From the Division of Medicinal Chemistry,
§ Division of Toxicology, College of Pharmacy and
¶ Graduate Programs in Biochemistry and Molecular Biology and the
Center for Molecular and Cellular Toxicology, University of Texas,
Austin, Texas 78712
Received for publication, September 26, 2000, and in revised form, October 31, 2000
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ABSTRACT |
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The mitogen-activated protein kinases
(MAPKs) are a family of enzymes conserved among eukaryotes that
regulate cellular activities in response to numerous external signals.
They are the terminal component of a three-kinase cascade that is
evolutionarily conserved and whose arrangement appears to offer
considerable flexibility in encompassing the diverse biological
situations for which they are employed. Although multistep protein
phosphorylation within mitogen-activated protein kinase (MAPK) cascades
can dramatically influence the sensitivity of signal propagation, an
investigation of the mechanism of multisite phosphorylation by a MAPK
has not been reported. Here we report a kinetic examination of the
phosphorylation of Thr-69 and Thr-71 of the glutathione
S-transferase fusion protein of the
trans-activation domain of activating transcription
factor-2 (GST-ATF2-(1-115)) by p38 MAPK The mitogen-activated protein kinases
(MAPKs)1 are a family of
enzymes conserved among eukaryotes that regulate cellular activities in
response to numerous external signals. They are implicated in processes
ranging from pheromone responses and cell wall formation in yeast to
mitogenesis, apoptosis, and stress responses in mammalian cells (1, 2).
They are the terminal components of a three-kinase cascade that is
evolutionarily conserved and whose arrangement appears to offer
considerable flexibility in encompassing the diverse biological
situations for which they are employed.
A MAP kinase cascade is generally composed of three protein kinases, a
MAP kinase kinase kinase (MAPKKK), a MAP kinase kinase (MAPKK), and a
MAP kinase (MAPK) arranged in a linear hierarchical fashion as shown in
Scheme 1. Although there are a large
number of MAPKKKs, which become activated through a variety of
mechanisms, MAPKKKs are highly specific for their conjugate MAPKK,
which they activate by dual phosphorylation. In turn MAPKKs activate
their conjugate MAPKs, also by dual phosphorylation, whereas MAPKs, which are the least specific kinases in these modules, phosphorylate many proteins, typically at more than one site (3). Although this
simple picture is somewhat obscured by the existence of an abundance of
different MAPKKKs and by the existence of MAPKK and MAPK isoforms, the
three-tiered hierarchy appears to be well conserved. The kinases do not
function in isolation, because protein phosphatases represent the force
against which they labor to drive the steady-state concentrations of
phosphorylated proteins beyond critical thresholds. There are several
protein phosphatases, some such as the MAPK phosphatases are specific
for certain MAPKs (4-6) whereas others such as protein phosphatases 1 (7) and 2 (8-10) dephosphorylate many cellular proteins.
(p38
) as a model system
for the phosphorylation of ATF2 by p38
. Our experiments demonstrated that GST-ATF2-(1-115) is phosphorylated in a two-step distributive mechanism, where p38
dissociates from GST-ATF2-(1-115) after the
initial phosphorylation of either Thr-69 or Thr-71. Whereas p38
showed similar specificity for Thr-71 and Thr-69 in the
unphosphorylated protein, it displayed a marked difference in
specificity toward the mono-phosphoisomers. Phosphorylation of Thr-71
had no significant effect on the rate of Thr-69 phosphorylation, but
Thr-69 phosphorylation reduced the specificity,
kcat/KM, of p38
for
Thr-71 by approximately 40-fold. Computer simulation of the mechanism suggests that the activation of ATF2 by p38
in vivo is
essentially Michaelian and provides insight into how the kinetics of a
two-step distributive mechanism can be adapted to modulate effectively the sensitivity of a signal transduction pathway. This work also suggests that whereas MAPKs utilize docking interactions to bind substrates, they can be weak and transient in nature, providing just
enough binding energy to promote the phosphorylation of a specific substrate.
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
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Scheme 1.
Although an arrangement of kinases in a cascade such as a MAPK cascade
generally leads to amplification of a signal, it also provides
considerable means for control. There has been substantial interest
recently in the mechanism of signal propagation through MAPK cascades
and, in particular, the generation of thresholds (11). Potential
sources of thresholds in a MAPK cascade include contributions from
zero-order ultrasensitivity2
(12, 13) and multistep mechanisms (12, 13), as well as other mechanisms
such as the regulated translocation of proteins between organelles
(14). Another potential source of ultrasensitivity is multistep
phosphorylation (11). Indeed, the dual phosphorylation of MAPK by MAPKK
has been shown to occur through a two-step distributive mechanism (15,
16), and theoretical calculations support the notion that this
mechanism contributes to the observed switch-like behavior of the MAPK
cascade responsible for the turning on and off of the cell cycle in
oocytes (17). Although the generation of thresholds within signaling
cascades is important, they may not always be necessary or even
beneficial. For example, control over a wider range of stimulus may be
more appropriate in some situations, such as when cells are exposed to
certain stresses. MAPKs activate many proteins by dual phosphorylation,
which is, potentially, an important source of control and which to our
knowledge has not been investigated before. Because they are known to
form complexes with potential substrates in cells and utilize discrete docking interactions to bind substrates, a major unanswered question is
whether or not they phosphorylate multiple sites on a substrate processively. To expand our current mechanistic knowledge of MAPK cascades and in particular the MAPK enzymes and to characterize further
possible mechanisms of control, we studied the kinetic mechanism of the
dual phosphorylation of the glutathione S-transferase fusion
protein of ATF2 (GST-ATF2-(1-115)) by the MAPK, p38. This fusion
protein corresponds to the trans-activation domain of ATF2 and is an excellent protein substrate for p38
(18) and an excellent model for the phosphorylation of the full-length protein.
p38 (19, 20) is involved in relaying stress-related signals in
mammalian cells. It is regulated by two MAPKKs termed MAP kinase kinase
3 (MKK3) and MAP kinase kinase 6 (MKK6) (21, 22), which phosphorylate
the activation loop of p38
twice, once on a tyrosine and once on a
threonine. Several activators of one or both of these enzymes have been
identified that include MEKK4 (MAP or ERK kinase kinase) (23-25),
apoptosis-stimulated kinase (26, 27), and transforming growth
factor-
-activated kinase (28-30). The basic region-leucine zipper
protein activating transcription factor 2 (ATF2) is a substrate of
p38
and other stress-activated MAPKs. It is a DNA-binding protein
that forms a homodimer or heterodimer with c-Jun, binds to cyclic
AMP-response elements (CREs), and stimulates CRE-dependent
transcription of genes. Recently, it was also shown to be a histone
acyltransferase, specific for histones H2B and H4 (31). Increases in
its transcriptional activity (32), its acyltransferase activity (31),
and its cellular stability (33) are associated with the dual
phosphorylation of ATF2 on Thr-69 and Thr-71.
