Identification of Protein Tyrosine Phosphatases with Specificity for the Ligand-Activated Growth Hormone Receptor
Christian Pasquali,
Marie-Laure Curchod,
Sébastien Wälchli,
Xavier Espanel,
Mireille Guerrier,
Fabrizio Arigoni,
Ger Strous and
Rob Hooft van Huijsduijnen
Serono Pharmaceutical Research Institute (C.P., M.-L.C., S.W., X.E., M.G., F.A., R.H.v.H.), 1228 Plan-les-Ouates, Geneva, Switzerland; and Department of Cell Biology (G.S.), University Medical Center Utrecht, Heidelberglaan 100 AZU-G02.525, 3584 CX Utrecht, The Netherlands
Address all correspondence and requests for reprints to: Rob Hooft van Huijsduijnen, Serono Pharmaceutical Research Institute 14, chemin des Aulx, 1228 Plan-les-Ouates, Geneva, Switzerland. E-mail: rob.hooft{at}serono.com.
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ABSTRACT
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Protein tyrosine phosphatases (PTPs) play key roles in switching off tyrosine phosphorylation cascades, such as initiated by cytokine receptors. We have used substrate-trapping mutants of a large set of PTPs to identify members of the PTP family that have substrate specificity for the phosphorylated human GH receptor (GHR) intracellular domain. Among 31 PTPs tested, T cell (TC)-PTP, PTP-ß, PTP1B, stomach cancer-associated PTP 1 (SAP-1), Pyst-2, Meg-2, and PTP-H1 showed specificity for phosphorylated GHR that had been produced by coexpression with a kinase in bacteria. We then used GH-induced, phosphorylated GH receptor, purified from overexpressing mammalian cells, in a Far Western-based approach to test whether these seven PTPs were also capable of recognizing ligand-induced, physiologically phosphorylated GHR. In this assay, only TC-PTP, PTP1B, PTP-H1, and SAP-1 interacted with the mature form of the phosphorylated GHR. In parallel, we show that these PTPs recognize very different subsets of the seven GHR tyrosines that are potentially phosphorylated. Finally, mRNA tissue distribution of these PTPs by RT-PCR analysis and coexpression of the wild-type PTPs to test their ability to dephosphorylate ligand-activated GHR suggest PTP-H1 and PTP1B as potential candidates involved in GHR signaling.
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INTRODUCTION
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CYTOKINE RECEPTOR SIGNALING typically involves ligand-mediated receptor dimerization and activation, which initiates a tyrosine phosphorylation cascade. Negative feedback to these signaling events is provided through the induction of suppressors of cytokine signaling (1), the action of signal regulatory proteins (2, 3, 4), protein tyrosine phosphatases (PTPs; Ref.5), and receptor endocytosis (6, 7). Among these, PTPs are enzymes and therefore of pharmaceutical interest as drugs targets for receptor agonists (8, 9, 10). The best-known example of a PTP that inhibits receptor activity is PTP1B, which has phosphorylated insulin receptor as a substrate (11), and also negatively regulates leptin receptor signaling (12, 13, 14, 15). Mice that are mutated for PTP1B show increased insulin sensitivity and are obesity resistant (16, 17). Blocking PTP1B gene expression using antisense oligonucleotides restores glucose levels in diabetic mice (18, 19).
GH signals through its receptor (GHR) by recruitment of Jak2 kinase. Activated Jak2 undergoes autotyrosine phosphorylation and phosphorylates GHR, STAT3, -5a and -5b (20, 21, 22). The STATs translocate to the nucleus as transcription factors (see Refs. 23, 24, 25, 26 for reviews). Physiological evidence indicates that GHR signaling is under exquisite control with different responses to continuous or pulsated GH release (27). Different regions of the GHR have been associated with STAT3 and STAT5 activation (20, 22, 28, 29), and various domains in the intracellular GHR have been identified that are responsible for specific transcriptional responses (reviewed in Ref.21). It has been suggested that GH induces phosphorylation of Tyr333 and/or Tyr338 in the rat GHR, which are possible Jak2 substrates (30, 31). However, more recent data point to Tyr487 and/or Tyr595 [human GHR (hGHR) numbering] as major players in signal transduction (32). Mutation of these tyrosines reduces binding of PTP Src-homology 2 domain (SH2)-containing phosphatase (SHP2) to the activated GHR, and results in prolonged Jak2 and signal transducer and activator of transcription (STAT)5b phosphorylation. SHP2 has been found in a complex consisting of GHR, Jak2 and (presumably) signal regulatory proteins-
, but SHP2 functionally enhances, rather than suppresses GHR signaling. Although these data clearly point to SHP2 recruitment to the activated GHR complex through the PTPs SH2 domain, and to the role of GHR tyrosines in modulating the GH response, the role of SHP2 and other PTPs in GHR signaling is far from clear. Also, fine-mapping of tyrosines involved in receptor signaling led to the conclusion that there is considerable redundancy among these tyrosines (21). Thus, Tyr534, Tyr566, and Tyr627 (porcine GHR numbering) have been implicated in Stat5 binding and signaling (22, 29).
