Mol Cell Biol, May 1998, p. 2608-2616, Vol. 18, No. 5
0270-7306/98/$04.00+0
Copyright © 1998, American Society for Microbiology. All rights reserved.
(PTP
) Binds to and Inhibits the First Catalytic Domain of
PTP
Division of Respiratory Research, The Hospital for Sick Children, Toronto, Ontario M5G 1X8, and Department of Biochemistry, University of Toronto, Toronto, Ontario M5S 1A8, Canada
Received 23 June 1997/Returned for modification 12 August 1997/Accepted 19 February 1998
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ABSTRACT |
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The LAR family protein tyrosine phosphatases (PTPs), including LAR,
PTP
, and PTP
, are transmembrane proteins composed of a cell
adhesion molecule-like ectodomain and two cytoplasmic catalytic domains: active D1 and inactive D2. We performed a yeast two-hybrid screen with the first catalytic domain of PTP
(PTP
-D1) as
bait to identify interacting regulatory proteins. Using this screen, we
identified the second catalytic domain of PTP
(PTP
-D2) as an interactor of PTP
-D1. Both yeast
two-hybrid binding assays and coprecipitation from
mammalian cells revealed strong binding between PTP
-D1 and
PTP
-D2, an association which required the presence of the wedge
sequence in PTP
-D1, a sequence recently shown to mediate D1-D1
homodimerization in the phosphatase RPTP
. This interaction was not
reciprocal, as PTP
-D1 did not bind PTP
-D2. Addition of a
glutathione S-transferase (GST)-PTP
-D2 fusion
protein (but not GST alone) to GST-PTP
-D1 led to ~50%
inhibition of the catalytic activity of PTP
-D1, as determined by
an in vitro phosphatase assay against
p-nitrophenylphosphate. A similar inhibition of PTP
-D1 activity was obtained with coimmunoprecipitated
PTP
-D2. Interestingly, the second catalytic domains of LAR
(LAR-D2) and PTP
(PTP
-D2), very similar in sequence to
PTP
-D2, bound poorly to PTP
-D1. PTP
-D1 and LAR-D1
were also able to bind PTP
-D2, but more weakly than
PTP
-D1, with a binding hierarchy of
PTP
-D1>>PTP
-D1>LAR-D1. These results suggest that
association between PTP
-D1 and PTP
-D2, possibly via
receptor heterodimerization, provides a negative regulatory
function and that the second catalytic domains of this and likely
other receptor PTPs, which are often inactive, may function instead
to regulate the activity of the first catalytic domains.
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INTRODUCTION |
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Tyrosine phosphorylation, controlled
by the activity of protein tyrosine kinases (PTKs) and protein tyrosine
phosphatases (PTPs), plays a critical role in the regulation of many
cellular processes, including cell proliferation and
differentiation. PTPs, like PTKs, can be classified into cytosolic and
receptor-type PTPs (11, 25). One subclass of receptor PTPs
(RPTPs) is represented by the LAR family of phosphatases, which
includes LAR and Drosophila DLAR (37, 39), PTP
(20, 23, 29), PTP
(also known as LAR-PTP2, PTP-P1,
CRYP
, PTP-NU3, PTP-NE3, and CPTP1 [27, 30, 34, 44, 45,
49, 52] and referred to herein as PTP
), and the three
related phosphatases PTP
, PTPµ, and PTP
(6, 12, 16).
These PTPs are characterized by an extracellular domain composed of
multiple immunoglobulin (Ig)-like and fibronectin type III (FNIII)
repeats, resembling cell adhesion molecules (CAM) such as N-CAM and L1
(7, 24) and several receptor PTKs. The CAM-like ectodomain
can also be expressed alone, due to either alternative splicing or
ectodomain shedding, thus disconnecting it from the intracellular
catalytic domains (16, 26, 36). Like most RPTPs, the LAR
family phosphatases contain a single transmembrane domain and two
tandemly repeated catalytic domains (D1 and D2). Mutation of the highly
conserved Cys in LAR-D1 abrogates PTP catalytic activity, suggesting
that D2 is inactive, as also demonstrated for CD45 (28, 38).
These results were also supported by direct measurements of the
catalytic activity of LAR-D1 and -D2, or PTP
-D1 and -D2, against
several artificial substrates (11a, 15, 44). Based on these
findings and the observation that in PTP
and RPTP
, the highly
conserved Cys in the second catalytic domain is replaced by Asp
(2, 19), it has been proposed that the second catalytic
domains of most RPTPs may have a regulatory rather than a catalytic
function or, alternatively, that the second catalytic domains have a
different substrate specificity than the first catalytic domains.