We found that the kinetic mechanism for the dual phosphorylation of the
two residues within GST-ATF2-(1-115), Thr-69 and Thr-71, occurs by a
two-step mechanism where p38 dissociates from the protein after each
phosphorylation event. The kinetics of this process is predicted to
have important implications for the activation of ATF2 by p38
in vivo and highlights the potential that two-step distributive mechanisms have for controlling the amplitude sensitivity of signal transduction pathways.
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EXPERIMENTAL PROCEDURES |
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Materials--
Trizma base was purchased from EM Reagents
(Gibbstown, NJ) and ammonium carbonate from Fisher. All other buffer
components and chemicals were obtained from Sigma. Qiagen Inc. (Santa
Clarita, CA) supplied nickel-nitrilotriacetic acid-agarose. GST fusion proteins were purified with glutathione-agarose from Sigma. Kinase assays were conducted with special quality ATP from Roche Molecular Biochemicals and [-32P]ATP from ICN (Costa Mesa, CA).
Proteins were isolated on Optitran nitrocellulose membrane from
Schleicher & Schuell and digested with sequencing grade trypsin from
Roche Molecular Biochemicals. Plasmids used to express
GST-ATF2-(1-115) (18) and His-p38
(35) have been reported previously.
Expression and Purification of Rat p38 MAP Kinase
--
Activated mouse p38
(19, 20) was expressed in
Escherichia coli (BL21 DE3 pLys S) with an N-terminal
hexahistidine tag (34) to facilitate its purification, essentially
according to the method of Khokhlatchev et al. (35). This
method typically yields ~0.25 mg of enzyme per liter of starting
culture with a specific activity of 970 nmol/min/mg (250 µM ATP and 50 µM GST-ATF2-(1-115)).
Expression and Purification of GST-ATF2-(1-115)-- The GST-ATF2-(1-115) construct was expressed as a glutathione S-transferase fusion protein in E. coli (BL21 DE3 pLys S) and purified essentially according to the method of LoGrasso et al. (18) with an additional high resolution Mono-Q anion exchange chromatography step. Protein fractions from a glutathione-agarose column (obtained by following standard protocols (18)) were applied to a Mono-Q HR 5/5 chromatography column pre-equilibrated with buffer B (50 mM Tris-HCl, pH 7.5, 0.1% (by volume), 2-mercaptoethanol, 0.2 mM phenylmethylsulfonyl fluoride, 0.1 mM N-tosyl-L-phenylalanine chloromethyl ketone, 1 mM benzamidine, 2 mM EDTA, 2 mM EGTA). The applied protein was fractionated using a stepwise NaCl gradient (0-0.3 M at increments of 0.5 mM NaCl per min at a flow rate of 0.8 ml/min and eluted as a single peak centered at 0.11 M NaCl). Selected aliquots were concentrated in a Centricon-10 (Amicon, Bedford, MA) at 0 °C to 3 mg/ml. Electrospray mass spectrometry confirmed the identity of the protein as GST-ATF2-(1-115) with a molecular mass of 39,655 Da.
Protein Kinase Assay of p38--
Protein kinase assays were
conducted at 27 °C in buffer C (50 mM HEPES, pH 7.4, 100 mM KCl, 2 mM dithiothreitol, 0.1 mM
EDTA, and 0.1 mM EGTA), containing 10 nM
p38
, 0.1-50 µM GST-ATF2-(1-115), 0.1-1.0
mM [
-32P]ATP (100-1000 cpm/pmol), 10 µg/ml bovine serum albumin, and 10 mM MgCl2
in a final volume of 50-100 µl. p38
and GST-ATF2-(1-115) were
preincubated for 5 min before the reaction was initiated by the
addition of ATP. Aliquots (5-10 µl) were taken at set time points
and applied to 2 × 2 cm2 P81 cellulose paper (36).
The papers were washed 3 times for 10 min in 50 mM
phosphoric acid (H3PO4) and then in acetone and dried, and the amount of labeled protein was determined by counting the
associated counts/min on a Packard Instrument Co. 1500 scintillation counter at a
value of 2. The (A280 1 mg
ml
1) of the proteins was determined by
denaturing them in 6 M guanidine chloride following the
method of Gill and von Hippel (37).
Product Analysis for the Phosphorylation of GST-ATF2-(1-115) by
p38--
p38
(42 nM) and GST-ATF2-(1-115) (25 µM) were incubated under the standard assay conditions in
buffer C (minus 100 mM KCl) in a 250-µl final volume. The
reaction was stopped by the addition of 250 µl of 0.2 M
EDTA, pH 8.0. The sample was applied to a Centricon-10 and the buffer
exchanged with ammonium carbonate 25 mM, pH 8.6 (300 µl
final volume). The protein was then incubated at 37 °C with 5 µg
of trypsin. After 3 h, another 5 µg of trypsin was added, and
the suspension was left for a further 12 h. After standing on ice
for 10 min, the suspension was centrifuged for 10 min at 13,000 × g. The supernatant containing >95% of the 32P
was removed and fractionated as described below. Fractions containing tryptic peptides were evaporated almost to dryness before the addition
of 0.5 ml of 50% (by volume) water/acetonitrile and analyzed by
electrospray mass spectrometry as described below.
Time Course for the Phosphorylation of GST-ATF2-(1-115) by
p38--
A time course was conducted under the standard assay
conditions with 50 nM p38
and 16.2 µM
GST-ATF2-(1-115) in a 100-µl final volume. Aliquots (10 µl) at
various times were added to hot 2× Laemmli buffer (60 mM
Tris-Cl, pH 6.8, 2% SDS, 10% (by volume) glycerol, 0.025% (by
weight) bromphenol blue) and were resolved by 10% SDS-polyacrylamide
gel electrophoresis. The samples were then transferred to
nitrocellulose membranes, and bands corresponding to GST-ATF2-(1-115)
were identified by staining with Ponceau S (0.1% w/v in 1% v/v acetic
acid). After destaining (3 × 500 ml washes of water), the
membrane was soaked in 0.5% polyvinylpyrrolidone (to prevent trypsin
from binding) and 0.1 M acetic acid for 30 min at 37 °C.
These bands were excised with a razor, shredded, and placed in a 1.5-ml
Eppendorf tube. Samples were washed with water (3 × 1 ml),
followed by 25 mM ammonium carbonate, pH 8.6 (2 × 1 ml). Samples were then incubated with shaking overnight at 37 °C
with 1 µg of sequencing grade trypsin (Roche Molecular Biochemicals)
in 200 µl of 25 mM ammonium carbonate, pH 8.6. The liquid
phase was removed, centrifuged at 13,000 × g for 10 min, and the supernatant placed in a fresh tube and dried on a
Speedvac. Greater than 95% of the counts present on the blot were
routinely recovered. The peptides were resuspended in 10 mM
ammonium acetate, pH 7.0, and applied to a 24 cm × 4 mM Vydac 218TP54 C18 column (Separations
Industries, Hesperia, CA), equilibrated with 10 mM ammonium
acetate, pH 7.0. The column was developed with a linear acetonitrile
gradient (0.25% acetonitrile/min) in 10 mM ammonium acetate, pH 7.0. The flow rate was 0.7 ml/min, and fractions of 0.35 ml
were collected and analyzed for 32P. Typically >95% of
the 32P applied to the column was recovered.
