Download The regulation of receptor protein tyrosine

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MINI REVIEW P 59-66
The regulation of receptor protein tyrosine
phosphatases by the dimerization of
intracellular domains
Ho-Chul Shin and Seung Jun Kim*
Disease Target Structure Research Center, Korea Research Institute of Bioscience and Biotechnology, Daejeon 34141, Korea
*Correspondence: [email protected]
Protein tyrosine phosphatases (PTPs) are critical in many signal transduction pathways for cell regulation. The activity of
PTPs is governed by interactions between various regulatory domains and their partners. Receptor-type PTPs (PTPRs)
also have many extracellular regulatory domains. The intracellular domain of some PTPRs consists of D1 and D2 domains,
similar to classical PTPs. The D1 domain is an active phosphatase domain, but the D2 domain has weak or no activity. The
D2 domain regulates the phosphatase activity of PTPRs containing the D1–D2 domain. Many studies have shown that the
dimerization of the D2 domain can inhibit the phosphatase activity of PTPRs. A few models have been proposed to explain
how phosphatase activity is inhibited by dimerization, but the precise mechanism is still not established. In this review, we
discuss the regulatory mechanism of the phosphatase activity of PTPRs via the intracellular domain.
INTRODUCTION
Protein phosphorylation is a well-known and common posttranslational modification. Phosphate can form an ester bond on
the hydroxyl group of various amino acids, i.e., serine, threonine,
and tyrosine. The phosphorylated residues carry a large negative
charge on the protein surface. This charge transition on the
surface induces changes in protein–protein interactions, and
these changes by phosphorylation are reversible. Because
phosphorylation is simple and reversible, but results in large
changes, it is used as an “on-off switch” in almost all signaling
pathways in eukaryotes.
Phosphorylation of a protein residue was first observed in
1932 by Fritz A. Lipmann and P. A. Levene. They extracted
phosphoserine from vitellin, a phosphoprotein in egg yolk
(Lipmann and Levene, 1932). Phosphothreonine has been
isolated from bovine casein, a phosphoprotein in milk (De Verdier,
1952). Since phosphoproteins and phosphorylated amino acids
were introduced, they had been considered a good nutrient
source for the young. Kinase and phosphatase, which mediate
protein phosphorylation and dephosphorylation, were discovered
around the same time. In 1946, Daniel L. Harris measured the
dephosphorylation activity of the protein extracted from frog
eggs (Harris, 1946). Kinase activity was measured in 1954 by
George Burnett and Eugene P. Kennedy using proteins in the
mitochondrial membranes of rat liver cells (Burnett and Kennedy,
1954).
Since protein kinases (PKs) were discovered, their importance
in many signal transduction pathways, e.g., cell migration, cell
proliferation, and differentiation, has been established. Although
it was previously thought that the role of protein phosphatases
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(PPs) is just the reverse of PKs, the importance of PPs is now
clearly established as well as PKs (Hunter, 2000; Muratore and
Cole, 2007; Tiganis and Bennett, 2007; Tonks, 2006).
PKs and PPs can be largely classified into three types based on
substrate specificity: serine/threonine-specific, tyrosine-specific,
and dual-specific. Phosphotyrosine regulation is particularly
important in many signaling pathways. Phosphotyrosine
(pY) sites account for a small portion of the human phosphoproteome, i.e., only 1.8%; in contrast, phosphoserine (pS)
and phosphothreonine (pT) sites account for 86.4% and 11.8%
of the phosphoproteome, respectively (Olsen et al., 2006).
Despite this small number of sites, protein tyrosine kinase (PTK)
and protein tyrosine phosphatase (PTP), which are regulatory
proteins for phosphotyrosine, occupy 17.4% (90 PTKs and 518
PKs) and 54.3% (107 PTPs and 197 PPs) of total PKs and PPs in
human cells (Sacco et al., 2012). Moreover, there are nearly equal
numbers of PTKs and PTPs, indicating that the balance between
PTKs and PTPs is important for signaling pathways.
The first PTKs discovered were v-Src and c-Src from
Rous sarcoma virus (RSV) and chickens (Oppermann et al.,
1979; Stehelin et al., 1977). This finding led to the idea that
external materials, like viruses, can promote cancer. Researchers
attempted to find an counteracting enzyme of PTK after the
discovery of Src, and measured the activity of PTPs from various
sources (Chernoff and Li, 1983; Clari et al., 1986; Gallis et al.,
1981; Horlein et al., 1982; Swarup et al., 1982). In 1988, N.