Dimerization of receptor PTKs has long been recognized as an essential
step in their autophosphorylation and activation (42). Recently, homodimerization of a tyrosine phosphatase, RPTP
, was demonstrated (3). Determination of the crystal structure of RPTP
has revealed that the first catalytic domain (D1) dimerizes, to
form a D1-D1 complex. This dimerization occurs by insertion of a
"wedge" sequence, located at the N terminus of each D1 and conforming to a helix-turn-helix structure, into the active site of the
partner D1 (3). Based on this dimeric structure, it was
proposed that a D1-D1 dimeric complex would inhibit catalytic activity.
Indeed, an epidermal growth factor receptor-CD45 chimera in which the
ecto- and transmembrane domain of the EGF receptor was linked to the
intracellular catalytic domains of CD45, was previously shown to
dimerize in response to epidermal growth factor and to inhibit
CD45-mediated T-cell activation, which requires intact catalytic
activity of CD45-D1 (9, 10). Thus, homodimerization or,
possibly, heterodimerization may provide a mechanism to regulate the
function of PTPs.
In this report, we describe the isolation of the second catalytic
domain of PTP
(PTP
-D2) in a two-hybrid screen using the first
catalytic domain of PTP
(PTP
-D1) as bait. Moreover, we show
strong interactions between these two domains in both two-hybrid binding assays and coprecipitations in mammalian cells, an interaction which requires the presence of the wedge region of PTP
-D1 and which leads to partial inhibition of the catalytic activity of PTP
.
The first catalytic domains of PTP
and LAR bind more weakly than
PTP
-D1, to PTP
-D2 and the binding of the D1 proteins of all
three LAR family members appears to be specific to PTP
-D2. We
thus propose that the association between the first and second catalytic domains of LAR family members, particularly
PTP
-D1 and PTP
-D2, may provide a negative regulatory
function and that the second catalytic domain of PTP
and, possibly,
those of other RPTPs, which are usually inactive, function instead to
regulate the activity of the first catalytic domains.
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MATERIALS AND METHODS |
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Yeast two-hybrid library screens.
A PCR fragment
encompassing nucleotides (nt) 3896 to 5002 (amino acids [aa] 1238 to
1606) that encodes the first catalytic domain (D1) of PTP
(rat
LAR-PTP2; accession no. L11587; reference 52) with a
C
S point mutation (M) in the signature motif (C1504S) and includes
a sequence encoding the HA epitope was subcloned into the LexA DNA
binding domain fusion vector pBTM116 (43); such a C
S
point mutation in the catalytic core of PTPs (including PTP
)
abolishes catalytic activity but still allows substrate binding
(40). The insert-containing plasmid [called
pBTM116HA
D1(M)] was used to transform Saccharomyces
cerevisiae L40 (MATa his3 LYS2:LexA-His3
URA3::LexA-lacZ) by the standard Li acetate method to
give Trp prototrophs. Transformants were tested for expression of the
LexA-PTP
-D1 fusion protein by immunoblotting using an anti-HA
antibody (Boehringer Mannheim). The L40 cells transformed with
pBTM116HA
D1(M) were cotransformed with either an adult rat lung cDNA
library or an 11-day mouse embryo library (Matchmaker; Clontech)
constructed in the pGAD10 (Gal4 activation domain) plasmid and selected
on medium lacking Trp, Leu, and His and containing 0 to 20 mM
3-aminotriazole. Plates were incubated for 5 days and
overlaid with replica filters, and cells were permeabilized by freezing
filters in liquid nitrogen and then thawing them at room
temperature. Filters were transferred onto Whatman 3MM paper saturated
with an X-Gal
(5-bromo-4-chloro-3-indolyl-
-D-galactopyranoside) solution and incubated at 30°C to monitor color development.
-Galactosidase (
-gal)-positive colonies were selected and
streaked on medium lacking Trp and Leu for further analysis. Total DNA
was extracted from these colonies, used to transform Escherichia
coli HB101. The bacteria were then plated on M9 minimal plates
or subjected to PCR with pGAD-specific primers, and religated into
pGAD10. Unique inserts were identified by sequencing. To test for true positives, the unique inserts were transformed into L40 cells either
alone, with pBTM116HA
D1(M), or with an unrelated pBTM116 construct.
Cloning of PTP
-D1.
The first catalytic domain of
PTP
(PTP
-D1), including its wedge sequence, was cloned from a
mouse brain cDNA library (Marathon cDNA library; Clontech) by PCR using
the 5' oligonucleotide AAAAGGAAGAGGGCAGAGTCGGACTCC and the
3' oligonucleotide TTTTGAGCTGGCTAGACGCTTAAATTC as primers.