The complexities of GHR signaling are further reflected in the discovery of at least two subtypes of Laron disease (dwarfism phenotypically similar to GH-deficiency) whose patients carry normal GH and GHR genes (33).
In GH-deficient children, GH needs to be administered frequently, and responses are erratic. Therefore, discovery of PTPs that dephosphorylate and inhibit GHR signaling may ultimately be of considerable interest for the development of orally available, effective GH agonists.
We have previously employed substrate trapping mutants of PTPs (34) to systematically test PTPs for activated insulin receptor substrate specificity (11). These mutants carry a single D-A mutation in the WPD amino acid sequence N-terminal of the PTP catalytic core HCSAG amino acid motif. Such mutants have unaltered substrate specificity but remain physically associated with their substrate (35, 36). Using a wide panel of mutated PTP catalytic domains in a novel experimental setup we identified PTP-1B among two PTPs that bound to autophosphorylated insulin receptor (34). In the work described here, we have used this same approach to identify PTPs with substrate specificity for the tyrosine phosphorylated GHR. Furthermore we have combined substrate trapping PTP mutants with a Far Western approach (37) to study PTP recognition of GHR stimulated by GH in cells. Of several PTP candidates able to bind the phosphorylated GHR in vitro, we show that in stable GHR-expressing cells, PTP-H1 and PTP1B dephosphorylate the ligand-activated GHR, arguing for their implication in GHR signaling.
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RESULTS
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Screening for Recognition of Phosphorylated GHR by a Panel of PTPs
To identify PTPs that recognize tyrosine phosphorylated GHR, we cloned, expressed and purified the human intracellular GHR domain as a His6-tagged protein from bacteria (see Materials and Methods). To obtain tyrosine-phosphorylated receptor, the same GHR protein was produced in commercially available bacteria that coexpressed tyrosine kinase Elk. The proteins were isolated and tested for purity (Fig. 1A
). The phosphorylated GHR was detected with an antibody against phospho-tyrosine containing protein (Fig. 1B
, left panel). In addition, we have produced a polyclonal antiserum against this GHR intracellular domain, which detects both native and phosphorylated receptor on Western blot (Fig. 1B
, right panel).

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Fig. 1. Expression of Recombinant GH
A, Coomassie gel with the human intracellular GHR domain produced in bacteria without (hGHR-NP, left panel) and with coexpressing Elk-kinase (hGHR-P, right panel). E1, E2, and E3 represent purification steps; M, size markers. B, Western blot showing that purified phosphorylated (P), but not nonphosphorylated (NP), hGHR is detected with an antityrosine-phosphate antibody, whereas both GHR species are detected with an antibody raised against GHR-P.
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The phosphorylated and native forms of the GHR were subsequently assayed for recognition by a panel of substrate-trapping PTPs, using a procedure developed earlier for the insulinR (11). Briefly, GHR and P-GHR were bound to nitrocellulose filters using a 96-well format dot-blot apparatus. After blocking the filter, each pair of wells was incubated with a different PTP trapping mutant, expressed as a glutathione-S-transferase (GST)-fusion protein. The entire filter was washed, and PTP-fusion proteins bound to their substrates were revealed using an anti-GST antibody. We have focused on single-specificity tyrosine PTPs, and excluded dual-specificity PTPs, such as the MAPK phosphatase, because these phosphatases tend to have soluble kinases rather than cytokine receptors as substrate. Among a panel of 31 PTPs, we found that TC-PTP, PTP-ß, PTP1B, SAP1, Pyst-2, megakaryocyte phosphatase (Meg-2), and PTP-H1 reproducibly bind phosphorylated, but not native GHR (Fig. 2A
). As a negative control, GST without a PTP fusion was used for incubation with GHR (GST-control); as a positive control, GST-PTP1B was directly blotted on the filter (C1 and C2). We then tested whether wild-type PTPs were able to dephosphorylate the GHR. The phosphorylated GHR was incubated with different nonmutated PTPs, and after different incubation times, vanadate (a generic, potent PTP inhibitor) was added to stop the reaction. The samples were then spotted on a filter for analysis with an antiphosphotyrosine antibody to examine the phosphorylation state of the GHR (Fig. 2B
). All PTPs whose trapping mutant bound phosphorylated GHR were active in this assay, but the dephosphorylation rates did not correlate perfectly with binding efficiencies. Thus, PTP-ß was a strong binder, but dephosphorylated the receptor slowly. One reason for a discrepancy between the results in Fig. 2
, A and B, could be that multiple tyrosines are phosphorylated in the GHR. Binding of a PTP trapping mutant may depend on only a subset of these, and the nonmutated version may be able to dephosphorylate this subset, but not other phosphotyrosines, resulting in some cases in a persistent signal in Fig. 2B
. Three active PTPs were subsequently compared with a phosphatase that tested negative in Fig. 2A
, namely cdi-1. As shown in Fig. 2C
, cdi-1 was still capable of dephosphorylating the phosphorylated GHR, albeit with clearly less efficacy than TC-PTP.

