein sequence. A,
schematic of a 2.5-kilobase human PME-1 cDNA. On the stick diagram,
the positions of the in-frame 5'-UTR stop codon (TGA), of the first two
potential start codons (ATGs), of tandem stop codons (TAGTGA) at the
end of the PME-1 ORF, and of the poly(A) tail (bracket) are
shown. The 3' end of the 3'-UTR, including the position of the poly(A)
tail, was deduced by analyzing overlapping PME-1 ESTs; all other
regions were directly sequenced. The sequence shown below the stick
diagram extends from the in-frame 5'-UTR stop codon (TGA;
overlined) to the second possible start ATG (double
underlined). The first possible start ATG (underlined
once in the sequence shown) was identified as the authentic start
site in vivo by making constructs whose
transcription/translation products in vitro would start with
one or the other of these two ATGs. 35S-Labeled in
vitro transcription/translation product starting at the first ATG,
but not the product starting at the second ATG, comigrated precisely on
two-dimensional gels with PME-1 from HeLa cell lysates (data not
shown). B, expression of PME-1 mRNA in different
tissues. Total RNA from the indicated mouse organs was separated by
electrophoresis and hybridized with a mouse PME-1 partial cDNA
probe from the 3'-UTR of mouse PME-1. In a separate experiment, the
size of the PME-1 transcript was calculated to be 2.6 ± 0.2 kilobases. The lower panel shows the 18 S rRNA from the same
blot visualized with methylene blue. C, the protein sequence
encoded by the human PME-1 cDNA is shown. Amino acid sequence
information for murine PME-1(p44A) obtained by tryptic peptide
microsequencing is underlined. Over 98% of the
microsequenced murine residues (107 of 109) were identical to the human
sequence. The double underlined serine at position 42 corresponds to a threonine in murine PME-1. D, alignment of
human, C. elegans (predicted), and S. cerevisiae
(YHN5) PME-1 protein sequences. Residues identical with human PME-1 are
shaded. Residues corresponding to the Prosite motif for
lipases employing an active site serine are boxed.
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Fig. 4.
PME-1 stably associates with H59Q but not
wild-type C subunit. HA tag immunoprecipitates prepared from
NIH3T3 (NIH) or MT-transformed NIH3T3 (NIHMT)
cell lines individually expressing HA-tagged wt (wt C sub) or mutant
(H59Q) C subunits were analyzed by SDS-PAGE and immunoblotting with HA
tag antibody and PME-1 anti-peptide antibody. The C subunits migrate as
tight doublets in these gels; whether doublets or a single band are
seen varies from gel to gel and does not appear to be due to
degradation (6, 13, 23). The panels and lanes shown are from the same
experiment and gel, but the lanes were not all originally adjacent.
Even on long exposure, the 44-kDa protein seen in the mutant lanes is
not seen in the wt lanes.
Comparison of the sequences surrounding the putative or known active
site serines of PME-1 proteins and CheB
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Fig. 5.
Human PME-1 is a PP2A methylesterase.
Immunoprecipitated PP2A C subunit was incubated with lysates from
bacteria either not expressing PME-1 (control) or expressing
PME-1 (PME-1), or with purified bacterially expressed PME-1
(~5 ng). Okadaic acid (O.A.) or PMSF was added to the
reactions to the indicated final concentrations. Reactions containing
1.25% dimethyl sulfoxide (DMSO) as a control to match the
level resulting from addition of okadaic acid or PMSF stock solutions
are noted. After incubation, the immunoprecipitated PP2A C subunits
were analyzed by SDS-PAGE. Proteins were transferred to nitrocellulose
and the membrane was probed with 4b7 (methylation-sensitive Ab), an
anti-C subunit antibody that only recognizes unmethylated C subunits.
Subsequently, the same membrane was probed with Transduction
Laboratories anti-PP2A C subunit antibody (methylation insensitive Ab),
which is insensitive to the methylation state of PP2A and therefore
reveals the total C subunit in each lane. The C subunits migrated as
doublets in this gel, but whether double or single bands are seen can
vary (see comments in legend to Fig. 4).
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Fig. 6.
Analysis of H59Q·PME-1 complex
formation. A, the PP2A inhibitors, okadaic acid, sodium
fluoride, and sodium pyrophosphate, reduce the amount of PME-1
complexed with the catalytically inactive H59Q C subunit. Seven
parallel dishes of NIH3T3 cells expressing HA-tagged H59Q were lysed as
described under "Experimental Procedures" in Nonidet P-40 lysis
buffer containing the indicated inhibitor(s) at the following
concentrations: sodium vanadate (1 mM), sodim fluoride (50 mM), okadaic acid (500 nM), phenylarsine oxide
(PAO, 10 µM), sodium pyrophosphate
(NaP~P, 20 mM). Anti-HA tag immunoprecipitates
were prepared from these lysates and analyzed by SDS-PAGE and
immunoblotting. The blot was probed sequentially with antibodies
detecting PME-1 and H59Q C subunit (via its HA tag). In a separate
experiment using phosphorylase a as substrate (not shown),
sodium fluoride, okadaic acid, and sodium pyrophosphate were,
respectively, found to inhibit PP2A 91 ± 10, 97 ± 4, and
>99%, while phenylarsine oxide and sodium vanadate, respectively,
showed no or 25 ± 18% inhibition. B, loss of the C
subunit carboxyl terminus reduces, but does not abolish, PME-1 binding.