Constructs and yeast two-hybrid binding assays.
PCR
fragments encompassing the first catalytic domain (D1) of PTP
(nt
3896 to 5002, aa 1238 to 1606), PTP
(nt 2494 to 3582, aa 673 to
1035), and LAR (nt 3846 to 4928, aa 1276 to 1636) or the first
catalytic domain of PTP
lacking its wedge sequence (nt 4138 to 5002, aa 1318 to 1606) were subcloned into the LexA DNA binding domain fusion
vector pBTM116. The second catalytic domains (D2) of PTP
(nt 5003 to
5776, aa 1607 to 1863), PTP
(nt 3577 to 4353, aa 1034 to 1291), and
LAR (nt 4938 to 5717, aa 1640 to 1898) were subcloned into the pGAD10
(Gal4 activation domain) plasmid. The same D2 fragments were also FLAG
tagged at their 3' termini and subcloned into pACT2 (Gal4 activation
domain); this plasmid has a stronger promoter than pGAD10, thus
allowing immunodetection of the expressed proteins. L40 cells were
transformed with these D2 constructs either alone or in combination
with the D1 constructs and grown on medium lacking Trp for the pBTM116 transformants, lacking Leu for the pGAD10 or pACT2 transformants, or
lacking Trp and Leu for the double transformants. Individual colonies
were streaked onto fresh medium for filter
-gal assays as described
above.
-gal assays were performed in accordance with the
manufacturer's (Clontech) instructions. Briefly, individual yeast transformant colonies were grown in 20 ml of selective medium at 30°C
until the cultures reached an optical density at 600 nm of ~1.3. An
aliquot (0.1 ml) of each culture was lysed and incubated with a
0.6-mg/ml o-nitrophenyl-
-D-galactopyranoside
solution at 30°C for 10 min. The reactions were then quenched, and
the absorbance of the supernatant was measured at 420 nm to quantify the release of o-nitrophenol. The same cultures used for the
-gal assays were analyzed for protein expression levels by
immunoblotting using an anti-HA antibody (Boehringer Mannheim) for the
LexA-D1 fusion proteins or an anti-FLAG antibody (IBI) for the
FLAG-tagged Gal4-D2 fusion proteins.
Preparation of GST fusion proteins in bacteria.
All
glutathione S-transferase (GST) fusion proteins were
prepared by PCR amplification of the appropriate regions of PTP
or
PTP
and subcloning into pGEX-KG, pGEX-4T1, or pGEX-4T2. The insert-containing plasmids were transformed into E. coli
HB101. Expression of fusion proteins was induced with 0.1 mM
isopropyl-
-D-thiogalactopyranoside, and bacteria were
collected and lysed in lysozyme buffer containing 33 mM Tris-HCl (pH
7.4), 2.5 mM EDTA, 10 mM
-mercaptoethanol, 1-mg/ml lysozyme, and
protease inhibitors (1 mM phenylmethylsulfonyl fluoride, 10-µg/ml
aprotinin, and 10-µg/ml leupeptin) by sonication. The lysate was
treated with MgCl2 and DNase I at final concentrations of 2 mM and 25 ng/ml, respectively. After 20 min of incubation at 25°C,
EDTA was added to a 4 mM concentration and Triton X-100 was added to
1% and incubation proceeded for an additional 10 min at 25°C. The
resulting lysate was cleared by centrifugation at 10,000 × g for 10 min (4°C), and the supernatant was incubated with
glutathione-agarose beads. The pellet was treated with 1.5% (wt/vol)
N-lauroylsarcosine-25 mM triethanolamine-1 mM EDTA (pH 8.0) and incubated for 15 min at 4°C. CaCl2 was added to
a 1 mM final concentration, and then the solubilized pellet was cleared by centrifugation at 10,000 × g for 10 min. The
supernatant was collected and pooled with the previous supernatant and
incubated with glutathione-agarose beads. The proteins were then eluted with 30 mM reduced glutathione (pH 8.0). This extensive purification procedure of the GST fusion proteins was necessary because the proteins
produced in bacteria were largely insoluble.
Transfections in mammalian cells and coprecipitations.