Peptide Microsequencing by Mass Spectrometry-- Phosphorylated peptides were analyzed by LC-MSMS with a Finnigan-MAT LCQ (Finnigan/ThermoQuest, San Jose, CA) electrospray, ion trap mass spectrometer coupled with a Magic 2002 microbore HPLC (Michrom BioResource, Auburn, CA). A 0.5 × 50 mm MAGIC MS C18 column (5 µm particle diameter, 200 Å pore size) with mobile phase A (acetonitrile/water/acetic acid/trifluoroacetic acid. 2:98:0.1:0.02) and phase B (acetonitrile/water/acetic acid/trifluoroacetic acid, 10:90:0.009:0.02) was used with 50% phase B at a flow rate of 20 µl/min. The full scan range in the MS mode was 300-2000 Da. The sequences of phosphorylated peptides were identified using the SEQUEST algorithm incorporated into the Finnigan-MAT BIOWORKS software to correlate the MSMS spectra with amino acid sequences in the OWL protein data base from the National Center for Biotechnology Information. The y and b series ions (38) detected in the secondary ion analysis of peptides were used to identify each peptide.
Pulse-Chase Analysis of the Phosphorylation of
Mono-phosphorylated GST-ATF2-(1-115) by p38--
The pulse-chase
experiment is composed of two parts, the pulse followed by the chase.
Initially, the substrate GST-ATF2-(1-115) (16.2 µM) was
incubated with p38
(24 nM) and
[
32P]ATP (1500 cpm/fmol) (1 µM) in a
volume of 100 µl in the standard kinase assay buffer. After 55 min,
5.4 × 107 cpm was incorporated into the
GST-ATF2-(1-115) corresponding to 1.1% of the theoretical (total)
incorporation into Thr-69 and Thr-71. A 30-µl aliquot was removed for
analysis, and 7 µl of 10.65 mM ATP and p38 MAP kinase
(0.27 µM) was added to give a chase solution (77 µl) of
GST-ATF2-(1-115) (16.2 µM), p38 MAP kinase
(46 nM), 0.97 mM ATP (0.99 cpm/fmol).3 Aliquots (20 µl) at various times were added to hot 2× Laemmli buffer and were
resolved by 10% SDS-polyacrylamide gel electrophoresis. The tryptic
peptide analysis was performed by reverse phase HPLC as above. Control
experiments were performed in the absence of a pulse, using ATP with a
specific activity identical to that in the chase. The maximum
incorporation into GST-ATF2-(1-115) in the control was 2.05 × 106 cpm which corresponds to 3.8% of the pulse.
Calculation of Stimulus Response Curves--
Numerical
integration of the appropriate rate equations was used to calculate the
steady-state concentrations of the species shown in
Scheme 2. Calculations were performed on
a Macintosh Powerbook Computer using KintekSim (Kintek Corp.)
and the PC emulator Virtual PC 3.0 (Connectix). Each step, represented
by an arrow in Scheme 2, was modeled as a two-step reaction
where enzyme and substrate combine (ka) to form a
Michaelis complex, which partitions between dissociation
(kd) and product formation and release
(kcat) (Equation 1). KintekSim has been widely
used in the scientific community to numerically solve complex rate equations and was developed from the original program Kinsim (39).
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(Eq. 1) |
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RESULTS |
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The Phosphorylation of GST-ATF2-(1-115) by p38--
The fusion
protein GST-ATF2-(1-115) was previously shown by LoGrasso et
al. (18) to be a good kinetic substrate for p38
, so we purified
it to homogeneity following their method, with some modifications to
ensure high purity. Once the p38
and GST-ATF2-(1-115) were purified
to homogeneity, we examined the rate and stoichiometry of
GST-ATF2-(1-115) phosphorylation by p38
. Under the conditions of a
standard kinase assay, the phosphorylation of GST-ATF2-(1-115) (16.2 µM) catalyzed by p38
(50 nM) was followed
for 120 min. After 120 min ~1.8 mol of phosphate per mol of
GST-ATF2-(1-115) was incorporated (Fig.
1). A final incorporation of 2.0 mol/mol of phosphate was achieved by incubating the reaction for longer times.
A careful analysis suggested that phosphate incorporation was biphasic,
where to a first approximation, 1.5 mol/mol of phosphate was
incorporated into GST-ATF2-(1-115) with a rate-constant that was
significantly faster than for the remaining 0.5 mol/mol. Steady-state Michaelis parameters were determined by the initial rate method used by
LoGrasso et al. (18). Apparent kcat
and KM(ATF2) parameters were
obtained by measuring the initial rates over the range of 0.5-50
µM GST-ATF2-(1-115) at a fixed concentration of ATP (250 µM). Values for kcat(app) = 0.63 ± 0.1 s
1 and
KM(app) (GST-ATF2-(1-115)) = 3.5 ± 0.5 × 10
6 M
were obtained4 by fitting the
kinetic data (not shown) to the Michaelis-Menten equation by nonlinear
regression according to the method of Cornish-Bowden (40), using the
program Kaleidagraph (3.08d, Synergy Software) for a MacIntosh
computer. These values are in close agreement with the values obtained
by LoGrasso et al. (18).
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Isolation and Identification of Phosphorylated Tryptic
Peptides--
The known p38 phosphorylation sites of ATF2, Thr-69,
and Thr-71 lie within the same tryptic peptide,
60NDSVIVADQTPTPTR74, so
we followed standard protocols to assay for their phosphorylation by
reverse phase HPLC. This approach involved immobilizing phosphorylated GST-ATF2-(1-115) on nitrocellulose membranes, after its fractionation by SDS-polyacrylamide gel electrophoresis, digesting it with high quality sequencing grade trypsin, and fractionating the phosphopeptides by reverse phase HPLC (see "Experimental Procedures"). To examine the phosphorylation of individual residues we conducted the time course
shown in Fig. 1, during the course of which aliquots were taken to
examine the progress of the reaction (Fig.
2, A-C). GST-ATF2-(1-115) was isolated and subjected to tryptic digestion, where the recovery of
the radiolabeled protein and peptides were monitored at each stage of
the analysis. The overall transfer of the protein to the membrane and
subsequent digestion were found to yield >90% of the initial
radiolabel.
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Fig. 2A shows that after 1 min three tryptic peaks, T1-T3,
could be resolved by reverse phase HPLC. Upon mass spectrometric analysis, peak T1 was found to contain a peptide of molecular mass
1774 ± 1 Da, corresponding to residues 60-74 of ATF2,
phosphorylated on two residues. T2 and T3 were found to contain a
peptide of molecular mass 1694 ± 1 Da, which corresponded to a
singly phosphorylated tryptic peptide
60NDSVIVADQTPTPTR74. The tryptic peptide
60NDSVIVADQTPTPTR74 contains four potential
phosphorylation sites as follows: Ser-62, Thr-69, Thr-71, and Thr-73.
