Biochem. J. (2013) 449, 741–749 (Printed in Great Britain)
741
doi:10.1042/BJ20121201
N-Myristoylation is essential for protein phosphatases PPM1A and PPM1B
to dephosphorylate their physiological substrates in cells
Toko CHIDA*†, Masakatsu ANDO*, Tasuku MATSUKI*, Yutaro MASU*, Yuko NAGAURA*, Teruko TAKANO-YAMAMOTO†,
Shinri TAMURA*1 and Takayasu KOBAYASHI*1
*Department of Biochemistry, Institute of Development, Aging and Cancer, Tohoku University, 4-1 Seiryo-machi, Aoba-ku, Sendai 980-8575, Japan, and †Division of Orthodontics and
Dentofacial Orthopedics, Tohoku University Graduate School of Dentistry, 4-1 Seiryo-machi, Aoba-ku, Sendai 980-8575, Japan
PPM [metal-dependent protein phosphatase, formerly called
PP2C (protein phosphatase 2C)] family members play essential
roles in regulating a variety of signalling pathways. While
searching for protein phosphatase(s) that act on AMPK (AMPactivated protein kinase), we found that PPM1A and PPM1B are
N-myristoylated and that this modification is essential for their
ability to dephosphorylate the α subunit of AMPK (AMPKα)
in cells. N-Myristoylation was also required for two other
functions of PPM1A and PPM1B in cells. Although a nonmyristoylated mutation (G2A) of PPM1A and PPM1B prevented
membrane association, this relocalization did not likely cause
the decreased activity towards AMPKα. In in vitro experiments,
the G2A mutants exhibited reduced activities towards AMPKα,
but much higher specific activity against an artificial substrate,
PNPP (p-nitrophenyl phosphate), compared with the wild-type
counterparts. Taken together, the results of the present study
suggest that N-myristoylation of PPM1A and PPM1B plays a
key role in recognition of their physiological substrates in cells.
INTRODUCTION
leading to exposure of the myristoyl moiety sequestered in a
hydrophobic pocket to the cytosol [8].
The PPM [metal-dependent protein phosphatase, formerly
called PP2C (protein phosphatase 2C)] family is one of two
major protein serine/threonine phosphatase families in eukaryotes
[9]. In contrast with members of the other major family, the
PPP (phosphoprotein phosphatase) family, which function as
oligomeric complexes, PPM members do not have regulatory
subunits but contain unique domains that are thought to regulate
catalytic activity, determine substrate specificity and affect
subcellular localization [10]. PPM members are insensitive to
any known protein phosphatase inhibitors, including okadaic acid
and microcystin-LR. The PPM family includes highly conserved
protein phosphatases with 17 distinct genes in the human genome.
These proteins play pivotal roles in regulating the stress response,
cell-cycle progression, apoptosis, Ca2 + signalling, BMP (bone
morphogenetic protein) signalling, metabolism, RNA splicing,
mitochondrial function and lipid transfer [10–13].
The cDNA of PPM1A (formerly PP2Cα) was first isolated
from a rat kidney cDNA library in 1989 [14]. Although PPM1B
(formerly PP2Cβ) is closely related to PPM1A (75 % identity),
knockout mice lacking these isoforms exhibited completely
different phenotypes, suggesting that they have different
physiological roles in vivo [15,16]. Molecular and cellular
biological studies have revealed that PPM1A dephosphorylates
p38 MAPK (mitogen-activated protein kinase), MKK (MAPK
kinase) 4 and MKK6 in the SAPK (stress-activated protein kinase)
cascade, and Smad2/3, proteins in the BMP signalling pathway,
whereas PPM1B dephosphorylates TAK1 [TGF (transforming
Protein N-myristoylation is the irreversible covalent linkage
of the 14-carbon saturated fatty acid myristic acid to the
N-terminal glycine residue of many eukaryotic and viral proteins.
N-Myristoylation provides some proteins with a relatively weak
membrane anchor and is important for correctly localizing them
within the cells [1–3]. N-Myristoylation also occurs on many
proteins that do not appear to associate with membranes. The
role of the myristoyl moiety in these proteins is less clear, but
may involve protein–protein interaction, stabilizing the protein
structure and regulation of the catalytic activity of the enzyme
[1–7]. An example of a protein whose structure is stabilized by a
myristoyl group is PKA (cAMP-dependent protein kinase/protein
kinase A), in which the myristoyl moiety binds in a deep hydrophobic cleft [4]. Although the non-myristoylated form of PKA
is kinetically indistinguishable from the myristoylated enzyme
and both are equal in their ability to associate with the regulatory
subunit, the myristate seems to increase thermal stability of the
kinase, possibly by binding to an adjacent hydrophobic region of
the protein. An example of an enzyme whose catalytic activity
is regulated by myristoylation is c-Abl tyrosine kinase [5,6]. In
c-Abl, the myristoyl group binds to a hydrophobic pocket and
induces a conformational change that allows the docking of the
SH2 (Src homology 2) and SH3 (Src homology 3) domains on
to the kinase domain leading to autoinhibition of the enzyme
[5,6]. Although myristoylation is an irreversible modification, its
function may be regulated via various ‘myristoyl switches’ which
rely on a ligand-dependent conformational change of the protein
Key words: AMP-activated protein kinase (AMPK), N-myristoylation, protein phosphatase 2C (PP2C), metal-dependent
protein phosphatase 1A (PPM1A), metal-dependent protein
phosphatase 1B (PPM1B).