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Fig. 2. Substrate Trapping Tyrosine Phosphatases Binding to the Phosphorylated GHR
A, Phosphorylated (white circles) and native GHR protein was pairwise bound to a membrane and incubated with trapping phosphatases. PTP nomenclature is as in (5 ). C1/C2, GST-PTP spotted directly on the membrane; GST-control: trapping with GST without a fused PTP catalytic domain. B, Dephosphorylation assay. Phosphorylated GHR was incubated with PTPs as indicated. At various time points, the reaction was stopped and the phosphorylation state of the GHR examined by anti-Tyr-P dot blot analysis. C, Dephosphorylation assay as in B, including negative control Cdi-1. Signals were measured using a densitometer and plotted, expressing signals at time zero as 100%.
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Testing Whether PTPs Have Naturally Tyrosine Phosphorylated GHR as Substrate
The phosphorylated GHR that we have used for Fig. 2
was prepared by coexpression in bacteria with Elk kinase. It is known that under saturating conditions tyrosine kinases phosphorylate tyrosine residues with reduced specificity, as was demonstrated for Fyn kinase, with a phage peptide library as substrate (38). Although we have identified in Fig. 2
a number of PTPs that recognize one or more of the full set of seven phosphotyrosines in the GHR, it is not certain that all of these are associated with normal GHR signaling. To investigate this issue further we have used a Far Western procedure described earlier (37) in which trapping PTPs are tested on GHR substrates purified from mammalian cells. Because GHR is generally only expressed at low levels, we have used a Chinese hamster ovary (CHO) cell line (CHO-ts20) that stably overexpressed the rabbit GHR (39) as a source for physiologically phosphorylated GHR. As shown in Fig. 3
, our antiserum against hGHR cross-reacted efficiently with the recombinant protein from this cell line. The antiserum recognized two GHR species: the GHR precursor (lower band), and the slower migrating glycosylated mature form (Fig. 3A
, lanes 23, arrows), as was described earlier for this cell line (39). Furthermore, treating these cells with GH resulted in tyrosine phosphorylation of the mature form, as detected by immunoprecipitation (IP) of the GHR followed by Western blotting with antiphosphotyrosine (Fig. 3B
, lane 3). No GHR was detectable in nontransfected CHO cells. Treating the CHO-ts20 cells with vanadate resulted in hyperphosphorylation of the GHR (Fig. 3B
, lane 4), attesting to the highly dynamic character of the intracellular phosphorylation-dephosphorylation equilibrium. Interestingly, vanadate treatment also led to a reduction in the amount of both mature and precursor GHR (Fig. 3A
, lane 4). This confirmed the importance of receptor down-regulation in GHR signaling, and suggested that phosphorylation is a major trigger (6, 7, 40, 41, 42). We reproducibly found that the vanadate- induced phosphorylated GHR migrated slower than the GH-induced one (Fig. 3A
, lane 4 vs. lanes 2 and 3). This suggests that vanadate induced phosphorylation on other (probably more) tyrosines than when the GHR was induced by GH. Finally, we showed that the GHR expressed in the CHO-ts20 cell line is functional, as judged by the tyrosine phosphorylation of STAT-3 and -5, but not STAT-1, that was induced by treatment of the cells with GH (Fig. 3C
).

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Fig. 3. GHR Expression, Phosphorylation, and Signaling in Cell Line CHO-ts20
A, CHO-ts20, but not native CHO cells express GHR, as detected by IP and Western blot analysis (W) with anti-GHR antibody. Arrows indicate the two GHR forms (see text). B, Tyrosine phosphorylation of the mature GHR in CHO-ts20 cells after treatment with GH or vanadate (Van), as measured by IP with anti-GHR and Western analysis with anti-Tyr-P. C, Signaling by GHR in CHO-ts20 cells. Cells were treated for the indicated time with GH and extracts analyzed by Western blot for proteins reacting with antibodies specific for phosphorylated or all forms of STAT-1, -3, and -5.