T. Tonks and colleagues first purified PTP and measured the
phosphatase activity of the protein PTP1B (Tonks et al., 1988).
The discoveries of Src and PTP1B promoted research on the
phosphorylation and dephosphorylation of tyrosine residues.
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The regulation of receptor protein tyrosine phosphatases by the dimerization of intracellular domains
The functions of PTKs are well studied, and several drugs
for PTK inhibition have already been developed and used
in clinical settings. However, the functions of PTPs are not
as well-known as those of PTKs are. Drugs targeting PTPs
have just recently been developed, even though many PTPs
are considered important and potential therapeutic targets
for cancer, diabetes, and degenerative brain disease (Das
et al., 2015; Krishnan et al., 2014).
PTPs can be classified into four classes: classes I to IV.
Almost all PTPs belong to class I, and are further divided
into classical PTPs and dual-specificity phosphatases.
Classical PTPs, which commonly have a highly conserved
phosphatase domain, are classified as cytosolic PTPs
and receptor-type PTPs (PTPRs). The cytosolic PTPs
have various regulatory domains or motifs, e.g., SH2,
PDZ, KIMKIS, and the zinc-binding domain. The features
of these cytosolic PTPs are various and they are part of
several signaling pathways. However, PTPRs each have
similar configurations. PTPRs have various regulatory
domains outside of the cell membrane, e.g., the Iglike domain, fibronectin-like domain, and carboxylation
site. These domains can interact with ligands and
environmental stresses, and these interactions can induce
signal transduction in cells. The PTPRs have only one
transmembrane region and one or two phosphatase
domains in the intracellular region after the extracellular
domains. The PTPRs are classified into 8 subtypes
according to their features (Table 1) (Andersen et al., 2001).
Five of 8 subtypes (R1/R6, R2A, R2B, R4 and R5) have
two classical PTP domains in tandem: the membrane
proximal D1 domain and distal D2 domain. The D1 domain
has strong phosphatase activity, but the D2 domain does
not. Despite little or no activity, the D2 domain has a highly
similar amino acid sequence and 3D structure to those of
classical PTPs. Several experimental results in vivo as well
as in vitro and structural analyses have shown that the D2
domain is a regulatory domain for phosphatase activity.
However, the mechanism of phosphatase activity regulation
by the D2 domain is still unclear. Here, we focus on the role
and the mechanism of the regulatory functions of the D2
domain.
TABLE I 1 Classification of PTPRs
Subtype
R1 / R6
Name of PTPR
PTPRC
R2A
PTPRK, PTPRM, PTPRT, PTPRU
R2B
PTPRD, PTPRF, PTPRS
R4
PTPRA, PTPRE
R5
PTPRG, PTPRZ1
R3
PTPRB, PTPRH, PTPRJ, PTPRO,
PTPRP
R7
PTPRR
R8
PTPRN, PTPRN2
D1-D2
in tandem
D1 only
A
B
STRUCTURAL FEATURES OF THE
PHOSPHATASE DOMAIN OF CLASSICAL
PTPS
First, we will describe the chemistry and mechanism of
classical PTPs, which have a highly conserved phosphatase
domain, including the active site and the overall structure.
Classical PTPs have four typical and important motifs:
PTP signature motif, KNRY motif, WPD loop, and
Q-loop (Figure 1A). A cysteine in the PTP signature
motif, (I/V)HCXXGXXR(S/T), also known as a P-loop, is a
catalytic residue, and nitrogen atoms of main chain and an
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FIGURE I 1 Important motifs for phosphatase activity and phosphatase
chemistry. (A) The left panel shows the D1 domain of PTPRS (PDB ID: 2FH7)
and the right panel shows the D1 domain of PTPRC (CD45) and its substrate
peptide (green) (PDB ID: 1YGR). The motifs for phosphatase activity are colored
in the 3D structure. WPD loop: red, KNRY motif: cyan, PTP signature motif: blue,
Q-loop: yellow. Important residues are displayed as sticks. (B) The hydrolysis
mechanism of phosphotyrosine at active sites of classical PTPs.
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Ho-Chul Shin and Seung Jun Kim
arginine of the PTP signature motif can stabilize the phosphate
of phosphotyrosyl substrates. A tyrosine residue in the KNRY
motif has specificity to the phosphotyrosine via a tyrosyl group.