Abbreviations used: AMPK, AMP-activated protein kinase; BMP, bone morphogenetic protein; DMEM, Dulbecco’s modified Eagle’s medium; FBS,
fetal bovine serum; HA, haemagglutinin; HEK, human embyonic kidney; HRP, horseradish peroxidase; ILKAP, ILK1 (integrin-linked kinase 1)-associated
protein; JNK, c-Jun N-terminal kinase; MAPK, mitogen-activated protein kinase; MKK, MAPK kinase; NF-κB, nuclear factor κB; PKA, cAMP-dependent
protein kinase/protein kinase A; PNPP, p -nitrophenyl phosphate; PP, protein phosphatase; PPM, metal-dependent protein phosphatase; SAPK, stressactivated protein kinase; S-HRP, HRP-conjugated S-protein; siRNA, small interfering RNA; SnRK1, SNF1-related kinase 1; TAK1, TGF (transforming growth
factor)-β-activated kinase 1; TNFα, tumour necrosis factor α; WT, wild-type.
1
Correspondence may be addressed to either of these authors (email [email protected] or [email protected]. ac.jp).
c The Authors Journal compilation c 2013 Biochemical Society
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T. Chida and others
growth factor)-β-activated kinase 1], a member of the SAPK
cascade [17–22]. However, cellular studies have shown that both
PPM1A and PPM1B dephosphorylate IKKβ {IκB [inhibitor
of NF-κB (nuclear factor κB)] kinase}, a protein kinase in the
NF-κB pathway [23,24]. These observations suggest that PPM1A
and PPM1B have both non-redundant and redundant roles in cells.
In in vitro experiments, bacterially expressed mammalian PPMA
has broad substrate specificity and even can dephosphorylate
small molecules such as PNPP (p-nitrophenyl phosphate) [25].
But PP2C purified from rabbit liver (probably a mixture of
PPM1A and PPM1B) does not act on this atypical substrate [26],
suggesting that the native enzyme is more selective; however, its
molecular basis remains unknown.
AMPK (AMP-activated protein kinase) acts as a sensor of
cellular energy status [27,28]. AMPK becomes activated in
response to increased AMP concentrations, which then induces
metabolic pathways that generate ATP, while also repressing
ATP consumption that is not essential for short-term survival.
AMPK is a heterotrimeric enzyme comprising a catalytic
α-subunit and regulatory β- and γ -subunits, each of which
contains two or more isoforms encoded by distinct genes. The αsubunits (α1 and α2) contain a kinase domain and are only active
after phosphorylation of Thr172 by upstream kinases, including
LKB1, CaMKKβ [CaM (calmodulin)-dependent protein kinase
kinase β] and TAK1. The β-subunits (β1 and β2) act as
scaffolds and are reportedly myristoylated at their N-terminus,
which is essential for the membrane localization of the AMPK
holoenzyme and to initiate AMPK signalling in response to
AMP [7,29]. The γ -subunits (γ 1–3) bind AMP, which activates
the kinase by promoting Thr172 phosphorylation by upstream
kinases, inhibiting dephosphorylation by protein phosphatases
and exerting direct allosteric effects. In 1991, Moore et al. [30]
proposed that PPM acts as an AMPKα phosphatase on the basis
of the finding that okadaic-acid-insensitive protein phosphatase
activity was effective in dephosphorylating AMPK in a cellfree assay. Experiments using recombinant proteins expressed
in bacteria, as well as siRNA (small interfering RNA), suggested
that PPM1A acts on AMPK [31,32]. Recently, however, Voss et
al. [33] reported that PPM1E (formerly CaMKP-N/POPX1), but
not PPM1A, was responsible for the dephosphorylation of AMPK
in HEK (human embryonic kidney)-293 cells. Their conclusion
was based on observations that siRNA-mediated knockdown of
PPM1E, but not PPM1A, increased AMPKα phosphorylation.
However, whether the other PPM family members also regulate
AMPKα remains to be elucidated. In addition, the specificity of
protein phosphatases for the two isoforms of AMPKα (1 and 2)
has not been determined.
In the present study, we provide evidence that among the
11 PPM isoforms tested, PPM1B, in addition to PPM1A and
PPM1E, contributes to the dephosphorylation of AMPKα in
HeLa cells. Furthermore, we found that PPM1A and PPM1B
are N-myristoylated and that the N-myristoyl moiety of PPM1A
and PPM1B is essential for their recognition of physiological
substrates in cells.
EXPERIMENTAL
Materials
cDNAs encoding mouse PPM1G (PP2Cγ ), mouse ILKAP [ILK1
(integrin-linked kinase 1)-associated protein] (PP2Cδ), mouse
PPM1D (wip1), mouse PPM1E (CaMKP-N/POPX1), human
PPM1F (CaMKP/POPX2), mouse PPM1H (NERPP), human PPM1K (PP2Cκ), human AMPKα1, human AMPKα2,
human AMPKβ1, human AMPKγ 1 and human LKB1 were
c The Authors Journal compilation c 2013 Biochemical Society
cloned using PCR and then subcloned into the pcDNA3 vector.
cDNAs for the expression constructs for mouse PPM1A (PP2Cα),
mouse PPM1B (PP2Cβ), mouse PPM1L (PP2Cε) and mouse
PPM1M (PP2Cη) have been described previously [13,21,34,35].
Details of the expression constructs used in the present study
are shown in Supplementary Table S1 (at http://www.biochemj.
org/bj/449/bj4490741add.htm). The modifying enzymes used for
DNA manipulation were obtained from New England Biolabs.
LipofectamineTM 2000, LipofectamineTM RNAiMax and siRNAs
were obtained from Invitrogen. Sequences of siRNA used in the
present study are shown in Supplementary Table S2 (at http://
www.biochemj.org/bj/449/bj4490741add.htm). PVDF membranes, ECL (enhanced chemiluminescence) kits and
[γ -32 P]ATP were obtained from GE Healthcare. S-protein agarose
beads and S-HRP [HRP (horseradish peroxidase)-conjugated
S-protein] were obtained from Merck. LKBtide (LSNLYHQGKFLQTFCGSPLYRRR, corresponding to amino acids 196–215
of human NUAK2) [36] was obtained from Millipore. RRApTVA
peptide was synthesized by Life Technologies. All other reagents
were purchased from Wako Pure Chemical Industries.