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To evaluate the feasibility of using GH-induced phospho-GHR purified from the CHO-ts20 cells as a substrate in trapping experiments, we tested the sensitivity of our Far Western assay as well as the amount of phospho-GHR that we could generate from this cell line. This is illustrated in Fig. 4A
in a titration experiment with bacterially produced phospho-GHR detected on a Far Western blot with mutated TC-PTP. We found that we could detect down to 12 ng of the receptor. Combined with an estimated copy number of 105 receptors per cell (Strous, G., unpublished) we calculated that this corresponds to the amount of GHR produced by 4 x 107 cells (see Materials and Methods). Figure 4
, BG, shows conventional and Far Western blots with cellular, phosphorylated GHR. As shown in Fig. 4
, BE (central two lanes), we observed efficient binding of PTP-H1, PTP-1B, TC-PTP, and Sap1 to GH-induced GHR, whereas Meg2, Pyst-2, and PTP-ß (not shown) where either not reproducible or negative. Fas-associated phosphatase (Fap)-1 (Fig. 4G
), was used as a negative control. We next controlled for the presence and phosphorylation state of the GHR by Western blotting with anti-GHR (Fig. 4
, BG, left lanes) and antiphosphotyrosine (right lanes), respectively. As expected, only the mature form of the GHR (upper band) reacted with antityrosine phosphate at the same position as detected with the trapping mutant PTPs. Interestingly, GH treatment consistently resulted in a reduction of the amount of mature, but not immature GHR isolated from cells (Fig. 4
, BD and F, left lanes). This is very similar to the effect of vanadate treatment of cells (Fig. 3B
) and may be part of a naturally occurring negative feedback mechanism that follows receptor phosphorylation.

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Fig. 4. Trapping PTPs Binding to GH-Induced Phosphorylated GHR
A, Far Western (FW) titration assay for bacterially produced, phosphorylated hGHR; indicated amounts of GHR were loaded on gel and detected with trapping TC-PTP. BG, FW/Western blots using GH-induced rabbit GHR from overexpressing CHO-ts20 cells. The immunoprecipitated receptor from GH-treated (GH) and untreated (-) cells was blotted and probed with anti-GHR ( -GHR), as FW with the various trapping mutant PTPs, or with an antibody against tyrosine-phosphate ( -P-Y). For the FW lanes, detection was with an anti-GST antibody; the PTPs were expressed as GST-fusion proteins. Numbers refer to protein gel size markers (kilodaltons).
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Recognition by PTPs of Individual GHR Phosphotyrosine Residues
Several studies have examined the requirements of individual, or subsets of GHR tyrosine residues in either GHR phosphorylation or downstream signaling as measured by Jak2 or STAT phosphorylation. However, these studies do not provide direct information on the tyrosines that are actually phosphorylated. Thus, two studies have implicated Tyr333 and Tyr338 in rat GHR phosphorylation (30, 31) but do not formally show that these tyrosines are themselves phosphorylated. By contrast, a more recent study shows that four other tyrosines, namely Tyr487, Tyr534, Tyr566, or Tyr627 (hGHR numbering) are individually sufficient to signal STAT5 phosphorylation (21).
To discover which tyrosines are recognized by GHR-trapping PTPs we used the SPOT technique (43, 44). In this setup, (phospho-)peptides are directly synthesized (anchored) on membranes, followed by incubation with trapping PTPs. Figure 5
shows the result for all seven tyrosines, as made in individual peptides, from the intracellular human GHR domain. All four PTPs that tested positive in Fig. 4
, BE, bound at least one of these seven phosphopeptides; however, binding affinities differed considerably. Thus, Sap1 bound only, but strongly, Tyr487; PTPH1 only bound Tyr534, whereas PTP1B and TC-PTP bound multiple phosphopeptides. Earlier work suggests that Tyr332, Tyr487, Tyr534, Tyr566, and Tyr627 are all phosphorylated after GH stimulation (21). Apart from Tyr627, all of these also appear good PTP substrates in Fig. 5
. Another study has implicated rat GHR Tyr333 and/or Tyr338 in STAT5 binding (30). The equivalent of these in the human GHR is Tyr436 (Tyr333 is not conserved in the hGHR), which is a very poor substrate for all of the PTPs tested.

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Fig. 5. Different PTPs Recognize Different GHR Phospho-Tyrosine Containing Peptides
Fourteen-tetradecamer peptides containing the seven tyrosines of the human GHR cytoplasmic region were directly synthesized on a membrane (see SPOT technique). Corresponding amino acid sequences are on the right side. Z, Phosphotyrosine and numbers indicate the position of the involved tyrosines in the GHR. All fusion proteins were radiolabeled with protein kinase A except GST-PTP-H1 which was revealed by chemiluminescence after anti-GST immunoblotting. The PTP mutants used to probe the membranes are indicated at the bottom.
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Tissue Distribution of PTPs that Bind the GHR
GHR is expressed in liver, kidney, adrenals, heart, muscle, ovary, mammary gland, gastrointestinal tract, and adipose tissue, but not in testis, thymus, and brain (45). Involvement of a PTP in GHR signaling implies that both proteins are expressed in the same tissue. Because expression data were not available for all PTPs we decided to investigate their expression in various tissues using real-time quantitative RT-PCR (46). Expression data for TC-PTP, PTP-ß, Sap-1, PTP-H1, and PTP1B are presented in Fig. 6
. The expression levels were calculated as a ratio to the GAPDH mRNA signals. Although tissue-to-tissue variations were probably well represented for each PTP mRNA, unknown differences in amplification efficiencies make comparisons between PTPs hazardous. Where data can be compared with existing literature, correlation is good. Thus, PTP-ß, like its mouse ortholog vascular endothelial PTP, was predominantly expressed in highly vascularized tissue like lung, placenta and spleen, as shown earlier (47). Sap1 is known to be expressed at very low levels. Also, this PTP has been associated with gastrointestinal cancers (48, 49), and it is therefore unlikely to be involved in GHR signaling. However, these data need to be interpreted with great caution; a low expression level does not necessarily mean that a gene has no function in the tissue or organ concerned. PTP1B expression is quite low in muscle, yet in PTP1B knockout animals many significant phenotypical observations have been made in muscle tissue (16, 17, 50).