Dephosphorylation proceeds by a two-step displacement
mechanism. The phosphoryl group of a substrate is transferred
to a thiol group as a nucleophile of the catalytic cysteine. An
aspartate in the WPD loop acts as both a general acid and a
base in two catalytic steps. In the next step, glutamines in the
Q-loop help activate a water molecule. The phosphate from
the thiophosphate intermediate is hydrolyzed using this water
molecule (Figure 1B). During dephosphorylation, the WPD loop
shows unique large movements via substrate binding (Figure 1A).
Without the substrate, the loop is located apart from the
active site, but once the substrate is bound to the active
site, the loop moves almost 6 Å and the aspartate of the
WPD loop attaches near the phosphate of phosphotyrosine.
The PTP D1 and D2 domains also have these motifs and the
typical scaffold of classical PTPs.
analysis of the amino acid sequence alignment (Figure 2). The
four motifs of the D1 domain are highly conserved, including
Tyr of the KNRY motif, Arg and Cys of the PTP signature motif,
two Glns of the Q-loop, and Asp of the WPD loop. However, Tyr
of the KNRY motif and Asp of the WPD loop of the D2 domains
are not conserved. Divergence in these motifs resulted in low or
no activity of the D2 domain. The D2 domain of PTPRA is also
inactive and has Asp and Glu, instead of Tyr of the KNRY motif
and Asp of the WPD loop. Substitutions from Asp and Glu to
Tyr and Asp nearly restored the activity levels to those of the D1
domain (Buist et al., 1999; Lim et al., 1998). When these residues
of the D2 domain of PTPRE were substituted to Tyr and Asp,
COMPARISON BETWEEN THE PTP D1 AND
PTP D2 DOMAINS
The available D1 and D2 of the PTPR structures were
superimposed and their RMSDs were calculated (Table
2). These data show a maximum RMSD of 1.951 Å and
minimum of 0.556 Å. The average was 0.830 Å and the
average excluding the row of the D2 domain of PTPRG
was 0.6827 Å. All RMSDs were less than 1.000 Å, except
that of the superimposed D1 domains with the D2 domain
of PTPRG. These low RMSDs indicate that the D1 and D2
domains have almost the same scaffold as classical PTPs.
Abnormally high, but generally low RMSDs between the
D2 domain of PTPRG and D1 domains are due to flexible
loops of the D2 domains of PTPRG, but the overall scaffold
of the D2 domain of PTPRG is also the same as that of the
classical PTPs.
Despite these similarities, there are critical differences
between the D1 domain and D2 domain based on an
FIGURE I 2 The alignment of amino acid sequences among PTPRs
containing D1 and D2 domains. ClustalX2 (Larkin et al., 2007) was used to
align the amino acid sequences of 12 PTPRs containing D1 and D2 domains.
Blue indicates the level of conservation. The residues in red boxes are important
for PTP activity, displayed as sticks in Figure 1A.
TABLE I 2 RMSDs among D1 domains and D2 domains of PTPRs with solved structures
D1 domain
D2 domain
A
E
F
G
S
A
E
F
G
S
K
M
T
Z1
0.780*
(195)#
0.704
(200)
0.577
(195)
1.246
(142)
0.575
(205)
0.709
(189)
0.642
(208)
0.556
(198)
1.309
(151)
0.557
(206)
0.853
(197)
0.930
(215)
0.688
(205)
1.344
(166)
0.602
(203)
0.727
(190)
0.778
(211)
0.763
(217)
1.951
(185)
0.831
(226)
0.806
(187)
0.836
(193)
0.668
(216)
1.235
(166)
0.657
(207)
0.684
(190)
0.660
(202)
0.580
(182)
1.642
(181)
0.561
(181)
0.701
(194)
0.693
(184)
0.653
(204)
1.177
(166)
0.599
(204)
0.634
(192)
0.570
(204)
0.566
(266)
1.170
(158)
0.567
(205)
0.723
(186)
0.792
(213)
0.752
(218)
1.679
(171)
0.602
(195)
*RMSDs (Å ) were calculated using PyMol (Schrodinger, 2010) #Numbers of superimposed amino acids.
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The regulation of receptor protein tyrosine phosphatases by the dimerization of intracellular domains
phosphatase activity of the D2 domain of PTPRE also increased,
but less than the activity of D1 domain. PTPRC (CD45) has an
additional substitution from Arg to Gln in the PTP signature motif.
When these three residues were replaced with the conserved
residues, phosphatase activity of the mutant D2 domain
increased slightly (Lim et al., 1999).