Antibodies
Anti-phospho-AMPKα, anti-AMPKα, anti-phospho-JNK (c-Jun
N-terminal kinase), anti-JNK, anti-phospho-p38 MAPK, anti-p38
MAPK, anti-PPM1A and anti-FLAG antibodies were obtained
from Cell Signaling Technologies. Anti-HA (haemagglutinin)
and anti-Myc antibodies were from Santa Cruz Biotechnology.
The anti-PPM1B antibody has been described previously [15].
Cell culture, transfection and S-pull-down assays
HEK-293 and HeLa cells were grown in DMEM (Dulbecco’s
modified Eagle’s medium, Invitrogen) supplemented with 10 %
(v/v) FBS (fetal bovine serum). Cells were transfected using
LipofectamineTM 2000 (plasmid) or LipofectamineTM RNAiMax
(siRNA) reagent. After a 48-h culture, cells were harvested,
washed twice with PBS, and then lysed with ice-cold lysis buffer
[50 mM Tris/HCl (pH 7.5), 1 % (v/v) Triton X-100, 150 mM
NaCl, 1 mM EGTA, 1 mM EDTA, 1 mM sodium orthovanadate,
50 mM sodium fluoride, 10 mM 2-glycerophosphate, 5 mM
sodium pyrophosphate and CompleteTM protease inhibitor
cocktail (Roche)]. For the S-pull-down assay, cell lysates
containing 200 μg of proteins were incubated for 0.5 h withS-protein agarose beads (5 μl). After washing the beads five times
with lysis buffer, the bound proteins were subjected to SDS/PAGE
(10 % gel) and then transferred on to PVDF membranes. The
membranes were incubated with primary antibody for 1 h at 25 ◦ C,
followed by an HRP-conjugated secondary antibody for 1 h at
25 ◦ C. The membranes were developed using chemiluminescence,
and then quantified using an image analyser (LAS4000 mini, GE
Healthcare).
In vitro kinase assay
Phosphorylation was carried out in a 50 μl incubation mixture,
containing 50 mM Hepes (pH 7.5), 10 mM MgCl2 , 0.1 %
2-mercaptoethanol, 0.1 mM EGTA, 10 nM microcystin-LR,
100 μM LKBtide and 100 μM [γ -32 P]ATP. After 15 min at 30 ◦ C,
the reaction was terminated by spotting aliquots on to P81
phosphocellulose paper, followed by immediate immersion in
75 mM phosphoric acid. The papers were washed, dried and
analysed by Cerenkov counting. One unit of kinase activity was
the amount of enzyme that phosphorylated 1 nmol of LKBtide
within 1 min.
PPM1A and PPM1B are N-myristoylated
Figure 1
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Identification of protein phosphatases acting on AMPKα at Thr172 in cells
(a) HeLa cells were co-transfected with expression plasmids encoding S-HA–AMPKα1, AMPKβ1–HA, Myc–AMPKγ 1, S-HA–LKB1 (except lane 1) and PPM1A–FLAG, PPM1B–FLAG, FLAG–PPM1E,
FLAG–PPM1F, PPM1G–FLAG, PPM1H–FLAG, PPM1L–FLAG, PPM1M–FLAG, ILKAP–FLAG, PPM1K–FLAG or PPM1D–FLAG. S-HA–AMPKα1 was recovered at 48 h post-transfection using
S-protein agarose beads and then immunoblotted with a phospho-specific antibody for AMPKα Thr172 (pT172) or an anti-HA antibody (total protein). The quantification of phospho-Thr172 signal
relative to the total protein level is shown as the mean +
− S.D. for three independent experiments. Whole-cell lysates were also analysed using an anti-FLAG antibody to determine the expression level
of the PPM1 family. (b) Same as (a), except that S-HA–AMPKα2 was used instead of S-HA–AMPKα1. (c) HeLa cells were transfected with the plasmids indicated and S-HA–LKB1 was recovered
from cell lysates (200 μg of protein) using S-protein agarose beads. An in vitro kinase assay was performed using LKBtide as the substrate. (d) HeLa cells were transfected with siRNA targeted
against PPM1A (α1 and α2) or PPM1B (β1 and β2), or a control siRNA for 48 h. The cells were harvested and the cell lysates were subjected to immunoblotting using the antibodies indicated.
Thr172 phosphorylation relative to the total AMPKα levels is shown as the mean +
− S.E.M. for three independent experiments. cont., control.
Metabolic labelling of protein with [3 H]myristic acid
HeLa cells were plated in 35 mm dishes, transfected with
the expression plasmids indicated using a modified calcium
phosphate method [37], and then incubated for 48 h. Cells were
washed twice with serum-free DMEM and incubated for 5 h at
37 ◦ C in DMEM with 2 % (v/v) FBS containing 100 μCi/ml
[3 H]myristic acid. Cells were washed three times with PBS and
lysed with ice-cold lysis buffer. After immunoprecipitation with
an anti-FLAG antibody, the samples were subjected to SDS/PAGE
®
and then fluorography with EN3 HANCE reagent (PerkinElmer).
a 23-gauge needle 25 times and centrifuged at 1000 g for 5 min.
The resulting supernatant was centrifuged at 16 000 g for 20 min.
The pellet was resuspended in buffer C [50 mM Hepes (pH 7.5),
0.25 M sorbitol, 70 mM potassium acetate, 2.5 mM magnesium
acetate, 5 mM EGTA and CompleteTM protease inhibitor cocktail
(Roche)], and then centrifuged again at 16 000 g for 3 min. The
resulting precipitates were resuspended in buffer C to obtain the
microsomal fraction.