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Fig. 6. Tissue Expression of PTPs
A, Real-time RT-PCR procedure was used to quantify PTP mRNA expression. Signals are represented as a percentage of GAPDH.
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Coexpression of PTPs in CHO-ts20 GHR Cells
To directly test if the PTPs we have identified can negatively regulate GHR signaling, we transfected the stably GHR-expressing cells with hemagglutinin (HA)-tagged, wild-type catalytic domains of PTP1B, PTP-H1, TC-PTP, and Fap-1 (negative control) and looked for their ability to dephosphorylate ligand-activated GHR in living cells. We decided not to test PTP-ß and PTP Sap1 in this study because their special distribution pattern (PTP-ß) or low detection level (Sap-1) as measured by quantitative PCR made it unlikely that these PTPs play an important role in GHR signaling. Figure 7A
shows tyrosine-phosphorylated GHR that was immunoprecipitated from GHR-overexpressing cells that had been treated with GH, and transiently transfected with various PTP expression vectors. As expected, for both PTP1B and PTP-H1 the ligand-stimulated GHR signal showed decreased phosphorylation as compared with the control from nontransfected cells. We have controlled for efficient HA-PTP expression by using an antibody against the HA-tag (Fig. 7B
). As expected, the second negative control, Fap-1, did not affect the GHR phosphorylation state. Surprisingly, however, TC-PTP showed no activity toward the GHR in living cells either, in contrast to what was seen in the previous binding study using PTP mutants (Figs. 2
, 4
, and 5
). This inability of TC-PTP to affect GHR phosphorylation in cellular assays was seen reproducibly (data not shown).

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Fig. 7. PTP Overexpression in Cells with Ligand-Stimulated GHR
A, Phosphotyrosine immunoblot analysis on different wild-type transfected PTPs immunoprecipitated from CHO-ts20 GHR expressing cells with GHR antibodies. Negative controls are non transfected (Ctrl) or transfected (Fap-1). Positive signal represents mature form of the GHR on hGHR ligand-stimulated cells (+). Difference in signal intensity is relative to the PTP activity toward the GHR. B, HA-tag expression immunoblot analysis of the different wt-PTPs used in A. C, Transfection of GHR overexpressing cells with trapping mutant PTPs (as in A). D, HA-tag expression immunoblot analysis of the different mutant-PTPs used in C.
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The results in Fig. 7A
do not allow one to conclude that the dephosphorylation of the GHR depended on the catalytic activity of the PTPs. We therefore repeated the experiment using substrate-trapping mutants of PTPH1, TC-PTP, and PTP1B. As shown in Fig. 7
, C and D, these mutants did not significantly modify the phosphorylation state of the receptor. We therefore conclude that the PTP enzymatic activity is required for GHR dephosphorylation.
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DISCUSSION
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GHR tyrosine phosphorylation and dephosphorylation events are both highly dynamic events and paramount in GHR signaling. We have used a funnel-like approach to identify a subset among a large panel of PTPs that potentially has the tyrosine phosphorylated GHR as a substrate and hence may be involved in GHR signaling regulation. Using GHR hyper-phosphorylated by Elk kinase, we have identified TC-PTP, PTP-ß, Pyst-2, SAP1, Meg-2, PTP1B, and PTPH1 as having substrate specificity for this receptor. In addition, we have shown that these same PTPs (or rather their nonmutated counterparts) can dephosphorylate the GHR. Interestingly, many of these PTPs are closely related. TC-PTP and PTP1B share close sequence similarity as well with PTP-H1 and Meg-2; the other subgroup is SAP1 and PTP-ß (51).