Divergence at these critical residues for catalytic activity leads
to the inactivation of the D2 domain, despite the similarity in the
3D structural scaffolds of the D1 domain and D2 domain. PTPRs
have a D2 domain, despite a lack of catalytic activity because the
domain can play a role in the regulation of phosphatase activity
in the intracellular domain.
A
RECEPTOR-TYPE PTP REGULATION VIA
D2 DOMAIN DIMERIZATION
B
The PTPRs can be dimerized or oligomerized via ligand binding
on extracellular domains (Fukada et al., 2006; Perez-Pinera
et al., 2007). This interaction can change the structure of the
intracellular domain and inactivate phosphatase activity. Several
studies have proven that the dimerization of the intracellular
domains of PTPR can decrease phosphatase activity.
A homodimer of the D2 domain of PTPRE has been detected
by immunoprecipitation, implying that the D2 domain can form a
dimer in vivo (Toledano-Katchalski et al., 2003). The phosphatase
activity of PTPRC is inhibited when the dimerization of a chimeric
protein, in which the extracellular domain and transmembrane
domain of PTPRC are replaced with EGFR (epithermal growth
factor receptor), is induced by EGF (epithermal growth factor)
(Desai et al., 1993). The chimeric protein of PTPRZ in which
FKBP is inserted between the transmembrane domain and D1
domain is dimerized by AP20187. This artificial dimerization
decreases the phosphatase activity of PTPRZ (Fukada et al.,
2006). The dimerization of the D2 domain, irrespective of whether
it is related to features of the extracellular domains or internal
features of the intracellular domain, can inhibit the phosphatase
activity of PTPRs.
In 1996, the structure of the D1 domain of PTPRA was
discovered. The structure displayed an antiparallel dimer of
two PTPRA D1 domains and a unique structure was observed
with respect to the location of the N-terminal wedge (N-terminal
helix-turn-helix) of the D1 domain, which blocks the active site
of another D1 molecule (Figure 3A). Furthermore, these wedge
regions of D1 domains are generally conserved at the amino acid
level (Figure 3B).
Researchers have suggested the wedge inhibition model, in
which the wedge domain can block the active site and inhibit the
phosphatase activity of the D1 domain after dimerization (Bilwes
et al., 1996).
In 1999, the structure of the intracellular domain of PTPRF
containing D1 and D2 domains was determined (Nam et al.,
1999). However, there was no blockage of the active site by a
wedge and accordingly the active site was still accessible in the
structure. After the discovery of this structure, no blockage of the
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C
FIGURE 3 I The dimer structure of PTPRA and its wedge region.
(A) The dimer structure of PTPRA (PDB ID: 1YFO). The wedge regions
are indicated by a blue line. The red circles are the target residues for
constitutive activation. Translucent red circles show the active sites of PTP.
(B) The amino acid sequences of the wedge region of PTPRs are aligned.
The red box shows the target residue for constitutive activation, labeled
by a red circle in (A). (C) Two PTPRFs (PDB ID: 1LAR) are superimposed
on the dimer of the PTPRA D1 domain on the right panel. The yellow box
displays a steric clash between two D2 domains of the superimposed
PTPRFs.
active site by a wedge region was detected in the structure of
the D1 domains and D1-D2 domains. Moreover, superimposing
two D1-D2 domains of PTPRF onto two D1 domains of PTPRA
suggested a wedge model with steric clashes between the D2
domains (Figure 3C) (Barr et al., 2009). The orientation of the
D1 and D2 domains is highly conserved (Figure 4A) and a linker
between D1 and D2 is too short and too tightly attached on the
N-terminus of the D2 domain, making a short beta-strand to
detach the D2 domain from the D1 domain (Barr et al., 2009).
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Ho-Chul Shin and Seung Jun Kim
Additionally, the linker region between the D1 and D2 domain is
highly conserved (GXTX(I/V)) (Barr et al., 2009). These analyses
suggested that the conformational changes between D1 and D2
are unlikely. In addition, the highly conserved orientation of the
D1-D2 domains suggests that the D1 and D2 domains interact
(Figure 4A). The residues at the interface between the D1 and
D2 domains are generally conserved (Figure 4B). Moreover,
∆iGs at the interface between the D1 and D2 domains of the
structure of PTPRs are sufficient to interact with each
other (for ∆ iG calculated using PISA (Krissinel and Henrick,
2007)), indicating that the interaction between D1 and D2 is an
important feature (Figure 4C). Based on these problems, Stefan
Knapp and colleagues suggested the inhibition model with a
head-to-toe arrangement, rather than a wedge inhibition model
(Barr et al., 2009). In the structure of PTPRG, the active site of
the D1 domain is blocked by the D2 domain of another PTPRG
in a head-to-toe arrangement (Barr et al., 2009). Their schematic
model suggested that this orientation, like the PTPRG structure,
could be formed under the membrane and could block the active
site (Barr et al., 2009).