PCR-generated fragments of PTP
corresponding to D1 or to D1 missing
the N-terminal wedge sequence (see above and Fig. 1A) were subcloned
into the mammalian expression vector pEBG to generate GST-PTP
-D1 and the GST-PTP
D1-W construct with the wedge
deleted, respectively. GST-PTP
-D1 and GST-LAR-D1 were
generated in pEBG in a similar fashion. The second catalytic domains
(D2) of PTP
, PTP
, and LAR were generated by PCR (as described
above) with HA tags and subcloned into the pCMV4 mammalian expression
vector. Insert-containing plasmids were transiently transfected (alone or in combination) into Cos7 cells in six-well plates by using Lipofectamine (Gibco). Transfected cells were lysed in 200 µl of
lysis buffer plus protease inhibitors (50 mM HEPES [pH 7.5], 150 mM
NaCl, 1.5 mM MgCl2, 1 mM EGTA, 10% glycerol, 1% Triton X-100, 1 mM phenylmethylsulfonyl fluoride, 10-µg/ml leupeptin, and 10-µg/ml aprotinin) per well; 20-µl aliquots were taken to verify the protein expression of each construct, and the remaining lysate was incubated with 25 to 30 µl of a 50% glutathione-agarose slurry for 1 h at 4°C. Beads were then washed four times with high-salt HNTG (20 mM HEPES [pH 7.5], 500 mM NaCl, 0.1% Triton X-100, 10% glycerol), and proteins were separated by sodium dodecyl sulfate (SDS)-10% polyacrylamide gel electrophoresis (PAGE),
transferred to nitrocellulose, and immunoblotted with either anti-GST
antibodies to detect the precipitated D1 domains of PTP
, PTP
, or
LAR or anti-HA antibodies to detect the coprecipitated D2
domains of these PTPs. In parallel sets of experiments, cells
transfected with GST-PTP
-D1, alone or together with
HA-PTP
-D2, were lysed and the lysate was incubated with
glutathione agarose beads to precipitate GST-PTP
-D1. The
precipitated PTP
-D1 was then analyzed for catalytic
(phosphatase) activity.
Phosphatase assays.
PTP activity of GST-PTP
-D1
precipitated from Cos7 cells, or generated in bacteria, was assayed by
using p-nitrophenylphosphate (PNPP) as a substrate. Assays
were performed at room temperature in 50 to 100 µl of reaction buffer
containing 100 mM PNPP, 100 mM
2-(N-morpholino)ethanesulfonic acid (MES) at pH 5.5, 10 mM dithiothreitol, 150 mM NaCl, and 2 mM EDTA. For assays performed on GST fusion proteins generated in bacteria, 200 to 500 ng of GST-PTP
-D1 alone (soluble or immobilized on agarose beads) or together with soluble GST-PTP
-D2 or with GST (control) was
added to the reaction mixture, and the reaction was allowed to proceed for 2 to 5 min. The reaction was then stopped with 900 µl of 1.0 M
NaOH, and the absorbance of p-nitrophenolate at 450 nm was
determined and compared against a standard curve. For assays performed
with GST-PTP
-D1 precipitated from Cos7 cells, the activity
against PNPP of the precipitated GST-PTP
-D1, alone or when
coprecipitated with HA-PTP
-D2, was analyzed exactly as
described above. Our unpublished work shows no difference in the amount
of substrate (PNPP) metabolized after 5 min in the presence of 100, 200, or 300 mM PNPP, suggesting that the substrate was not limiting in our assays. Moreover, preincubation of the PNPP-containing reaction mixture with the D2 of PTP
or PTP
did not have a significant effect on the amount of PNPP metabolized by PTP
-D1, suggesting that substrate sequestration by the (inactive) D2 domains is not significant.
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RESULTS |
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Identification of PTP
-D2 in two-hybrid screens using
PTP
-D1 as bait.
We have been studying the role of
PTP
in mammalian development. To gain insight into the possible
regulation of this phosphatase, we performed a yeast
two-hybrid screen with the first catalytic domain of rat PTP
(PTP
-D1, fused to the LexA DNA binding domain) as bait (Fig.
1A) to identify interacting, possibly
regulatory, proteins. The bait sequence used (aa 1238 to 1606;
reference 52) also included the wedge region of
PTP
(aa 1285 to 1317) and contained a Cys-to-Ser mutation at the
highly conserved (V/I)HCxAGxxR(T/S)G signature
motif. Our screens of either a rat lung library or a mouse embryonic
(11-day) library resulted in the isolation of several strong positive
clones corresponding to the second catalytic domain of PTP
(Fig.
1B). The clones isolated from the mouse embryonic library were 26 and
48 aa shorter at the N terminus than the rat clone (the shortest clone
was missing most of the highly conserved DYINAS sequence [Fig. 1B and
2]), suggesting that these N-terminal amino acids in PTP
-D2 are not necessary for binding. Comparison of the second catalytic domain of rat PTP
to that of the
previously cloned mouse PTP
(23) reveals 97%
sequence identity at the amino acid level (Fig. 2).