Because we suspected that the GHR we had used was hyperphosphorylated at tyrosines that are not physiologically relevant we performed an analogous experiment in a Far Western setup with GH-induced GHR, purified from eukaryotic cells, and probed with the subset of PTPs. We identified PTP-H1, PTP-1B, TC-PTP, and Sap1 as binding also to this form of the receptor. In parallel, we have analyzed mRNA tissue distribution for these PTPs. The distribution patterns of PTP-ß and Sap1 makes it unlikely that these PTPs play an important role in GHR signaling. This, and their nonuniform tissue distribution argue against their involvement in GHR signaling. Taking together, the different results oriented us toward PTP1B, TC-PTP, and PTP-H1 as potential candidates for GHR dephosphorylation. Among these, PTP1B nor TC-PTP knockout mice show a GH-related phenotype (11, 12). Nevertheless, these PTPs may be involved in a modulation of the sensitivity of the GHR toward its ligand. TC-PTP has been showed to be associated with cell cycle progression, and TC-PTP knockout mice die early after birth (52, 53). PTP-H1 has been associated with Jak 2 (54). Because Jak2 is thought to be responsible for GHR- and autophosphorylation in response to GH-induced signaling, this PTP may be involved in GHR dephosphorylation. PTP1B, but not TC-PTP has also been implicated in Jak2 down-regulation (12, 13, 14). Based on these considerations, we decided to test whether the three PTPs were able to dephosphorylate the GHR in the stably transfected CHO-Ts20 cells. Wild-type, HA-tagged PTP expression vectors were generated and transiently transfected into the GHR-overexpressing cell line, treated or not with GH. The phosphorylation state of the GHR was examined by IP and Western blot analysis with anti-P-Tyr (Fig. 7
). PTPH1 and PTP1B were able to dephosphorylate the receptor, whereas to our surprise, TC-PTP, which recognized different GHR phospho-tyrosine-containing peptides (Fig. 5
), did not emerge as positive candidate.
In the final interpretation of our results, one needs to consider that different PTPs may be involved in different aspects of GHR signaling. This is made plausible by the observation that different PTPs recognize different GHR phosphotyrosines, and the many signaling pathways that initiate at the activated GHR. Although we have cast a wide web by initially using Elk-phosphorylated GHR we cannot exclude the possibility that we may have missed PTPs that dock at or near activated GHR through other means. Such a recruitment may render a PTPs catalytic domain substrate specificity less relevant. Our inability to trap SHP2 to the GHR may reflect this. For the two SH2 domain- containing PTPs (SHP1 and SHP2), substrate specificity may reside in their SH2- and other domains involved in protein-to-protein interactions rather than in their catalytic domains. Thus, our approach may have missed relevant PTPs.
Our results and those from others seem to focus on a small, related subset of PTPs that consist of PTP1B, TC-PTP, and PTP-H1, which share substrate selectivity for the GHR, the insulin receptor and their immediate downstream second messengers insulin receptor substrate-1 and Jak-2. Because the GHR is phosphorylated by Jak-2, which is also autophosphorylated on a related target sequence, it is not too surprising that the same PTP-H1 has substrate selectivity for the GHR and Jak-2. Similarly, the insulin receptor and insulin receptor substrate-1 are both phosphorylated by the insulin receptor kinase at similar sites, both of which are good substrates for PTP1B.
We have begun a systematic search for PTPs that potentially have the activated hGHR as substrate. Our confidence in this approach is based on our earlier success in a systematic trapping assay that correctly identified PTP1B as a major PTP for the insulin receptor. This work indicates that substrate specificity is an important aspect of PTP functional selectivity and that this specificity is mediated in large part through the PTPs catalytic domain.
The recent publication of the public (55) and privately owned (56) human genome drafts allows for an updated estimate of the number of functional PTP genes; the total count of bona fide single specificity PTP genes is not higher than 35. Our analysis involved therefore nearly all PTPs in this set. It is important to stress that the PTPs we have identified represent a broadest selection of potential enzymes for the GHR. Whether these PTPs are actually involved requires additional knowledge of the PTPs intracellular colocalization, and their posttranslational control. In this context it is useful to reflect on the very large number of PTPs, besides PTP1B, whose overexpression results in insulinR dephosphorylation (reviewed in Ref.8), illustrating the difficulty of predicting gene function by biochemical means alone. If GHR regulation depends on PTP-H1, or PTP1B alone (i.e. absence of redundancy among these PTPs), then only the use and careful analysis of knockout animals and cell lines, antisense reagents or specific chemical inhibitors will allow one to unambiguously implicate these PTPs in GHR signaling.
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MATERIALS AND METHODS
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Cells and Media
Nontransfected CHO and full-length rabbit GHR overexpressing cells (CHO-ts20) were cultured and seeded on 10-cm Petri dishes. CHO cells were grown at 37 C, whereas CHO-ts20 at 30 C as previously described (39). CHO cells were grown in MEM with Earles salts and L-glutamine [Life Technologies (Gaithersburg, MD) no. 31095-029] supplemented with 10% FBS, penicillin and streptomycin. CHO-ts20 cells were grown in Eagles MEM
with L-glutamine, ribonucleosides and deoxyribonucleosides from Life Technologies no. 22571-020 supplemented with 10% FBS, penicillin, streptomycin, and 0.45 mg/ml of geneticin.