Phosphoamino acid analysis (not shown) showed that only threonine
residues were phosphorylated by p38, which ruled out the
phosphorylation of Ser-62. To distinguish between the possible
phosphothreonine isomers, a secondary ion analysis was used to provide
support for the presence of particular phosphorylated peptides. The
y-series of ions, which corresponds to the series of ions generated by
the sequential fragmentation of the M+ or M2+
(38) ion from the C terminus, proved to be diagnostic for the assignment of each peptide (Table I). The
peptide associated with peak T1 corresponded to the peptide,
60NDSVIVADQTPPTPPTR74
(where TP indicates phosphothreonine)
(M = 1773.7 calculated), phosphorylated on Thr-69 and
Thr-71 (Fig. 3). The peptide associated
with peak T2 corresponded to the peptide
60NDSVIVADQTPPTPTR74
(M = 1693.7), mono-phosphorylated on Thr-69. The
peptide associated with peak T3 corresponded to the peptide
60NDSVIVADQTPTPPTR74
(M = 1693.7), phosphorylated on Thr-71. In summary,
both the M+ and M2+ peaks of the expected
molecular mass were observed, and the fragmentation patterns of these
ions are consistent with their assignment. The presence and/or absence
of critical ions, which are summarized in Table I, provide strong
evidence for the unambiguous assignment of each peptide.
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The Kinetics of GST-ATF2-(1-115) Phosphorylation Is Not
Processive--
A primary goal of this work was to establish whether
p38 catalyzed the phosphorylation of GST-ATF2-(1-115) with a
processive (Scheme 3, upper
pathway) or a two-step distributive mechanism (Scheme 3,
lower pathway). ATF2 contains a putative docking sequence similar to the docking sequence termed the docking site for
ERK and JNK, LXL, the DEJL-docking
sequence (3), found in a number of MAPK substrates. This putative
docking sequence is likely to dock on to Asp-313 and Asp-316 of p38
(41), which lies >30 Å outside of the catalytic active site (34). If
this docking hypothesis is correct the DEJL sequence should form a
common binding platform for both phosphorylation events. Thus a
processive mechanism could be favorable, because repositioning of the
threonine residues (Thr-69 and Thr-71) in the active site could be
brought about through minimal reorganization of the intermediate
enzyme-substrate complexes, while contact is maintained at the docking
site.
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With the identity of the peptides in peaks T1-T3 in hand, we were able
to analyze the kinetic mechanism of p38 in more detail. Under
saturating conditions, Thr-69 and Thr-71 were phosphorylated at almost
identical rates (Fig. 2A), despite lying within a different sequence context. The product ratio of an enzymatic reaction is determined by the ratio of the specificity constants
(kcat/KM) under all
conditions (40). Therefore our analysis suggests that p38
is
approximately only 1.2-fold more specific for Thr-71 than for Thr-69 in
the unphosphorylated protein. This is consistent with previous
observations that the specificity of MAP kinases, with respect to the
sequences that surround the phosphorylation sites (SP or TP motifs),
are fairly relaxed (42). By cutting out the peaks and weighing the
paper we were able to determine that the concentration of the
phosphorylated proteins, ATF2
69P, ATF2
71P,
and ATF2
69P/71P, after a 1-min incubation,
were ~21-, 25-, and 3-fold in excess of the enzyme, p38
. This
excess of the mono-phosphorylated substrates is strong evidence that
the mechanism of dual phosphorylation is not processive. It strongly
suggests that the mechanism is a two-step distributive mechanism, where
the intermediary substrates (S*) dissociate from the enzyme faster than
at least one of the following steps: nucleotide exchange, repositioning
of S*, or phosphoryl transfer (Scheme 3) (15).
Pulse-Chase Analysis of the Dual Phosphorylation of
GST-ATF2-(1-115)--
The experiment described above does not
rigorously prove that the formation of the mono-phosphorylated proteins
ATF269P and ATF2
71P
precedes the formation of the dual-phosphorylated product
ATF2
69P/71P. To provide direct evidence that
both ATF2
69P and
ATF2
71P are directly converted to the
dual-phosphorylated product ATF2
69P/71P, a
pulse-chase experiment was performed. In this experiment radiolabeled ATF2
69P and ATF2
71P
were first formed in the pulse phase using [
-32P]ATP
(carrier-free). During the chase phase, incorporation of unlabeled
phosphate into the free site occurs to give labeled ATF2
69P/71P. By monitoring both the
disappearance of the mono-phospho forms of ATF2-(1-115),
ATF2
69P, and
ATF2
71P and the appearance of the
dual-phosphorylated product ATF2
69P/71P, one
can provide evidence for the direct conversion of the
mono-phosphorylated substrates to the dual-phosphorylated product. To
label GST-ATF2-(1-115) in the pulse phase, p38
(24 nM),
GST-ATF2-(1-115) (16.2 µM), and MgATP (1 µM) were combined in a small volume and incubated for 55 min before the ATP was diluted >1500-fold by the addition of 1 mM carrier ATP. The progress of the reaction was then
monitored in the same manner as the time course experiments described
above. During the pulse phase, approximately equal proportions of
ATF2
69P (0.09 µM) and
ATF2
71P (0.09 µM) were formed
as well as a small amount of the doubly labeled
ATF2
69P/71P (less than 0.001 µM) (Fig. 2D). The addition of 1 mM ATP resulted in the fast conversion of
ATF2
71P to
ATF2
69P/71P with a half-life of less than 5 min and the slower conversion of ATF2
69P to
ATF2
69P/71P with a half-life of ~60 min
(see Fig. 2, D-F). The rate of disappearance of peaks
corresponding to ATF2
69P and
ATF2
71P matched the rate of appearance of
label in T1, corresponding to ATF2
69P/71P.
This is consistent with the direct conversion of both
ATF2
69P and ATF2
71P
to the dual-phosphorylated product.
Quantifying the Kinetic Mechanism--
A careful examination of
the products provided insight into the biphasic kinetics shown in Fig.
1. After ~1.5 mol/mol of phosphate was incorporated (20 min, data not
shown) only substrate mono-phosphorylated on Thr-69 remained. No
ATF271P was present, because it had all been
transformed to the dually phosphorylated product
ATF2
69P/71P.
ATF2
69P is considerably less reactive as
witnessed by its presence (~3-4 µM) after 60 min (Fig.
2B). After a more extensive incubation, however, it was
fully converted to the dually phosphorylated product (Fig.
2C). We can quantitatively account for the observations using the mechanism of Scheme 2A (in the absence of
phosphatases), where SP1 and SP2 correspond to
the two mono-phosphoisomers, ATF2
69P and
ATF2
71P, respectively. The line through the
data in Fig. 1 was obtained by using the experimentally observed values
for kcat(app) and KM(app), for all of the steps in Scheme
2A, except for the transformation of
ATF2
69P to
ATF2
69P/71P, whose parameters were allowed to
float. The best fit5 to the
data using Kinteksim gave a
kcat/KM for the
transformation of ATF2
69P to
ATF2
69P/71P that was 40-fold lower than
kcat(app)/KM(app).