Generation of 293 Flp-In cell lines
Isolation of microsomal membranes from cultured cells
Cells were scraped into ice-cold PBS and resuspended in buffer
B [10 mM Hepes (pH 7.5), 0.25 M sorbitol, 10 mM potassium
acetate, 1.5 mM magnesium acetate and CompleteTM protease
inhibitor cocktail (Roche)]. The suspension was passed through
To generate cell lines that stably express PPM1A–FLAG and
PPM1B–FLAG in a doxycycline-inducible manner, the Flp-In
T-REx system (Invitrogen) was used. cDNAs encoding PPM1A
and PPM1B were inserted into the pcDNA5-FRT/TO vector and
introduced into Flp-In 293 cells according to the manufacturer’s
instructions (Invitrogen).
c The Authors Journal compilation c 2013 Biochemical Society
744
Figure 2
T. Chida and others
PPM1A and PPM1B are N-myristoylated
(a) Sequence alignment of the N-terminus of human AMPKβ, c-Src, PPM1A and PPM1B. (b) HeLa cells were transfected with expression plasmids encoding PPM1A–FLAG or PPM1B–FLAG using
a modified calcium phosphate method and incubated for 48 h. The cells were washed twice with serum-free DMEM and incubated for 5 h at 37 ◦ C in DMEM with 2 % (v/v) FBS containing 100 μCi/ml
[3 H]myristic acid. Labelled proteins were recovered by immunoprecipitation and the samples were subjected to SDS/PAGE, followed by fluorography. The bottom panel shows a gel stained with
Coomassie Brilliant Blue (CBB). IgG, immunoglobulin used for immunoprecipitation. (c) HEK-293 cells were transiently transfected with the plasmids indicated using a modified calcium phosphate
method and incubated for 48 h. The cells were resuspended in buffer B, passed through a 23-gauge needle 25 times, and centrifuged at 1000 g for 5 min at 4 ◦ C. The resulting supernatant was
centrifuged at 16 000 g for 20 min at 4 ◦ C to obtain the cytosolic (supernatant) and membranous (pellet) fractions. The fractions were immunoblotted with an anti-FLAG antibody. PPM1L (membrane)
and N-PPM1L (cytosol) were used as controls.
Cell proliferation assay
To evaluate the cell proliferation rate, 293 Flp-In cell lines were
seeded in 96-well plates at a concentration of 5×103 cells/well
in 100 μl of DMEM. The next day, the medium was changed
to DMEM containing 30 ng/ml doxycycline and the cells were
incubated for an additional 24 and 48 h. At 1 h after adding 10 μl
of WST-8 solution (Dojin) to the culture medium, the absorbance
at 450 nm was measured using a microplate reader.
activity was defined as the amount of enzyme that
dephosphorylated 1 nmol of substrate in 1 min. For the kinetic
parameter K metal , the metal ion (Mn2 + ) concentration was altered
from 0.01 to 1 mM. K metal values were determined using the
Hanes–Woolf linearization method. PNPP phosphatase activity
was measured in a reaction mixture (50 μl) containing 50 mM
Hepes (pH 7.5), 0.5 mM MnCl2 , 0.02 % 2-mercaptoethanol and
20 mM PNPP. The reaction was carried out at 30 ◦ C for 45 min
and then terminated by adding 0.5 M NaOH, after which the
absorbance at 405 nm was measured.
Purification of proteins expressed in mammalian cells
HEK-293 cells plated in 10 cm dishes were transiently transfected
with plasmid constructs encoding PPM1A–FLAG [WT (wildtype) or G2A mutant], PPM1B (WT or G2A) or the AMPK
complex (co-expression of FLAG–AMPKα1, AMPKβ1–HA and
Myc–AMPKγ 1) using a modified calcium phosphate method
[37]. After transfection, the cells were cultured for 48 h and then
harvested. Cells transfected with the AMPK complex were treated
with 10 mM 2-deoxyglucose for 1 h before harvest to activate the
AMPK complex. The cells were washed twice with PBS and
lysed with ice-cold lysis buffer. Cell lysates were incubated for
2 h with anti-FLAG agarose beads (50 μl, Sigma). The beads
were washed three times with lysis buffer and twice with HBSSG [50 mM Hepes (pH 7.5), 150 mM NaCl, 0.25 M sucrose and
2 % (v/v) glycerol] and then the bound proteins were eluted with
HBS-SG containing 200 μg/ml 3×FLAG peptide (Sigma). All
in vitro experiments were performed using proteins that were
expressed in mammalian cells.
Protein phosphatase activity assay
Protein phosphatase activity was determined by measuring the
release of Pi from RRApTVA phosphopeptide as a substrate.