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Binding of PTP
-D2, but not LAR-D2 or PTP
-D2,
to PTP
-D1 in a yeast two-hybrid binding assay.
The second catalytic domains of PTP
(PTP
-D2), LAR
(LAR-D2), and PTP
(PTP
-D2) share a high degree of
sequence similarity (Fig. 2). We therefore wanted to investigate
whether LAR-D2 and PTP
-D2 can also bind to PTP
-D1 by
using yeast two-hybrid binding assays. As shown in Fig.
3A, cotransformation in yeast of
PTP
-D1 (fused to the DNA binding domain) together with
PTP
-D2 (fused to the transactivation domain) caused strong
expression of
-gal, a marker enzyme indicating an interaction
between the two proteins. Only basal levels of
-gal activity were
detected when either clone was transformed alone (Fig. 3A) or when
PTP
-D1 was cotransformed with an unrelated protein (Nedd4; data
not shown). In contrast to the PTP
-D1-PTP
-D2
interaction, cotransformation of PTP
-D1 with LAR-D2 or
with PTP
-D2 resulted in very weak
-gal expression, similar to
that of the negative control (PTP
-D1 alone, LAR-D2 alone, or
PTP
-D2 alone) (Fig. 3A). This lack of interaction was also
apparent upon cotransformation of PTP
-D1 and PTP
-D2
expressed in the reciprocal vectors, i.e., PTP
-D2 fused to the
LexA DNA binding domain and PTP
-D1 fused to the transactivation
domain (Fig. 3A). To ensure that this lack of interaction was not
caused by too low or inappropriate expression of PTP
-D2 relative
to PTP
-D2 in yeast cells, we epitope tagged both D2 domains with a FLAG tag and expressed them in the pACT vector, which leads to higher
levels of protein expression. As can be seen in Fig. 3B, both proteins
were highly expressed in yeast cells, yet only PTP
-D2, and not
PTP
-D2, was able to interact with PTP
-D1. Moreover, unlike the reported homodimerization of the D1 domain of RPTP
(3), there was no detectable
PTP
-D1-PTP
-D1 association when the domain was
expressed on both the DNA binding and the transactivation domains (data
not shown). These results demonstrate that PTP
-D1 preferentially
associates with PTP
-D2 and not with LAR-D2 or with
PTP
-D2, despite close sequence similarity between the second catalytic domains of all three PTPs. They also suggest a D1-D2 heterodimerization rather than a D1-D1 homodimerization type of interaction between these domains.
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-D1-PTP
-D2 binding, we
wanted to test whether the association is reciprocal, i.e., if PTP
-D1 can bind PTP
-D2. We therefore isolated
PTP
-D1 by PCR cloning, generated LexA-PTP
-D1 and
Gal4-PTP
-D2 constructs, and analyzed the binding of these
fusion proteins in yeast two-hybrid binding assays. Our results show no
interaction between these domains, despite the expression of both
proteins in yeast L40 cells (Fig. 3C), suggesting that the two
phosphatases interact in a unidirectional manner, by association of
PTP
-D1 with PTP
-D2, and not vice versa.
Coprecipitation of PTP
-D1 with PTP
-D2 expressed in
mammalian cells.
To test whether the interaction between
PTP
-D1 and PTP
-D2 is not just an anomaly associated with
the yeast two-hybrid binding assay, we transfected the above-described
PTP catalytic domains into mammalian cells and tested their
interactions by coprecipitation. Thus, GST-tagged PTP
-D1 (in
vector pEBG) was cotransfected with HA-tagged PTP
-D2 (in vector
pCMV4) into Cos7 cells. Transfected cells were lysed, and the lysate
was incubated with glutathione agarose beads to precipitate the
GST-tagged (PTP
-D1) proteins. The proteins were then separated
by SDS-PAGE and immunoblotted with anti-HA antibodies to detect the
coprecipitation of HA-tagged PTP
-D2. As shown in Fig.