This model quantitatively accounts for the progress of phosphate
incorporation into GST-ATF2-(1-115), as well as the absence of
ATF2
71P after 20 min and the presence of 3-4
µM ATF2
69P (calculated 4.5 µM) after 60 min. Product inhibition cannot account for
the difference in the rate of disappearance of the mono-phosphoisomers, because in the pulse-chase experiment (Fig. 2, D and
E) the concentration of phosphorylated product is initially
submicromolar. Furthermore, inhibition of p38
by phosphorylated
products would be expected to have an equal effect on the initial
turnover of both mono-phosphoisomers in the pulse-chase experiment.
The Two-step Distributive Mechanism Allows for Easy Adaptation of
Sensitivity--
The difference in the specificity of p38 for the
mono-phosphorylated intermediates was surprising, and therefore we
examined the possible consequences of such differential specificity on the activation of a substrate by a two-step distributive mechanism in vivo (Scheme 2A). To do this we modeled the
two-step distributive mechanism shown in Scheme 2A and
examined the effect of decreasing the specificity of the enzyme for one
mono-phosphoform of the protein, SP1, by 10-fold. This
conservatively represents the decreased affinity of p38
for
ATF2
69P that we observed experimentally. The
parameters chosen were not chosen to be specific for p38
but were
chosen to be generally representative of an MAPK. For comparison, we
also modeled a processive mechanism (Scheme 2B), which is
not expected to display significant intrinsic ultrasensitivity in the
absence of zero-order effects (12, 13). Initially, we assumed
reasonable values for enzyme and substrate concentrations, taking into
account both total protein concentration and subcellular localization.
Reasonable KM values were assumed based on
experimentally measured values for typical protein substrates.
In the two-step distributive mechanism (Scheme 2A) the dual-phosphorylated product, SP1P2, is formed in two discrete steps. In step 1 the enzyme and substrate combine to give a mono-phosphorylated protein SP1 or SP2. The intermediary enzyme-protein complexes (E·SP1 or E·SP2) then fully dissociate, before rebinding with the enzyme to give the dual-phosphorylated protein (SP1P2). Each phosphorylation step is reversed by a protein phosphatase. In the processive mechanism (Scheme 2B) formation of SP1P2 occurs after just one enzyme-substrate collision (middle pathway) and the protein phosphatases reverse this process by the upper and lower pathways. It is assumed that SP1 and SP2 are substrates of the MAPK. Typical cellular concentrations of MAPKs, for example ERK2, are in the range of 1-2 µM, but because MAPKs are localized within cells it can be reasonably assumed that not all of the MAPK present in a cell are available to phosphorylate any one substrate. Furthermore, the catalytic subunits of protein phosphatase 1 (PP1) and protein phosphatase 2 (PP2) are also targeted to specific subcellular locations through complexation with a variety of other proteins (7, 10). Therefore, assuming a 20-fold lower concentration of the available protein phosphatases, compared with the activated MAPK, we set the phosphatase concentration at 50 nM. MAPKs and protein phosphatases typically have KM values in the 1-10 µM range for bona fide protein substrates, and therefore we used a value of KM = 5 µM for all substrates. Substrate concentration was fixed at 500 nM, corresponding to the approximate concentration of several MAPK substrates, such as the p90rsk isoforms, in mammalian cells. Finally, a separate cycle was introduced (Scheme 2C) to account for competitive substrate inhibition of the MAPK and the phosphatases by other cellular substrates. The concentration of total cellular substrate was estimated to be ~5 µM, based on the assumption that an MAPK has between 20 and 40 substrates at cellular concentrations of 250-500 nM. In all the simulations the signal was interpreted as the concentration of activated MAPK. The concentration of dually phosphorylated product (SP1P2), both free and complexed to protein phosphatases, was used as the signal readout, and the accuracy of the numerical integration was verified by numerically checking the law of conservation of mass. The concentration of magnesium and ATP was assumed to be constant. Values were taken 20 half-lives after the initiation of the input.
The predicted stimulus/response curve for Scheme 2, A and
B, are shown in Fig. 4. The
stimulus/response curve predicted for the processive mechanism (Scheme
2B) and the two-step distributive mechanism (Scheme
2A) are both sigmoidal. Koshland et al.
(13) and more recently Huang and Ferrell (17) have discussed the general shapes of stimulus/response curves resulting from covalent modification of proteins at length. Briefly, the predicted curves are
similar in shape to those given by the Hill equation (y = Xn/(C + Xn) but are
steeper at low stimulus levels and less steep at high stimulus levels.
For comparison, Fig. 4 includes a plot of a signal/response curve for a
Michaelis-Menten enzyme and an ultrasensitive enzyme, where
an 81- and a 9-fold increase in stimulus, respectively, are required to
drive the enzyme response from 10% maximal to 90% maximal. The main
conclusion from these calculations is that the decrease in specificity
of the kinase for SP1 (line D* in Fig. 4)
results in a significant increase in the hyperbolic character of the
simulated signal/response curve (compare curve D*
with curve D). A 17- and 29-fold increase in stimulus (MAPK
activity) is needed to drive the concentration of the
dual-phosphorylated protein, SP1P2, from 10% maximal to
90% maximal for the distributive (line D in Fig. 4) and
processive (line P in Fig. 4) mechanisms, respectively. In
contrast a 58-fold increase in stimulus is needed to drive the same
change in activity for the D* mechanism. A similar relative increase in
the hyperbolic character of the curves (an increase in the control
parameter R (12))6
was observed when the KM of the phosphatase was
varied between 1 and 50 µM or the total substrate
concentration varied from 1.5 to 20 µM. As mentioned
above the ultrasensitivity predicted for the processive mechanism
(R = 29) is due to contributions from zero-order
ultrasensitivity (12, 13). These contributions were estimated by
extrapolating to conditions where the enzymes are not saturated by
substrates (12, 13). Under these conditions (in the absence of a
competitive substrate (Scheme 2C), a processive mechanism is
predicted to be essentially Michaelian (R = 75), a
two-step distributive mechanism ultrasensitive (R = 37), and the D* mechanism subsensitive (R = 115). These
calculations show how the sensitivity through a two-step distributive
mechanism can be modulated to be either ultrasensitive or subsensitive
by changes in the specificity of an enzyme for an intermediate
substrate brought about by phosphorylation.
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DISCUSSION |
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In this study we show that the stress-activated MAPK, p38,
dually phosphorylates the transcription factor ATF2 by a two-step (double collision) distributive mechanism and not by a one-step (single
collision) processive mechanism. Our observation was surprising, because the manner that substrates are recognized by MAPKs appears to
be ideal to promote a processive mechanism of substrate
phosphorylation. There are two possible mechanisms of dual
phosphorylation that can be distinguished by the relative order of
substrate binding (Scheme 3). The upper pathway in Scheme 3 represents
a processive mechanism of phosphorylation, where the protein substrate
does not dissociate from the enzyme until after the second
phosphoryl transfer, k'p, has occurred. The
requirements for this mechanism are fairly stringent because nucleotide
exchange, substrate repositioning, and phosphoryl transfer must
all occur faster than the dissociation of the
mono-phosphorylated intermediate (S*) from the respective ternary or
binary complexes. The lower pathway corresponds to the less stringent,
two-step, distributive pathway, where S* dissociates from the enzyme
after the first phosphorylation, kp, and then
recombines before the second, k'p.