However, from a different perspective, these conserved
features indicate the possibility of mechanical conformational
changes of the D1-D2 domain by dimerization to inhibit
phosphatase activity. Interactions between D1 and D2 are not
strong enough to stabilize the D1-D2 orientation. This means
that some forces or changes in D1-D2 can induce changes in the
D1-D2 orientation, and a steric clash between the D2 domains
can be avoided. In fact, there is evidence supporting the
wedge inhibition model in vivo (Tertoolen et al., 2001; ToledanoKatchalski et al., 2003). A mutation from Glu to Arg in the
wedge region constitutively activates the phosphatase activity
of PTPRC (CD45) (Majeti et al., 2000). This residue is in close
proximity to the active site in the dimer structure of the PTPRA
D1 domain, and a substitution of an arginine residue from
a glutamate causes repulsion at the active site via a charge
inversion and a structural clash owing to a large guanine group
(Figure 3A, B). Phosphatase activation by the ER mutant on the
wedge region of PTPRC decreases leukocyte recruitment and
drives autoimmune diseases (Germena et al., 2015; Hermiston
et al., 2005; Zikherman et al., 2013). Moreover, the activation
of PTPRS with this mutation limits neuronal regeneration.
Recently, several researches show the wedge peptide of PTPRs
can inhibit the phosphatase activity of PTPRs. Proliferation
of PC12 cells is increased by treatment of wedge peptide
of PTPRF and the wedge peptide of PTPRM blocks outgrowth
of retinal neurite (Xie et al., 2006). Jerry Silver and colleagues
have shown that the wedge peptide of PTPRS causes a corn
growth of neurons (Lang et al., 2015). This peptide also promotes
regeneration of sympathetic axon in the cardiac scar by inhibition
of PTPRS in vivo. R. T. Gardner and colleagues showed the
possibility that wedge peptide of PTPRS could be used clinically
to inhibit the activity of PTPRS (Gardner et al., 2015).
Although dimerization of the intracellular domain, which may
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A
B
C
FIGURE 4 I Conservation between the D1 domain and D2 domain. (A)
The four PTPRs are superimposed to compare the orientations of the D1
and D2 domains. The conserved residues in the amino acid sequence
alignments are presented in the inset. Blue areas indicate the level of
conservation. (B) The interfaces between the D1 domain and the D2
domain are presented as a surface model. Blue areas indicate the level of
conservation, similar to (A). The area in the red line is the contact region of
the D1 domain and the D2 domain. (C) Estimated ∆iG and interface areas
are shown for the structures for which the intracellular domain of PTPR are
solved.
be induced by ligand binding, can decrease the phosphatase
activity of PTPRs, the inhibitory mechanism of phosphatase
activity by dimerization is still unclear.
RECEPTOR-TYPE PTP REGULATION BY
OXIDATION
Another potential inhibitory mechanism is the reversible oxidation
of the intracellular domain. Oxidation is a well-known regulatory
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The regulation of receptor protein tyrosine phosphatases by the dimerization of intracellular domains
mechanism for enzymes that have a cysteine as the catalytic
residue, and most PTPs have a cysteine as a catalytic residue.
It had been reported that exogenous H2O2 stimulates the insulin
receptor, which is a PTK, and increases the phosphorylated
tyrosine levels of its substrates (Heffetz et al., 1990; Heffetz
and Zick, 1989). These reports suggested that the inhibition
of PTP activity may be mediated by oxidation. PTP is oxidized
by exogenous H 2O 2 directly and reduced by GSH reversibly
in vitro (Hecht and Zick, 1992). H2O2 increases endogenously
via platelet-derived growth factor (PDGF) (Sundaresan et al.,
1995). These results demonstrate the role of H2O2 as a signal
transduction molecule. Subsequently, Rhee and colleagues
discovered that the phosphatase activity of PTP1B decreases
in response to endogenous H 2O 2 induced by EGF and this
oxidation is reversible (Lee et al., 1998). Since these discoveries,
signal transduction related to reversible oxidation of PTPs
has been confirmed by several analyses, including structural
evidence (Buhrman et al., 2005; Jeon et al., 2013; Lee et al.,
2015; Salmeen et al., 2003; van Montfort et al., 2003).