4A, precipitation of PTP
-D1
resulted in coprecipitation of PTP
-D2 with it, confirming our
above-described yeast two-hybrid binding results. That this
association is not a result of nonspecific binding is evident from the
observation that coexpression of an unrelated protein (HA-tagged
rENaC) together with GST-PTP
-D1 did not lead to
coprecipitation of these proteins (Fig. 4A). The
GST-PTP
-D1
HA-PTP
-D2 interaction was equally strong when PTP
-D1 contained a Cys
Ser mutation in the
signature motif (data not shown). In contrast to the strong
association between GST-PTP
-D1 and HA-PTP
-D2,
cotransfection of GST-PTP
-D1 together with either HA-LAR-D2 or
HA-PTP
-D2 did not yield significant binding between
PTP
-D1 and these D2 domains (Fig. 4A), despite similar
levels of protein expression in all transfected cells (Fig. 4B to
D). Upon greater overexpression of the proteins, weak binding of
PTP
-D1 to PTP
-D2 or LAR-D2 was also observed (data not
shown). Collectively, these results are in agreement with the results
of our yeast two-hybrid binding assays described above, as well as with
those of our preliminary in vitro binding assays (data not shown), and
confirm the selectivity for PTP
-D1-PTP
-D2 interactions.
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Inhibition of PTP
-D1 catalytic activity by
PTP
-D2.
In the LAR family PTPs studied to date (including
LAR and PTP
), the first, but not the second, catalytic domains are
active (11a, 38, 44). To test whether the association
between PTP
-D1 and PTP
-D2 had any effect on the catalytic
activity of PTP
-D1, we initially performed a series of
phosphatase assays using PNPP as a substrate for PTP
-D1
and added PTP
-D2 to the reaction mixture. For these experiments,
PTP
-D1 and PTP
-D2 were expressed in bacteria as GST
fusion proteins. Figure 5A shows that
upon addition of soluble GST-PTP
-D2 to the reaction mixture,
dephosphorylation of PNPP by soluble or immobilized GST-PTP
-D1
(Fig. 5A) was reduced by ~50%, an inhibition not seen with GST alone
(Fig. 5A). PTP
-D2 was equally effective in inhibiting the
catalytic activity of the full intracellular domain (which includes
both D1 and D2) of PTP
(data not shown). Moreover, PTP
-D1
expressed in Cos7 cells and then immunoprecipitated was catalytically
active against PNPP and its activity was inhibited by ~50% in cells
cotransfected with PTP
-D2 (Fig. 5B), suggesting that the
coprecipitated PTP
-D2 was partially blocking PTP
-D1
activity. These results, therefore, demonstrate that the association of
PTP
-D2 with PTP
-D1 leads to partial inhibition of the
catalytic activity of the latter.
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Association between PTP
-D1 and PTP
-D2 requires the
wedge sequence.
A recent report describing the dimerization of the
first catalytic domains (D1) of RPTP
identified the wedge sequence
(a helix-turn-helix motif) located in the N terminus of the domain as
the sequence responsible for binding to the active site of the dimer
partner (3). Because the wedge sequence is also conserved in
LAR family members (3; Fig.
6A), we tested
whether this sequence may be responsible or necessary for the
association of PTP
-D1 with PTP
-D2. We therefore repeated
our yeast two-hybrid binding assays and coprecipitation experiments
with mammalian cells by using a wedge-deleted PTP
-D1
(PTP
D1-W) instead of wild-type PTP
-D1 (Fig. 6B). Our
results show that removal of the wedge sequence from PTP
-D1
drastically reduced binding to PTP
-D2, as determined
by yeast two-hybrid binding assays (Fig. 6C) and by
coprecipitation in mammalian cells (Fig. 6D). This effect was seen
despite similar levels of protein expression of PTP
-D1 and PTP
D1-W in both yeast (Fig. 6C, inset) and mammalian (Fig. 6D, two bottom panels) cells. Thus, the wedge region, previously shown to
mediate RPTP
D1-D1 homodimerization (3), is also involved in the D1-D2 heterodimerization of PTP
and PTP
.
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Weak binding of the D1 domains of other LAR family members to
PTP
-D2.
Our results described above demonstrate an
association between PTP
-D1 and PTP
-D2. To test whether
the D1 domains of PTP
and LAR are also able to bind PTP
-D2,
we expressed either of these domains as a LexA fusion protein and
cotransformed yeast L40 cells with this construct together with
Gal4-PTP
-D2. Our results show that PTP
-D1 was also able
to bind PTP
-D2, but the interaction was weaker (~20%) than
that seen with PTP
-D1; the interaction of LAR-D1 with
PTP
-D2 was even weaker (~7%) (Fig. 7A). Accordingly, cotransfection of
LAR-D1 and PTP
-D2 revealed poor binding between the two
proteins, unlike the strong association observed between PTP
-D1
and PTP
-D2 (Fig. 7B). The interaction between PTP
-D1
and PTP
-D2 could not be assessed in Cos7 cells due to the
instability of the former protein in these cells. As observed
above with PTP
-D1, neither PTP
-D1 nor LAR-D1 was able to
bind the D2 domain of PTP
or LAR, as determined by yeast two-hybrid binding assays or by coprecipitation experiments with mammalian cells
(data not shown). Thus, these result suggest that the first catalytic domain of LAR family members, especially PTP
, are able to
bind the second catalytic domain of PTP
(but not other D2 domains) and that the hierarchy of interactions is
PTP
-D1>>PTP
-D1>LAR-D1.