MAPKs are believed to utilize substrate-docking interactions that appear to be well situated to provide a common platform for the phosphorylation of multiple residues within a substrate through a processive mechanism (3, 43, 44). By using a docking site to maintain contact with a substrate and then simply "sliding" the active site from one phosphorylation site to the next, an MAPK could phosphorylate several residues without dissociating fully from the substrate. The catalytic advantage of such a process, over a nonprocessive mechanism, while not expected to be large for the dual phosphorylation of a protein, could be significant within the context of a signal transduction cascade, where the efficient propagation of a signal through the cascade may be critical.7 Interestingly, it has been proposed and also examined theoretically, although not proven experimentally, that scaffold proteins facilitate the processive phosphorylation of proteins in signal transduction cascades (45).
ATF2 contains a putative MAPK docking site that is similar to the DEJL
docking sequence,
(K/R)(K/R)(K/R)X(1-5)(L/I)X(L/I), found in many other MAPK substrates (3). This DEJL-like sequence, 46KHKHEMTL53,
found N-terminal to the phosphorylation sites, Thr-69 and Thr-71, within the trans-activation domain of ATF2, probably
interacts with Asp-313 and Asp-316 of p38 and is predicted to be a
primary specificity determinant (41). Whereas the DEJL-like domain
appears to be ideally situated to promote a processive mechanism, it is clear from our data that this does not happen and that the
mono-phosphorylated GST-ATF2-(1-115) dissociates from p38
with a
rate constant greater than, or equal to 0.6 s
1, the magnitude of GST-ATF2-(1-115)
turnover. Previous studies with p38
uncovered a dependence of
kcat, for the phosphorylation of a peptide, on
viscosity and also a small thiol effect (46). This suggests that
phosphoryl transfer is not rate-limiting for the phosphorylation of a
peptide substrate and that a viscosity-sensitive conformational change
or a product dissociation step is rate-limiting instead. We are
currently investigating the kinetic mechanism of p38
in more detail
to determine the rate-limiting step for the phosphorylation of
GST-ATF2-(1-115). There are several possible reasons why the mechanism
of ATF2 phosphorylation by p38
is not processive. One possibility is
that the putative docking interactions are not the only interactions
required to form a stable p38
-ATF2 complex and that other
interactions within or near the active site of p38
are also
necessary. This is supported by mutagenesis experiments on p38 isoforms
that showed that docking site interactions are not the only
interactions that determine the specificity of the interactions between
p38 isoforms and protein substrates (47). If other interactions are
critical for complex formation and become disrupted after the first
phosphoryl transfer, ATF2 dissociation might quickly ensue, leading to
a distributive mechanism.
It will be interesting to determine whether other MAPKs, which exhibit potentially tighter binding interactions, phosphorylate their substrates with a processive mechanism. There have been several reports that suggest that in some cases MAPKs can bind tightly to docking domains. For example, the binding of the MAPK ERK2 to a glutathione S-transferase fusion protein of the Ets domain transcription factor Elk-1 (GST-Elk1) immobilized on glutathione-agarose beads was shown to be strong enough to survive multiple washes by buffer (48). It was shown that the docking site interactions were both necessary and sufficient to maintain the ERK2-Elk-1 complex on the beads. Similar results were observed for several other MAPK/substrate interactions using a similar approach (49). However, it is possible that the tight binding of MAPK-substrate complexes seen on glutathione-agarose beads are due to nonspecific binding associated with the matrix that increases the affinity of the interactions.
Although p38 showed similar specificity for Thr-71 and Thr-69 in
GST-ATF2-(1-115) (Fig. 2A), it displayed a marked
difference in the specificity toward the mono-phosphoisomers. This
demonstrates that although local sequence is not the primary
determinant of p38
specificity; it can be very important
nevertheless. Cantley and co-workers (42) recently showed that ERK2 has
a preference for a proline at the P-2 position, and therefore it is
possible that phosphorylation of a threonine at this position can
hinder binding of a substrate to p38
.
Interestingly, many MAP kinase substrates appear to be phosphorylated, at least twice by a MAP kinase, before they become activated. Numerous examples of protein conformations regulated by single phosphorylations exist in the literature, so dual phosphorylation is not required per se. Why then is dual phosphorylation so common in MAPK cascades? It has been suggested that dual phosphorylation provides an added check against the inappropriate low rate activation by another protein kinase present in the cell (15). This may be true for the activation of MAPK substrates, because different MAPKs, although similar in both structure and specificity, activate different proteins with different functions in vivo. Another possibility, however, is that multiple phosphorylation mechanisms (by the same enzyme) provide a simple and effective way of influencing the amplitude sensitivity of a pathway. The recent argument by Huang and Ferrell (17) that a two-step distributive mechanism can lead to an increase in ultrasensitivity of a signaling pathway is compelling and could help explain the existence of the exquisite switch-like behavior of a MAPK cascade in Xenopus oocytes. Their laboratory investigated how the variable concentration of the maturation-inducing hormone progesterone, which activates the MAPKKK, Mos, is converted into a switch-like activation of MAPK. They concluded that the observed behavior is consistent with a model where at low concentrations of progesterone the MAPK remains inactive, but upon passing through a narrow concentration threshold of progesterone the MAPK becomes fully activated (50). Sigmoidal stimulus-response curves are important because they provide a way of switching from off to on over a narrow range of input and provide a threshold that can filter out base-line noise.
Computer simulation of our experimentally observed kinetic results
(Fig. 4) shows, however, that a distributive mechanism does not
necessarily result in an increase in ultrasensitivity in
vivo. In fact it appears to offer the potential for considerable variation in the degree of sensitivity. We observed a 40-fold reduction
in the specificity of p38 toward ATF2
71P
in vitro, and we used this as the basis for our
calculations. Rather than model the reaction of p38
specifically, we
examined the effect of decreasing the specificity of a MAPK for one of two intermediates formed as part of a two-step distributive mechanism (Scheme 2A). We assumed that the intermediates were
initially formed in equal amounts from the unphosphorylated substrate
but that the MAPK displays a 10-fold decrease in specificity toward one
of them. The decrease in specificity was predicted to lead to a 3-fold
increase in the range of MAPK activation required to drive the
formation of the product from 10% to 90% maximum, assuming that the
phosphatase did not discriminate between the phosphorylated products
(Fig. 4). This conclusion is qualitatively independent of the level of
the contribution from zero-order ultrasensitivity and illustrates how
the degree of activation of a system in response to a signal could be
adapted by mutations that alter the kinetic parameters of activation.