PTP cDNA Cloning
Cloning and expression of PTP catalytic domains and their trapping mutants was described earlier (11). Briefly, the PTPs used for this study contained the catalytic domain of the original PTP, which was fused to GST and mutated to form a trapping mutant after conversion of a D-to-A in the catalytic domain. For SAP-1 TC-PTP, PTP-1B et PTP-ß the pGEX-4TK3 vector corresponds to the pGEX-2TK vector (Amersham Pharmacia Biotech AB, Uppsala, Sweden) with multicloning sites of the pGEX-4T3 vector (Amersham Pharmacia Biotech). Consequently, pGEX-4TK3, as well as pGEX-2TK, encodes a phosphorylation site for protein kinase A after the GST domain.
GHR Production and Tyrosine Phosphorylation
The intracellular domain of the hGHR (SwissProt P10912; http://us.expasy.org/sprot/) was cloned by PCR-amplification from a publicly available EST clone, (IMAGE no 116954; Invitrogen, Groningen, The Netherlands) using the following oligonucleotides: phGHR-1: 5'-atc gcc atg gtg CTG CCC CCA GTT CCA GTT CCA G-3'; and phGHR-2: 5'-tcg act cga gTT AGT GGT GGT GGT GGT GGT GAG GCA TGA TTT TGT TCA GTT G-3'. The primers were designed such that the resulting PCR product was flanked by an NcoI site upstream of the GHR sequence and a XhoI site downstream of it. In addition, primer phGHR-2 introduced an in-frame C-terminal His6 tag to the translated soluble human GHR sequence. The PCR fragment was digested by NcoI and XhoI and cloned into the Escherichia coli expression vector pET24-d (Novagen, Madison, WI). Soluble nonphosphorylated human GHR-his6 was produced from E. coli strain BL21(DE3) (Novagen), whereas phosphorylated hGHR-his6 was produced from a strain expressing the Elk tyrosine kinase (TKB1, Stratagene, La Jolla, CA).
Immunization
A total of 320 µg of highly purified phospho-hGHR was injected on d 1 (in Complete Freunds Adjuvants), 14, 28, and 56 (Incomplete) into two New Zealand rabbits, for a final bleed on d 80 (Eurogentec), according to ISO 9002 (specific pathogen-free conditions) and European guidelines for animal health.
Dot-Blot Trapping
Phosphorylated and native GHR was spotted on nitrocellulose membranes and tested for binding to trapping mutant GST-PTPs and bound PTPs detected with anti-GST as described previously (11). Briefly, phosphorylated and nonphosphorylated hGHR (0.5 µg in 50 µl of PBS) was spotted on nitrocellulose membrane [BA83 0.2 µm (Schleicher & Schuell, Keene, NH) no. 401388] and fitted in a 96-well dot-blot apparatus [Bio-Rad (Hercules, CA) no. 170-6545]. After 1-h incubation, samples were allowed to block for an additional hour in 200 µl washing buffer (PBS, 0.2% Tween 20) plus 5% nonfat milk powder. After this, the GST-PTP protein was incubated at 1 ng/µl (100 µl) for 1 h at 4 C, followed by washing of the wells. The membrane was taken out of the dot-blot apparatus, washed, then blocked in 10 ml washing buffer plus milk. The membrane was washed again and incubated with anti-GST [mouse monoclonal, 1:1500 (Santa Cruz Biotechnology, Inc., Santa Cruz, CA) B-14 sc-138] for 1 h at room temperature followed by washing and incubation with goat antimouse Ig-horseradish peroxidase (HRP) [1:1500, Dako (Carpinteria, CA) P0447]. Detection was by chemiluminescence (ECL kit, Amersham Pharmacia Biotech, RPN 2106).
Dephosphorylation Assay of hGHR
Essentially as described in (11). Briefly, GST-PTP (wt) proteins were incubated at 0.5 ng in 10 µl with 10 ng phosphorylated hGHR in 40 mM Tris (pH 7), 50 mM NaCl, 1 mM EDTA. The reaction was stopped with 0.4 mM (final) pervanadate after 0.5, 2, 15, or 30 min. The stop solution contained 0.1 M Na3VO4 (Sigma S-6508) and 0.3 M H2O2. The samples were spotted on membrane (BA83 0.2 µm; Schleicher & Schuell no. 401388) and the blot was blocked in 10 ml washing buffer (PBS, 0.2% Tween 20) plus 5% nonfat milk powder. The blot was washed and incubated 1 h at room temperature with antiphosphotyrosine antibody [1:1500, clone 4G10 mouse monoclonal IgG2b
(Upstate Biotechnology, Lake Placid, NY) no. 05-321]. The blot was washed again and incubated 1 h with goat anti-mouse antiserum coupled to HRP (Dako no. P0447) and visualized by ECL as described above.