The standard assay mixture contained 50 mM Hepes (pH 7.5),
0.02 % 2-mercaptoethanol, 100 μM RRApTVA phosphopeptide
and 0.5 mM MnCl2 . The enzyme reaction was carried out at 30 ◦ C
for 45 min. The reaction was started by adding the enzyme and
terminated by adding 100 μl of Biomol Green reagent (Enzo Life
Sciences). The absorbance at 620 nm was measured. Pi release
was calculated using a standard curve with phosphoric acid
that was treated as described above. One unit of phosphatase
c The Authors Journal compilation c 2013 Biochemical Society
RESULTS
AMPKα is dephosphorylated by PPM1A and PPM1B in cells
To identify the PPM family members that are responsible
for dephosphorylating the α isoform of AMPK (α1 and α2)
in HeLa cells, we examined the effects of overexpressing
PPM members on LKB1-dependent phosphorylation of AMPKα
at Thr172 . Expressing S-HA–LKB1 alone in HeLa cells,
which do not express LKB1 due to abnormal methylation of
the promoter region, induced phosphorylation of endogenous
AMPKα. However, we could not distinguish the phosphorylation
level of AMPKα1 from that of AMPKα2 because a phosphospecific antibody against phospho-Thr172 of AMPKα detected
both isoforms (results not shown). Therefore we transfected
HeLa cells with expression plasmids for two different AMPK
complexes (AMPKβ1–HA, Myc–AMPKγ 1, and either S-HA–
AMPKα1 or α2) and S-HA–LKB1 and then recovered
S-HA–AMPKα from the cell lysates to detect the phosphorylation
of AMPKα1 and α2 separately. Under these conditions, both
S-HA–AMPKα1 and α2 were phosphorylated in an LKB1dependent manner, and co-expressing PPM1A and PPM1B
caused a large decrease in the phosphorylation of both
isoforms with no effect on LKB1 activity (Figures 1a–1c).
Furthermore, siRNAs targeting PPM1A or PPM1B increased
AMPKα phosphorylation in HeLa cells (Figure 1d). These results
indicate that both PPM1A and PPM1B are responsible for the
dephosphorylation of AMPKα1 and α2 in cells. In contrast,
PPM1E overexpression decreased phosphorylation of AMPKα2,
but not AMPKα1 (Figures 1a and 1b).
PPM1A and PPM1B are N-myristoylated
745
PPM1A and PPM1B are N-myristoylated
The β-subunit of AMPK is N-myristoylated and this modification
was reported to be involved in regulating the key steps of AMPK
activation [7,29]. Although the N-termini of PPM1A and PPM1B
do not completely match the consensus sequence for N-myristoyl
transferase (MGXXXS/T, where X is any amino acid residue), a
portion of the N-terminal sequence was same as that of c-Src, a
well-known N-myristoylated protein (Figure 2a) [38]. These facts
motivated us to examine whether PPM1A and PPM1B are also Nmyristoylated. HeLa cells expressing PPM1A–FLAG or PPM1B–
FLAG were metabolically labelled with [3 H]myristic acid and the
incorporation of radioactivity into these proteins was analysed.
As shown in Figure 2(b), [3 H]myristic acid was incorporated into
WT but not the G2A mutant, indicating that these proteins were
N-myristoylated.
Since N-myristoylation has been shown to facilitate protein–
membrane interactions [3], we examined whether the G2A
mutation affected the subcellular localization of PPM1A and
PPM1B. When extracts of HEK-293 cells expressing these
proteins were separated into cytosolic and membrane fractions,
WT PPM1A and PPM1B were predominantly recovered in the
cytosolic fraction. However, a small, but significant, amount
of these proteins were recovered in the membrane fraction
(Figure 2c). Distribution to the membrane fraction markedly
decreased in cells expressing the G2A mutants (Figure 2c).
These results suggest that PPM1A and PPM1B weakly associate
at the membrane and N-myristoylation mediates the membrane
localization of these isoforms.
PPM1A and PPM1B must be N-myristoylated to dephosphorylate
their physiological substrates in cells
To investigate the possible role of N-myristoylation in the
activities of PPM1A and PPM1B, we examined whether the G2A
mutants could dephosphorylate the α-subunit of AMPK in cells.
In contrast with ectopically expressed WT PPM1A and PPM1B,
which caused a large decrease in AMPKα phosphorylation, the
G2A mutants were unable to reduce AMPKα phosphorylation.
Essentially there was no significant alteration in the expression
levels of AMPKα/β/γ , and LKB1 was observed upon expression
of either WT or the mutant PPM1A/PPM1B in cells (Figure 3a).
We then determined whether N-myristoylation is required for
the general function of these two isoforms. Since PPM1A and
PPM1B reportedly regulate the SAPK cascade [17,21,22], we
examined the effects of overexpressing PPM1A and PPM1B with
or without the G2A mutation on TNFα (tumour necrosis factor
α)-dependent phosphorylation of p38 MAPK and JNK, the most
downstream components of the SAPK cascade. While ectopic
expression of both WT PPM1A and PPM1B in cells caused a
large decrease in TNFα-enhanced phosphorylation of both JNK
and p38 MAPK, the G2A mutants had little effect (Figure 3b).
Ofek et al. [39] reported that PPM1A inhibited the proliferation
of HEK-293 cells through activation of the p53 protein [39].
Therefore we tested the effects of expressing PPM1A or its G2A
mutants on HEK-293 cell proliferation. As shown in Figure 3(c),
expressing WT PPM1A inhibited cell growth at 48 h after the
addition of doxycycline. In contrast, the proliferation rate of cells
expressing the G2A mutant was similar to control cells.
Taken together, these results suggest that N-myristoylation
is indispensable for the ability of PPM1A and PPM1B to
dephosphorylate not only AMPK but also the other endogenous
substrates involved in the stress response and cell growth
regulation.
Figure 3 The G2A mutants of PPM1A and PPM1B are unable to
dephosphorylate their physiological substrates in cells
(a) HeLa cells were transiently transfected with the indicated plasmids and incubated for 48 h.
S-HA–AMPKα1 was recovered from cell lysates using S-protein agarose beads. The samples
were subjected to immunoblotting with anti-phospho-AMPKα (pT172) and anti-HA antibodies.