The kinetics of ATF2 activation by p38 suggests that a further
increase in ultrasensitivity at this level of the pathway is
unnecessary. It will be interesting to determine whether the activation
of p38
displays "switch-like" ultrasensitivity in response to
cell stresses or whether its response is more hyperbolic, where control
is exerted over a wider range of stimulus. It will also be interesting
to compare the activation of other p38
substrates such as the
transcription factors MEF2 (51) and CHOP (52), because in each case,
activation is brought about by the phosphorylation of two sites that
lie close together in primary sequence. It will also be fascinating to
determine how the activation of ATF2 by the other subfamily of
stress-activated MAPKs, the c-Jun N-terminal protein kinases (JNKs),
compares to p38
. Differences in the specificity could reflect
differences in the dependence of ATF2 activation on the activity of the
two subfamilies.
In summary, the dual phosphorylation of protein substrates by a
two-step distributive mechanism appears to offer considerable flexibility in the control of the amplitude-sensitivity of a pathway and is likely to contribute significantly to the signal/response profiles of numerous signal transduction pathways. At one extreme a
distributive mechanism of multiple phosphorylation can increase the
ultrasensitivity of a pathway, as is probably the case for the
activation of MAPK by MAPKK in Xenopus oocytes (17). At the
other extreme the same mechanism could help maintain a broader threshold, allowing the system to respond to a wider range of stimulus.
These factors are likely to be important for the activation of a large
number of enzymes and proteins regulated by MAPK cascades in eukaryotic organisms.
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ACKNOWLEDGEMENTS |
---|
We thank Dr. Melanie Cobb (University of
Texas Southwestern Medical Center) and Dr. Philip LoGrasso (Merck) for
DNA encoding His6-p38 and GST-ATF2-(1-115),
respectively. We also thank Dr. Christian P. Whitman (University of
Texas, Austin) for critically reading the manuscript.
![]() |
FOOTNOTES |
---|
* This work was supported in part by the NIEHS/Center for Research on Environmental Disease, Pilot Project Program, the Analytical Instrumentation Facility Core Grant P30 ES07784, and the Welch Foundation Grant F-1390.The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
To whom correspondence should be addressed: Division of Medicinal
Chemistry, College of Pharmacy, University of Texas, Austin, TX 78712. Tel.: 512-471-9267; Fax: 512-232-2606; E-mail:
Dalby@mail.utexas.edu.
Published, JBC Papers in Press, November 7, 2000, DOI 10.1074/jbc.M008787200
2 Golbeter and Koshland used the term ultrasensitive (and subsensitive) to define any system that requires less (more) than an 81-fold increase in input to drive it from 10% maximal to 90% maximal activation (13).
3
This was calculated by accounting for the ATP
(0.64 µM in 70 µl = 4.48 × 1011 mol) that remained after 55 min of the
pulse phase and the 1516-fold dilution by the cold ATP in the chase.
4 These values are apparent because the assay does not distinguish between the phosphorylation of Thr-69 and Thr-71.
5
Within the limits kcat = 0.1-0.6 s1 and KM = 25-150 µM for this step, little differences in the fit
were seen.
6 The parameter R is the concentration of the stimulus that causes a 90% maximal response divided by the concentration of the stimulus that causes a 10% maximal response.
7 Typically a one-step processive enzymatic mechanism will give 50% product ~2.5-fold faster than a two-step distributive mechanism, assuming comparable rate constants for the binding and chemical steps.
![]() |
ABBREVIATIONS |
---|
The abbreviations used are:
MAPK, mitogen-activated protein kinase;
MAP, mitogen-activated protein;
MAPKK, MAP kinase kinase;
MAPKKK, MAP kinase kinase kinase;
MKK6, mitogen-activated protein kinase kinase 6;
MKK3, mitogen-activated
protein kinase kinase 3;
p38, p38 MAP kinase
;
ATF2, activating
transcription factor 2;
ERK, extra signal regulated protein kinase;
MEKK4, MAP or ERK kinase kinase 4;
HPLC, high pressure liquid
chromatography;
JNK, c-Jun N-terminal protein kinase;
MS, bombarded
mass fragmentation pattern of a molecule;
MS2, bombarded mass
fragmentation pattern of a single m/z ion derived from a MS;
GST, glutathione S-transferase;
ATF269P, ATF271P, and ATF269P/71P correspond to
GST-ATF2-(1-115) phosphorylated on Thr-69, Thr-71, or both Thr-69 and
Thr-71.
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REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1. | Cobb, M. H. (1999) Prog. Biophys. Mol. Biol. 71, 479-500[CrossRef][Medline] [Order article via Infotrieve] |
2. |
Widmann, C.,
Gibson, S.,
Jarpe, M. B.,
and Johnson, G. L.
(1999)
Physiol. Rev.
79,
143-180 |
3. |
Jacobs, D.,
Glossip, D.,
Xing, H.,
Muslin, A. J.,
and Kornfeld, K.
(1999)
Genes Dev.
13,
163-175 |
4. | Keyse, S. M. (2000) Curr. Opin. Cell Biol. 12, 186-192[CrossRef][Medline] [Order article via Infotrieve] |
5. |
Camps, M.,
Nichols, A.,
and Arkinstall, S.
(2000)
FASEB J.
14,
6-16 |
6. | Keyse, S. M. (1999) Free Radic. Res. 31, 341-349[Medline] [Order article via Infotrieve] |
7. | Aggen, J. B., Nairn, A. C., and Chamberlin, R. (2000) Chem. Biol. 7, R13-R23[CrossRef][Medline] [Order article via Infotrieve] |
8. | Virshup, D. M. (2000) Curr. Opin. Cell Biol. 12, 180-185[CrossRef][Medline] [Order article via Infotrieve] |
9. | Millward, T. A., Zolnierowicz, S., and Hemmings, B. A. (1999) Trends Biochem. Sci. 24, 186-191[CrossRef][Medline] [Order article via Infotrieve] |
10. | Goldberg, Y. (1999) Biochem. Pharmacol. 57, 321-328[CrossRef][Medline] [Order article via Infotrieve] |
11. | Ferrell, J. E., Jr. (1996) Trends Biochem. Sci. 21, 460-466[CrossRef][Medline] [Order article via Infotrieve] |
12. | Goldbeter, A., and Koshland, D. E., Jr. (1981) Proc. Natl. Acad. Sci. U. S. A. 78, 6840-6844[Abstract] |
13. | Koshland, D. E., Jr., Goldbeter, A., and Stock, J. B. (1982) Science 217, 220-225[Medline] [Order article via Infotrieve] |
14. | Ferrell, J. E., Jr. (1998) Trends Biochem. Sci. 23, 461-465[CrossRef][Medline] [Order article via Infotrieve] |
15. |
Ferrell, J. E., Jr.,
and Bhatt, R. R.
(1997)
J. Biol. Chem.
272,
19008-19016 |
16. | Burack, W. R., and Sturgill, T. W. (1997) Biochemistry 36, 5929-5933[CrossRef][Medline] [Order article via Infotrieve] |
17. |
Huang, C. Y.,
and Ferrell, J. E., Jr.
(1996)
Proc. Natl. Acad. Sci. U. S. A.