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DISCUSSION |
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In this report, we demonstrate that the first catalytic domain of
PTP
binds to the second catalytic domain of PTP
, an interaction which requires the presence of the wedge sequence of PTP
and which
leads to partial inhibition of the catalytic activity of PTP
. The
first catalytic domains of other LAR family members can also bind
PTP
-D2, although more weakly, and none of the LAR family D1
domains is able to bind D2 domains other than that of PTP
.
A recent determination of the tertiary structure of RPTP
-D1
revealed that the domain crystallizes as a homodimer. This D1-D1 dimerization is mediated by an ~30-aa helix-turn-helix (wedge) sequence located at the N terminus of RPTP
-D1 which is tucked into
the active site of the opposing partner of the dimer (3). Based on this structure, it was predicted that such dimerization would
inhibit catalytic activity, because the active site is occupied by the
wedge sequence. Our PTP
-D1-PTP
-D2 heterodimerization results provide a variation on this theme, but with a fundamental difference; we believe that the wedge sequence of PTP
-D1 indeed binds to the "pseudoactive" site of PTP
-D2 (i.e., homologous to the active site of D1), which, like other LAR family D2 domains, is
catalytically inactive (11a). The D2 domains of LAR family RPTPs (and other RPTPs) do not possess an N-terminal wedge sequence, and moreover, all of the PTP
-D2 sequences that we isolated in the yeast two-hybrid screens did not contain their N termini. Thus, the
observed inhibition of PTP
-D1 catalytic activity
suggests either the existence of a downstream inhibitory region(s)
in the D2 domain of PTP
which may bind to and inhibits the active
site of PTP
-D1 or that binding of PTP
-D2 to the wedge
sequence of PTP
-D1 somehow distorts the active site of
PTP
-D1 or, alternatively, interferes with substrate
accessibility. Determination of the tertiary structure of D2 domains
alone or in complex with D1 domains, not yet available, should help in
the identification of the exact mode of D1-D2 interactions and
D2-mediated inhibition of D1 catalytic activity. Whatever the
mechanism(s) of binding, the observation of partial inhibition of
the PTP activity of a D1 domain by a D2 domain could have important
biological implications (see below). More importantly, it may
provide an explanation for the long-standing observation that the
D2 domains of many RPTPs are inactive; our work suggests that the
role of these D2 domains is to regulate the activity of the D1 domains.
In LAR family members, this regulation is likely mediated by
intermolecular interactions between these closely related
phosphatases, although we cannot preclude the possibility of a weak
intermolecular association between the two catalytic domains of PTP
.
Based on our lack of PTP
D1-D1 binding, we believe that the observed
D1-D1 homodimerization of RPTP
(3) may represent a
different mode of regulation of that phosphatase; indeed, unlike most
receptor PTPs, both catalytic domains of RPTP
are catalytically
active (47).
An alternative possibility to explain our data, although less likely,
is that the PTP
-D1-PTP
-D2 association somehow induces PTP
D1-D1 dimerization which was undetected by our yeast two-hybrid binding assays.
The second catalytic domains of LAR, PTP
, and PTP
are very
similar in sequence, with only minor substitutions, mostly in nonconserved amino acids (Fig. 2). It is therefore difficult to explain
the vast difference between PTP
-D2 and the D2 domain of PTP
or LAR in the ability to bind to PTP
-D1 (or other LAR family D1
domains), demonstrated here by yeast two-hybrid binding assays and
coprecipitations from mammalian cells. Detailed mutation analysis is
required to sort out the source of this specificity.