Whole-cell lysates (WCL) were also analysed to determine the cellular expression level of each
protein. EV, empty vector. (b) 293 Flp-In cell lines expressing the indicated constructs were
seeded into 12-well plates and incubated for 24 h (EV, empty vector). The medium was replaced
with serum-free DMEM containing doxycycline (30 ng/ml) and then incubated for an additional
24 h. After treating the cells with TNFα (20 ng/ml) for 10 min, cell lysates were prepared and
subjected to immunoblotting using the antibodies indicated. (c) 293 Flp-In cell lines expressing
the indicated constructs were seeded into 96-well plates and incubated for 24 h (EV, empty
vector). The medium was replaced with DMEM containing doxycycline (30 ng/ml) and then the
cells were incubated for 24 h or 48 h. After adding WST-8 solution to the culture medium, the
absorbance at 450 nm was measured. The results are expressed as the ratio of the absorbance
at 24 h and 48 h to that obtained at 0 h. The results are shown as the means +
− S.D. for three
independent experiments.
N-Myristoylation is required for PPM1A and PPM1B to recognize
their physiological substrates in cells
Since mutating the N-myristoylation site in PPM1A and
PPM1B affected the membrane association of these proteins,
we suspected that proper intracellular localization mediated
by N-myristoylation was required for PPM1A and PPM1B to
dephosphorylate their physiological substrates. It was previously
reported that membrane association of the AMPK complex
mediated by N-myristoylation of the β-subunit is required for
the metabolic stress-sensing function of the AMPK complex
[7]. Therefore we asked whether co-localization of the AMPK
complex and PPM1A or PPM1B at the membrane was required
for these phosphatases to dephosphorylate AMPKα. To address
this question, we examined whether PPM1A or PPM1B could
c The Authors Journal compilation c 2013 Biochemical Society
746
Figure 4
T. Chida and others
N-Myristoylation is an essential determinant of substrate specificity for PPM1A and PPM1B
(a) HeLa cells were transiently transfected with S-HA–LKB1, AMPKβ1-S (WT or G2A) and increasing amounts of PPM1A–FLAG or PPM1B–FLAG and then incubated for 48 h. The AMPK complex
was recovered by S-tag-mediated pull down of AMPKβ1-S, and phosphorylation of endogenous AMPKα in the complex was analysed using an anti-phospho-Thr172 antibody. The membrane was
reprobed with anti-AMPKα and S-HRP to confirm the amount of AMPKα, S-HA–LKB1 and AMPKβ1-S. Whole-cell lysates (WCL) were also subjected to immunoblotting with an anti-FLAG antibody.
In the absence of PPM1A or PPM1B expression, AMPKα complexed with the mutant β-subunit was more highly phosphorylated than AMPKα complexed with the WT β-subunit, as was previously
reported [29]. The efficiency of AMPKα dephosphorylation was quantified and is shown on the right-hand side. (b) A phosphatase assay was performed in a reaction mixture (50 μl) containing
50 mM Hepes (pH 7.5), 0.5 mM MnCl2 , 0.02 % 2-mercaptoethanol, 100 μM RRApTVA peptide and 8 μg/ml protein phosphatase. The enzyme reaction was carried out at 30 ◦ C for 45 min and
then terminated by adding Biomol Green Reagent, after which the absorbance at 650 nm was measured using a plate reader. The results are shown as the means +
− S.E.M. for three independent
experiments. (c) Phosphatase activity against RRApTVA phosphopeptide was measured with increasing concentrations of MnCl2 . The results are expressed as a ratio of the activity to the highest
activity. (d) The AMPK complex was incubated with PPM1A (WT), PPM1A (G2A), PPM1B (WT) or PPM1B (G2A) in a reaction mixture (50 μl) containing 50 mM Hepes (pH 7.5), 0.5 mM MnCl2
and 0.02 % 2-mercaptoethanol at 30 ◦ C for 20 min. The amount of protein phosphatases added to the reaction was equalized for their ability to dephosphorylate RRApTVA peptide (0.15, 0.3, 0.45 and
0.6 units/ml). At the end of the reaction, the samples were analysed by immunoblotting using anti-pAMPKα (phospho-Thr172 ) and anti-AMPKα antibodies. (e) A phosphatase assay was performed
in a reaction mixture (50 μl) containing 50 mM Hepes (pH 7.5), 0.5 mM MnCl2 , 0.02 % 2-mercaptoethanol, 20 mM PNPP and 8 μg/ml protein phosphatase. The reaction was carried out at 30 ◦ C
for 45 min and then terminated by adding 0.5 M NaOH, after which the absorbance at 405 nm was measured using a plate reader. The results are shown as the means +
− S.E.M. for three independent
c The Authors Journal compilation c 2013 Biochemical Society
PPM1A and PPM1B are N-myristoylated
dephosphorylate AMPKα that forms a complex with a nonmyristoylated mutant of AMPKβ (G2A). LKB1 and AMPKβ-S
(WT or G2A) were co-expressed with PPM1A or PPM1B in HeLa
cells and phosphorylation of endogenous AMPKα complexed
with AMPKβ-S (WT or G2A) was analysed after the AMPK
complex was recovered by S-tag-mediated pull down of AMPKβS. PPM1A and PPM1B similarly dephosphorylated AMPKα
that was complexed with either WT or the G2A version of
AMPKβ (Figure 4a). These results indicated that both PPM1A
and PPM1B could dephosphorylate AMPKα, irrespective of its
subcellular localization, suggesting that PPM1A and PPM1B
do not have to co-localize with the AMPK complex at the
membrane to dephosphorylate AMPKα. This assumption is
supported by evidence that PPM1E, which has a disrupted Nmyristoylation site, is also able to dephosphorylate AMPKα in
cells.