93,
10078-10083 |
18. | LoGrasso, P. V., Frantz, B., Rolando, A. M., O'Keefe, S. J., Hermes, J. D., and O'Neill, E. A. (1997) Biochemistry 36, 10422-10427[CrossRef][Medline] [Order article via Infotrieve] |
19. | Han, J., Lee, J. D., Bibbs, L., and Ulevitch, R. J. (1994) Science 265, 808-811[Medline] [Order article via Infotrieve] |
20. | Lee, J. C., Laydon, J. T., McDonnell, P. C., Gallagher, T. F., Kumar, S., Green, D., McNulty, D., Blumenthal, M. J., Heys, J. R., Landvatter, S. W., et al.. (1994) Nature 372, 739-746[CrossRef][Medline] [Order article via Infotrieve] |
21. |
Han, J.,
Lee, J. D.,
Jiang, Y.,
Li, Z.,
Feng, L.,
and Ulevitch, R. J.
(1996)
J. Biol. Chem.
271,
2886-2891 |
22. |
Cuenda, A.,
Cohen, P.,
Buee-Scherrer, V.,
and Goedert, M.
(1997)
EMBO J.
16,
295-305 |
23. | Porter, A. C., Fanger, G. R., and Vaillancourt, R. R. (1999) Oncogene 18, 7794-7802[CrossRef][Medline] [Order article via Infotrieve] |
24. | Takekawa, M., and Saito, H. (1998) Cell 95, 521-530[Medline] [Order article via Infotrieve] |
25. |
Gerwins, P.,
Blank, J. L.,
and Johnson, G. L.
(1997)
J. Biol. Chem.
272,
8288-8295 |
26. | Tobiume, K., Inage, T., Takeda, K., Enomoto, S., Miyazono, K., and Ichijo, H. (1997) Biochem. Biophys. Res. Commun. 239, 905-910[CrossRef][Medline] [Order article via Infotrieve] |
27. |
Ichijo, H.,
Nishida, E.,
Irie, K.,
ten Dijke, P.,
Saitoh, M.,
Moriguchi, T.,
Takagi, M.,
Matsumoto, K.,
Miyazono, K.,
and Gotoh, Y.
(1997)
Science
275,
90-94 |
28. | Hirose, T., Fujimoto, W., Tamaai, T., Kim, K. H., Matsuura, H., and Jetten, A. M. (1994) Mol. Endocrinol. 8, 1667-1680[Abstract] |
29. |
Moriguchi, T.,
Kuroyanagi, N.,
Yamaguchi, K.,
Gotoh, Y.,
Irie, K.,
Kano, T.,
Shirakabe, K.,
Muro, Y.,
Shibuya, H.,
Matsumoto, K.,
Nishida, E.,
and Hagiwara, M.
(1996)
J. Biol. Chem.
271,
13675-13679 |
30. | Yamaguchi, K., Shirakabe, K., Shibuya, H., Irie, K., Oishi, I., Ueno, N., Taniguchi, T., Nishida, E., and Matsumoto, K. (1995) Science 270, 2008-2011[Abstract] |
31. | Kawasaki, H., Schiltz, L., Chiu, R., Itakura, K., Taira, K., Nakatani, Y., and Yokoyama, K. K. (2000) Nature 405, 195-200[CrossRef][Medline] [Order article via Infotrieve] |
32. | van Dam, H., Wilhelm, D., Herr, I., Steffen, A., Herrlich, P., and Angel, P. (1995) EMBO J. 14, 1798-1811[Abstract] |
33. |
Fuchs, S. Y.,
Tappin, I.,
and Ronai, Z.
(2000)
J. Biol. Chem.
275,
12560-12564 |
34. |
Wang, Z.,
Harkins, P. C.,
Ulevitch, R. J.,
Han, J.,
Cobb, M. H.,
and Goldsmith, E. J.
(1997)
Proc. Natl. Acad. Sci. U. S. A.
94,
2327-2332 |
35. |
Khokhlatchev, A.,
Xu, S.,
English, J.,
Wu, P.,
Schaefer, E.,
and Cobb, M. H.
(1997)
J. Biol. Chem.
272,
11057-11062 |
36. | Kemp, B. E., Benjamini, E., and Krebs, E. G. (1976) Proc. Natl. Acad. Sci. U. S. A. 73, 1038-1042[Abstract] |
37. | Gill, S. C., and von Hippel, P. H. (1989) Anal. Biochem. 182, 319-326[Medline] [Order article via Infotrieve] |
38. | Roepstorff, P., and Fohlman, J. (1984) Biomed. Mass Spectrom. 11, 601[Medline] [Order article via Infotrieve] |
39. | Barshop, B. A., Wrenn, R. F., and Frieden, C. (1983) Anal. Biochem. 130, 134-145[Medline] [Order article via Infotrieve] |
40. | Cornish-Bowden, A. (1995) Fundamentals of Enzyme Kinetics , pp. 297-315, Portland Press Ltd., London |
41. | Tanoue, T., Adachi, M., Moriguchi, T., and Nishida, E. (2000) Nat. Cell Biol. 2, 110-116[CrossRef][Medline] [Order article via Infotrieve] |
42. | Songyang, Z., Blechner, S., Hoagland, N., Hoekstra, M. F., Piwnica-Worms, H., and Cantley, L. C. (1994) Curr. Biol. 4, 973-982[Medline] [Order article via Infotrieve] |
43. | Gavin, A. C., and Nebreda, A. R. (1999) Curr. Biol. 9, 281-284[CrossRef][Medline] [Order article via Infotrieve] |
44. |
Smith, J. A.,
Poteet-Smith, C. E.,
Malarkey, K.,
and Sturgill, T. W.
(1999)
J. Biol. Chem.
274,
2893-2898 |
45. |
Levchenko, A.,
Bruck, J.,
and Sternberg, P. W.
(2000)
Proc. Natl. Acad. Sci. U. S. A.
97,
5818-5823 |
46. | Chen, G., Porter, M. D., Bristol, J. R., Fitzgibbon, M. J., and Pazhanisamy, S. (2000) Biochemistry 39, 2079-2087[CrossRef][Medline] [Order article via Infotrieve] |
47. | Gum, R. J., and Young, P. R. (1999) Biochem. Biophys. Res. Commun. 266, 284-289[CrossRef][Medline] [Order article via Infotrieve] |
48. |
Yang, S. H.,
Yates, P. R.,
Whitmarsh, A. J.,
Davis, R. J.,
and Sharrocks, A. D.
(1998)
Mol. Cell. Biol.
18,
710-720 |
49. | Kallunki, T., Su, B., Tsigelny, I., Sluss, H. K., Derijard, B., Moore, G., Davis, R., and Karin, M. (1994) Genes Dev. 8, 2996-3007[Abstract] |
50. |
Ferrell, J. E., Jr.,
and Machleder, E. M.
(1998)
Science
280,
895-898 |
51. | Wang, X. Z., and Ron, D. (1996) Science 272, 1347-1349[Abstract] |
52. |
Zhao, M.,
New, L.,
Kravchenko, V. V.,
Kato, Y.,
Gram, H.,
di Padova, F.,
Olson, E. N.,
Ulevitch, R. J.,
and Han, J.
(1999)
Mol. Cell. Biol.
19,
21-30 |