The ectodomains of LAR, PTP
, and PTP
are composed of Ig and FNIII
repeats, resembling the cell adhesion molecules N-CAM, fasciclin, and
L1. CAMs such as N-CAM or Ng-CAM have been demonstrated to aggregate
through homophilic interactions (14). Indeed, recent studies
have demonstrated that PTP
, PTPµ, and PTP
, a subfamily of
phosphatases closely resembling LAR, can each aggregate via its
extracellular domain in a homophilic, but not heterophilic, manner
(4-6, 13, 31, 53); such interactions, however, have no
effect on the catalytic activity of these PTPs (5, 31). So
far, homophilic interactions of LAR, PTP
, and PTP
have not been
demonstrated, raising the possibility that the ectodomains of these
PTPs interact either with other extracellular components (e.g.,
extracellular matrix proteins) or, possibly, with each other in a
heterophilic manner. Our results described here demonstrate that these
LAR family PTPs can form heterocomplexes via their intracellular
domains. Moreover, such putative heterodimerization is likely to
inhibit the catalytic activity of at least one of the binding partners.
Although it is not known whether these LAR family members are
coexpressed in the same cells, this is likely, since recent reports
have demonstrated expression of these PTPs in the same types of
neuronal and epithelial tissues or cells. For example, both PTP
and
PTP
have been shown to be expressed in the hippocampus, especially
in the pyramidal cell layer and granular layer of dentate gyrus
(23, 45, 46, 49), and we and others have found that LAR,
PTP
, and PTP
are expressed in fetal alveolar epithelial cells
(11a, 17, 18). In addition, a recent report has demonstrated
colocalization of PTP
and LAR in adhesion plaques of A431 cells
(1).
The physiological substrates for most PTPs, including LAR family
members, are not known. Several proteins that interact with LAR family
members have been described recently, but unlike the PTP
-D2
described here, none seem to affect PTP activity. A coiled-coil phosphoserine called LAR-interacting protein was shown to bind to the
second catalytic domains of LAR, PTP
, and PTP
and appears to
localize LAR to focal adhesions (29, 32). Recently, it was
demonstrated that LAR family members can associate with the
-catenin-cadherin complex and can dephosphorylate
-catenin in vitro (1, 22). The cadherin-
-,
-, or
-catenin
complex is associated with the cytoskeleton and is found in regions of cell-cell contact. The presence of these phosphatases in such regions
suggests that the interactions may regulate tyrosine dephosphorylation of
-catenin, thus affecting the integrity of the cadherin-
-,
-, or
-catenin complex and therefore that of cell adhesion. This
could potentially have major implications for tissue development, particularly for events associated with neurite outgrowth and epithelial differentiation. Several drosophila receptor PTPs, including DLAR, have been shown to be expressed in a subset of the developing axons and pioneer neurons in the central nervous system
(41, 50) and were recently demonstrated to be necessary for
motor axon guidance in the Drosophila embryo (8,
21). This suggests that LAR or its other family members may have
a parallel role in vertebrates as well. Indeed, PTP
, PTP
,
and LAR were previously shown to be strongly expressed during
development in selected regions within the central nervous system and
the peripheral nervous system, as well as in other epithelial and neuroepithelial cells (17, 18, 23, 34, 35, 45, 46, 49), and
a recent gene knockout of LAR has demonstrated a reduction in the size
of cholinergic neurons and defects in hippocampal cholinergic
innervation (51).
The biological meaning of our observed association between LAR family
members, especially between PTP
-D1 and PTP
-D2, and the
resultant inhibition of PTP activity, is not known. It is possible,
however, that such an association keeps one or both binding partners in
an inactive state, perhaps analogous to the intramolecular interactions
recently identified in src family members which keep the kinase domain
inactive (33, 48). We speculate that upon arrival of the
appropriate (high-affinity) tyrosine-phosphorylated substrate, the
D1-D2 intermolecular complex is likely to dissociate, allowing
substrate dephosphorylation (Fig. 8). The
identification of a biological substrate(s) for PTP
, the role that
this PTP and other LAR family members play in neuronal and epithelial
morphogenesis and development, and the possible inhibitory role of
PTP
in these processes, are important questions that now need to be
addressed.
|
| |
ACKNOWLEDGMENTS |
|---|
M.J.W. and C.F. contributed equally to this work.
We thank Barry Goldstein for LAR and LAR-PTP2 (PTP
) cDNA.
This work was supported by a Group Grant on Lung Development from the Medical Research Council (MRC) of Canada, an Operating Grant from the Canadian MRC, the Canadian CF Foundation, and the International Human Frontier Science Program (to D.R.). D.R. was a recipient of a Scholarship from the Canadian MRC. J.B. and M.J.W. are recipients of a Fellowship from the Canadian Lung Association/Canadian MRC.
| |
FOOTNOTES |
|---|
* Corresponding author. Mailing address: The Hospital for Sick Children, 555 University Avenue, Toronto, Ontario, Canada M5G 1X8. Phone: (416) 813-5098. Fax: (416) 813-5771. E-mail: drotin{at}sickkids.on.ca.
| |
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