To test the possibility that N-myristoylation of PPM1A and
PPM1B directly affects their activities, we expressed WT and the
G2A mutant of PPM1A–FLAG and PPM1B–FLAG in HEK293 cells, immunoprecipitated these proteins with an antiFLAG antibody, and then measured the enzymatic activities of
these proteins towards various substrates in vitro. When the
RRApTVA phosphopeptide, which is widely used for in vitro
phosphatase assays, was used as the substrate, the G2A mutants
of PPM1A and PPM1B showed a slight and 50 % increase in
specific activity compared with the WT enzymes respectively
(Figure 4b). Furthermore, there was no apparent difference in
Mn2 + -dependency [Figure 4c, K metal values for PPM1A WT,
PPM1A (G2A), PPM1B WT and PPM1B (G2A) were 0.011,
0.019, 0.015 and 0.011 mM respectively]. These results suggest
that the non-myristoylated forms of PPM1A and PPM1B have
catalytic activities towards the phosphopeptide that are similar
to the WT enzymes. We then analysed the dephosphorylation
activity towards an AMPK complex that had been activated
with 2-deoxyglucose and then affinity-purified from HEK-293
cells. Both WT PPM1A and PPM1B efficiently dephosphorylated
AMPKα, whereas the G2A mutants had much lower activities
towards AMPKα, although the amounts of the phosphatases
in the reaction mixture were adjusted to obtain equal activities
towards the RRApTVA peptide (Figure 4d). Interestingly, when
an artificial substrate, PNPP, was used, both PPM1A (G2A) and
PPM1B (G2A) had much higher specific activities compared
with the WT enzymes (Figure 4e). These results indicated that
PPM1A and PPM1B showed lower enzymatic activity towards
the physiological substrate (AMPK), but had higher specific
activity against the artificial substrate (PNPP) in the absence of the
N-myristoyl group. Taken together, these results suggest that
the N-myristoyl group is essential for the recognition of
physiological substrates by PPM1A and PPM1B in cells.
DISCUSSION
In the present study we have provided evidence that Nmyristoylation is required for PPM1A and PPM1B to function
in cells. Importantly, N-myristoylation is essential for the
recognition of physiological substrates by PPM1A and PPM1B
in cells, but dispensable for their catalytic activity itself. This
conclusion is based on the following observations. First, a nonmyristoylated mutant (G2A) of PPM1A and PPM1B could not
747
dephosphorylate endogenous substrates, including AMPKα, p53
and components of the SAPK cascade, in cells (Figure 3).
Secondly, in in vitro experiments, the G2A mutants exhibited
reduced activities towards AMPKα, whereas they exhibited
higher specific activity against an artificial substrate (PNPP) than
the WT protein (Figures 4d and 4e). Finally, the G2A mutation did
not affect Mn2 + -dependency of PPM1A and PPM1B (Figure 4c).
There are two possible mechanisms by which N-myristoylation
is involved in substrate recognition: (i) the myristoyl group
may increase the affinity of the enzyme for its substrate by
interacting directly with the substrate protein and (ii) the myristoyl
group may interact with the intramolecule-binding pocket and
induce conformational changes in the enzyme that affect its
affinity for the substrate protein. Considering our observation
that G2A mutants exhibited higher specific activity against an
artificial substrate (PNPP) than the WT protein, it is unlikely that
myristoylation directly associates with the substrate. Rather, the
myristoyl group may preserve the conformation of the catalytic
pocket by interacting with hydrophobic residues near the active
site, which allows only the physiological substrate proteins to
access the catalytic site. In the absence of the modification, the
structure around the active site may be destabilized, so that the
artificial substrates with low molecular mass readily approach
the catalytic site. This conclusion may be reinforced by the
fact that Gly2 is located at the entrance of the catalytic pocket
(Figure 4f) [40]. It remains uncertain whether N-myristoylation
is involved in regulation of PPM1A and PPM1B function in cells,
but displacement of the myristoyl moiety from the hydrophobic
pocket by an as yet unidentified mechanism might lead to change
of substrate recognition of these isoforms.
In the present study we also provide evidence that PPM1A and
PPM1B redundantly dephosphorylate the α-subunit of AMPK.
Although these isoforms are closely related (75 % identity),
they were shown to be distinctly localized in cells [15,19,41],
raising the possibility that they might act on AMPK heterotrimeric
complexes at different subcellular locations within cells. Recently,
Voss et al. [33] reported that PPM1E is a protein phosphatase that
acts on AMPK. In our assay, PPM1E decreased phosphorylation
only of AMPKα2 and had little effect on AMPKα1 (Figures 1a
and 1b). Considering the observation by Voss et al. [33]
that endogenous PPM1E specifically interacted with AMPKα2,
PPM1E may act as an AMPKα2-containing-complex-specific
phosphatase. In the same paper [33], the authors claimed that
PPM1A was not involved in the dephosphorylation of AMPK on
the basis of the finding that PPM1A-specific shRNA (short hairpin
RNA) did not increase AMPKα phosphorylation. Although
the reason for the discrepancy between their observations and the
findings of the present study is not known, there might be
differences in relative PPM1A and PPM1E expression in the
cell lines used in the knockdown studies, resulting in different
responses with the PPM1A-specific siRNA.
PPM1A and PPM1B expressed in Escherichia coli have
been generally used by researchers for in vitro experiments.
However, it should be noted that bacteria do not have an
N-myristoyltransferase gene [3] and therefore are not capable
of modifying PPM1A and PPM1B. Therefore using recombinant
PPM1A and PPM1B produced in mammalian or other eukaryotic
cells is highly recommended for in vitro experiments.