Non-immune (N) and HA tag (I) immunoprecipitates
were prepared from MT-transformed NIH3T3 cells expressing vector only
(GREonly), HA-tagged H59Q, or HA-tagged H59Q/301Stop double
mutant which lacks nine carboxyl-terminal amino acids. Immune complexes
were analyzed by SDS-PAGE; proteins were transferred to nitrocellulose;
and immunoblotting was performed with antibodies directed against A
subunit, PME-1, and C subunit (anti-HA tag). The C subunits migrate as
doublets in this gel, but whether double or single bands are seen can
vary (see comments in legend to Fig. 4). The band seen in all lanes in
the PME-1 panel is from the immunoprecipitating antibodies.
Chemiluminescent quantitation (using a Bio-Rad Fluor-S Max MultiImager
or a Roche Molecular Biochemicals Lumi-imager) was used in seven
separate experiments with mixtures of clones to quantify the ratio of
PME-1 to C subunit signal in each lane. In six of seven experiments
with mixes of clones, the double mutant bound less PME-1 than did H59Q,
with a mean reduction of 56 ± 30% and a median value of 39 (range of 8-87%). Thus, PME-1 binding is clearly reduced by loss of
the carboxyl terminus. In a seventh experiment, for unknown reasons,
the double mutant bound 235% of the H59Q level of PME-1, lowering the
overall mean reduction to 28% (median = 40). C, C
subunit carboxyl-terminal antibodies immunoprecipitate reduced amounts
of H59Q·PME-1 complex. Immunoprecipitates were prepared from
MT-transformed NIH3T3 cells expressing HA-tagged H59Q using control
antibody, HA-tag antibody (12CA5), or carboxyl-terminal C subunit
antibodies (1D6, 4B7, 4E1). The immune complexes were analyzed by
SDS-PAGE, proteins were transferred to nitrocellulose, and
immunoblotting was performed with anti-A subunit antibody (upper
panel), anti-PME-1 antibody (middle panel), and anti-C
subunit antibody recognizing both endogenous and HA tagged proteins (1D6 lower panel). The positions of A subunit, the
immunoprecipitating antibody heavy chains (Ab), PME-1,
HA-tagged H59Q C subunit, and untagged, endogenous wt C subunit are
indicated. The C subunits migrate as single bands in this gel, but
whether double or single bands are seen can vary (see comments in
legend to Fig. 4). HA-tagged H59Q C subunit migrates more slowly than
endogenous wt C subunit because of the HA tag.
DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
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ACKNOWLEDGEMENTS |
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We thank Brian Hemmings for the C subunit cDNA, John White for the pGRE vector, and Richard Kahn for providing human cDNA libraries. We are also grateful to Carrie Weaver, Danielle McKelton, Tatiana Mendez, Thomas Loregger, Ingrid Mudrak, Richard Green, and Jay Elliott for expert technical assistance; Jie Yang for advice/help especially on RACE PCR; R. Robinson, J. Neveu, V. Bailey, and E. Spooner of the Harvard Microchemistry Facility for expertise in the high performance liquid chromatography, mass spectrometry, and peptide sequencing; and Cori Beychok, Carlos Moreno, and Anand Viswanathan for critical reading of the manuscript. Under agreements between Upstate Biotechnology Inc. and Emory University and Calbiochem and Emory University, David Pallas is entitled to a share of sales royalty received by the University from these companies. In addition, this same author serves as a consultant to Upstate Biotechnology Inc. The terms of this arrangement have been reviewed and approved by the University in accordance with its conflict of interest policies.
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FOOTNOTES |
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* This work was supported in part by National Institutes of Health Grant CA57327 (to D. C. P.).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.
¶ Supported by an Erwin Schrödinger Postdoctoral Fellowship and Austrian Science Foundation Grant FWF, P12523-MOB.
Contributed equally to the results of this work.
Present address: Cubist Pharmaceuticals, Inc., Cambridge, MA 02139.
¶¶ To whom correspondence should be addressed: Dept. of Biochemistry, Emory University School of Medicine, 1510 Clifton Rd., Atlanta, GA 30322. Tel.: 404-727-5620; Fax: 404-727-3231; E-mail: dpallas{at}emory.edu.
2 L. Haehnel and D. C. Pallas, unpublished data.
3 X. X. Yu, X. Du, E. Ogris, R. E. Green, Q. Feng, L. Clon, and David Pallas, submitted for publication.
4 E. Ogris, I. Mudrak, E. Mak, D. Gibson, and D. C. Pallas, submitted for publication.
5 D. C. Pallas, unpublished data.
6 X. Du and D. C. Pallas, unpublished data.
7 K. C. Nelson and D. C. Pallas, unpublished data.
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ABBREVIATIONS |
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The abbreviations used are: PP2A, protein phosphatase 2A; MT, middle tumor antigen; ST, small tumor antigen; EST, expressed sequence tag; RT-PCR, reverse transcriptase-polymerase chain reaction; PAGE, polyacrylamide gel electrophoresis; UTR, untranslated region; wt, wild type; ORF, open reading frame; PMSF, phenylmethylsulfonyl fluoride.
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REFERENCES |
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