The reversible redox regulation occurs at a catalytic cysteine.
This residue can be easily oxidized because the thiol group of
cysteine, as a catalytic residue, is highly reactive in the cytosolic
environment. The thiol group of cysteine can have several
oxidized forms. In general, the thiol group is able to change
sequentially to sulfenate, sulfinate, and sulfonate by oxidation.
If another cysteine is located near the target cysteine, they
can form a disulfide bond by oxidation. In special cases, the
catalytic residue can make a sulfenyl-amide bond with a nitrogen
atom between the thiol group in cysteine and the main chain
nitrogen atom in the following residue (Figure 5). The sulfenic
acid, disulfide, and sulfenyl-amide intermediate can be reversibly
converted to a thiol group (Figure 5). These reversible oxidations
are the key step for controllable redox regulatory signaling.
PTPRs have this reversible redox regulation pathway because
PTPRs also have cysteine as a catalytic residue. The various
oxidized D2 domain structures of PTPRA have been solved and
analyzed (Yang et al., 2007). One of the PTPRA structures has
a sulfenyl- amide bond at Cys723 and Ser724, indicating that
PTPRA can be regulated by reversible redox signaling.
The redox regulation of PTPRs involves not only the direct
inhibition of the catalytic cysteine by oxidation, but also two
unique characteristics. First, the D2 dimerization is increased
by oxidation of the intracellular domain. According to previous
studies, the extent of homodimer formation of the intracellular
domains of PTPRA and PTPRE was increased by H 2 O 2
(Blanchetot et al., 2002; Toledano-Katchalski et al., 2003).
Second, the D2 domain is more sensitive to oxidation than the
D1 domain (Groen et al., 2008; Persson et al., 2004). These two
unique characteristics could be closely related to phosphatase
activity regulation by the dimerization of D2 domains. According
to limited proteolysis results for PTPRA, PTPRM, PTPRC,
and PTPRS, it is possible that conformational changes of the
intracellular domain can be induced by oxidation (Groen et al.,
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FIGURE 5 I The oxidation mechanism of a thiol group of a cysteine
residue. Oxidation starts with a thiol group. ROS is a reactive oxygen
species and RSH is a reductant containing a thiol group.
2008), and it is possible that these conformational changes
induce the dimerization of the two intracellular domains.
However, based on the recently solved structure of the
oxidized intracellular domain of PTPRS, only the D1 domain
is oxidized and there is no significant conformational change
compared with the previously solved PTPRS (RMSD 0.197
Å). Analyses of the oxidized intracellular domain of PTPRS by
using mass spectrometry have revealed that only D1 is oxidized
(Jeon et al., 2013). Despite evidence for the induction of D2
dimerization by oxidation, the mechanism is still unclear and
controversial.
A SCHEMATIC REGULATORY MODEL FOR PTPR
The inhibition of phosphatase activity by the dimerization of D2
domains is clearly established, but the mechanism is not yet
known. The dimerization of D2 domains can be induced by two
mechanisms. First, it is possible that the intracellular domains
come close to each other via the dimer- or oligomerization
of the extracellular domain. This could induce mechanical
conformational changes between the two intracellular domains,
potentially via a steric clash. Alternatively, the oxidation of
the D2 domain could induce conformational changes at the
interface between the D1 and D2 domains. These two possible
conformational changes could facilitate the dimerization of the
D2 domains. After D2 domains form a dimer, the active site of the
D1 domain could be blocked by the wedge region or by head-
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Ho-Chul Shin and Seung Jun Kim
to-toe dimer formation. There is no clear and direct evidence to
support these mechanisms. Additional data, e.g., structural and
biochemical data, are necessary to establish the mechanism of
phosphatase inhibition by intracellular domains.
ACKNOWLEDGEMENTS
This work was supported by the Bio & Medical Technology Development
Programs of the National Research Foundation, funded by the Korean
Government (2011-0030027) and the Korea Research Institute of
Bioscience and Biotechnology Research Initiative Program.
AUTHOR INFORMATION The authors declare no potential conflicts of interest.
Original Submission: May 20, 2016
Revised Version Received: Jun 13, 2016
Accepted: Jun 14, 2016
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