During the preparation of the present paper, Tsugama et al. [42]
reported that WT PP2C74, a PPM family protein phosphatase
experiments. (f) Two orthogonal views of the molecular-surface representation of PPM1A. Gly2 is indicated in red. Amino acid residues composing the active site (Asp38 , Asp60 , Thr128 , Asp146 ,
Asp199 , Asp239 and Asp282 ) are coloured in yellow, and hydrophobic residues (leucine, valine and isoleucine) are shown in cyan. The catalytic metals in the catalytic pocket are difficult to see in this
projection. The structure is displayed using PyMOL (http://www.pymol.org). The PDB code for PPM1A is 3FXJ.
c The Authors Journal compilation c 2013 Biochemical Society
748
T. Chida and others
in Arabidopsis, but not its mutant with an N-terminal glycineto-alanine substitution, localizes to the plasma membrane in
Arabidopsis and interacts with SnRK1 (SNF1-related kinase
1), the plant orthologue of mammalian AMPK [42]. Using in
silico analysis, they found that the N-terminal glycine residue
of PP2C74 could be a consensus site for myristoylation and
proposed that its myristoylation is required for the plasma
membrane localization of SnRK1 and its association with SnRK1.
Although they did not demonstrate that PP2C74 is myristoylated
or show that its activity is altered upon myristoylation in cells,
these observations raise the possibility that myristoylation is
a conserved mechanism to functionally regulate PPM family
members in a variety of organisms.
AUTHOR CONTRIBUTION
Takayasu Kobayashi and Shinri Tamura designed the overall experiments and wrote the
paper. Toko Chida executed the major parts of this work. Masakatsu Ando, Tasuku Matsuki
and Yutaro Masu helped with the initial experiments. Yuko Nagaura and Teruko TakanoYamamoto contributed to a comprehensive interpretation of the results and data analyses.
FUNDING
This work was supported, in part, by a grant-in-aid for Scientific Research from the Ministry
of Education, Culture, Sports, Science and Technology of Japan.
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Received 30 July 2012/19 October 2012; accepted 22 October 2012
Published as BJ Immediate Publication 22 October 2012, doi:10.1042/BJ20121201
c The Authors Journal compilation c 2013 Biochemical Society
Biochem. J. (2013) 449, 741–749 (Printed in Great Britain)
doi:10.1042/BJ20121201
SUPPLEMENTARY ONLINE DATA
N-Myristoylation is essential for protein phosphatases PPM1A and PPM1B
to dephosphorylate their physiological substrates in cells
Toko CHIDA*†, Masakatsu ANDO*, Tasuku MATSUKI*, Yutaro MASU*, Yuko NAGAURA*, Teruko TAKANO-YAMAMOTO†,
Shinri TAMURA*1 and Takayasu KOBAYASHI*1
*Department of Biochemistry, Institute of Development, Aging and Cancer, Tohoku University, 4-1 Seiryo-machi, Aoba-ku, Sendai 980-8575, Japan, and †Division of Orthodontics and
Dentofacial Orthopedics, Tohoku University Graduate School of Dentistry, 4-1 Seiryo-machi, Aoba-ku, Sendai 980-8575, Japan
Table S1
Expression constructs used in the present study
®
Name
Synonym
Species
GenBank accession No.
Tag
Tag position
Reference
PPM1A
PPM1B
PPM1D
PPM1E
PPM1F
PPM1G
PPM1H
PPM1K
PPM1L
PPM1M
ILKAP
AMPKα1
AMPKα2
AMPKβ1
AMPKγ 1
LKB1
PP2Cα
PP2Cβ
Wip1
CaMKP-N/POPX1
CaMKP/POPX2
PP2Cγ
NERPP
PP2Ck/PP2Cm
PP2Cε
PP2Cη-2
PP2Cδ
Mouse
Mouse
Mouse
Mouse
Human
Mouse
Mouse
Human
Mouse
Mouse
Mouse
Human
Human
Human
Human
Human
NM_008910
NM_001159496
AF200464
XM_282986
D13640
BC009004
AK051434
NM_152542
AY184801
AB474373
BC026953
NM_006251
NM_006252
NM_006253
NM_002733
U63333
FLAG
FLAG
FLAG
FLAG
FLAG
FLAG
FLAG
FLAG
FLAG
FLAG
FLAG
S-HA, Myc
S-HA
HA-S
Myc
S-HA, Myc
C-terminal
C-terminal
C-terminal
N-terminal
N-terminal
C-terminal
C-terminal
C-terminal
C-terminal
C-terminal
C-terminal
N-terminal
N-terminal
C-terminal
N-terminal
N-terminal
The present study
The present study
The present study
The present study
The present study
The present study
The present study
The present study
[1]
[2]
The present study
The present study
The present study
The present study
The present study
The present study
Table S2
Sequences of siRNA used in the present study
siRNA
Sequence (5 →3 )
Target
A1
A2
B1
B2
UUAAGACACCUACAGCUGUUGACCC
AAAUUAAGACACCUACAGCUGUUGA
UUAGGUGAAAUCAUAACUCCCACUG
UUAACACGUUGUAUCAUCACGCUGC
Human PPM1A
Human PPM1A
Human PPM1B
Human PPM1B
REFERENCES
1 Saito, S., Matsui, H., Kawano, M., Kumagai, K., Tomishige, N., Hanada, K., Echigo, S.,
Tamura, S. and Kobayashi, T. (2008) Protein phosphatase 2Cε is an endoplasmic reticulum
integral membrane protein that dephosphorylates the ceramide transport protein CERT to
enhance its association with organelle membranes. J. Biol. Chem. 283, 6584–6593
2 Henmi, T., Amano, K., Nagaura, Y., Matsumoto, K., Echigo, S., Tamura, S. and Kobayashi,
T. (2009) A mechanism for the suppression of interleukin-1-induced nuclear factor κB
activation by protein phosphatase 2Cη-2. Biochem. J. 423, 71–78
Received 30 July 2012/19 October 2012; accepted 22 October 2012
Published as BJ Immediate Publication 22 October 2012, doi:10.1042/BJ20121201
1
Correspondence may be addressed to either of these authors (email [email protected] or [email protected]. ac.jp).
c The Authors Journal compilation c 2013 Biochemical Society