SDS-PAGE and Western Blot Detection
Gels (7% Tris-acetate; Novex, Frankfurt, Germany) were run according to the manufacturers instructions and protein were transferred onto Hyperbond C extra, nitrocellulose (no. RPN303E, Amersham Life Science, Arlington Heights, IL) membrane using a semi-dry (homemade) transfer apparatus for 3 h at 4 C until 25 V was reached. Protein was stained with Ponceau-S to confirm equal loading of samples and blocking was done in 0.1% PBST washing buffer (0.1% Tween 20 in 1x PBS) plus 5% milk for 30 min. Incubation with mouse antityrosine phosphate antibody was as described above in blocking buffer for 1 h at room temperature but at a concentration of 1:3000. Membranes were rinsed twice for 1 min followed by three 10-min washes. Secondary antibody was goat antimouse coupled to HRP (1:1500, Dako no. P0447) for 1 h at RT. Membranes were rinsed again twice for 1 min and washed three times in washing buffer. Immunoreactive bands were revealed by ECL as mentioned above. For rabbit GHR antisera antibody, dilution was 1:2000 in blocking solution and secondary antibody was goat antirabbit coupled to HRP (1:2000, Bio-Rad no. 1706515). Anti-Stat and phospho-Stat 1 (Tyr701), 3 (Tyr705), and 5 (Tyr694) rabbit antibodies (1:1000), were from Cell Signaling Technology (Beverly, MA)/Biolabs except rabbit anti-Stat5a, which was from Santa Cruz Biotechnology, Inc. Molecular weights were Mark 12 from Invitrogen.
Immunoprecipitation
CHO-ts-20 cells were seeded at 2 x 106 cells per 10 cm plate and allowed to grow 20 h up to 6070% confluency. Cells were then incubated overnight with 10 mM sodium butyrate followed by 4 h of starvation in serum-free medium supplemented with 10 mM sodium butyrate. Cells were then either left untreated, or induced with 1 mM Pervanadate (200 µl of a mix of 100 µl of 0.1 M Na3VO4 and 100 µl of 0.3 M H2O2) for 30 min or induced with hGH, 8 nM for 15 min (100 µl of 0.8 µM Serostim from Serono, Switzerland) diluted 226 times in distilled water). After medium removal, cells were scraped and lysed in 500 µl RIPA buffer [20 mM Tris (pH 7.5), 150 mM NaCl, 0.1% sodium dodecyl sulfate, 0.5% deoxycholate, 1% Triton X-100, 2 mM EDTA] containing protease and phosphatase inhibitors [1 mM Na3VO4, 30 nM okadaic acid, 25 nM microcystin, 10 mM NaF, 40 mM ß-Glycero-PO4, 1 mM Na-Pyro-PO4, 10 µg/ml aprotinin, 10 µg/ml leupeptin, 1 mM Pefabloc SC (Roche, Basel, Switzerland)], homogenized, vortexed and incubated for 20 min at 4 C under rotation. Lysates were centrifuged for 5 min at 14,000 x rpm and supernatants precleared against 40 µl of washed protein G Sepharose packed beads (PGS 4 Fast Flow, Amersham Pharmacia Biotech) for 30 min under rotation. For IP, 400 µl of the precleared lysate was mixed together with GHR antibody coupled to beads (20 µl of packed beads precoupled for 2 h with 10 µl rabbit antiserum and 500 µl of 0.5x RIPA buffer) and incubated at 4 C for 2 h under rotation. Total lysate aliquots were kept separate for protein measurement [Coomassie Plus Protein Assay Reagent from Pierce (Rockford, IL)] and Western blot control analysis. Immunoprecipitated samples were then washed twice with 500 µl of 0.5x washing buffer (RIPA buffer and PBS plus all inhibitors at a ratio of 1:1 vol:vol) followed by one additional wash with RIPA buffer diluted ten times in distilled water. For normal IP Western blot analysis, samples were stopped by addition of 20 µl of Laemmli sample buffer twice and heated, whereas for Far Western immunoblotting analysis, IPs were stopped by addition of 10 µl of Laemmli sample buffer five times, heated 5 min as above, centrifuged and the eluate reused to elute a second parallel IP to increase the number of immunoprecipitated GHR. Twenty microliters of IP eluate were then subjected to SDS-PAGE analysis and first analyzed by Western blot for sample quality control (normal IP) and then for Far Western immunoblotting (coupled Ips).
For GHR IP performed on cells transfected with different PTPs (Fig. 7
), experiments were performed as above except that two plates were used for the assay. Subsequent phosphotyrosine immunoblot analysis were performed using 4G10 (1:1500) mouse monoclonal IgG2b
from Upstate Biotechnology.
Coexpression of PTPs in CHO-ts20 GHR- Expressing Cells
Ten-centimeter plates were transfected with three microgram of DNA (pHM6 vector, Roche) of wild-type or trapping mutant, HA-tagged PTP1B, PTP-H1 or TC-PTP according to FuGENE manufacturers instructions. Controls were nontransfected cells and wt-HA tagged PTP-Fap-1-transfected cells (a PTP that tested negative in the in vitro binding assay). For ligand stimulation and IP, cells were treated as described above except that cell growth was for 30 h instead of 20 h.
Far Western Immunoblotting
Substrate recognition by trapping mutants on cell material after protein transfer was described elsewhere (37) except that washing buffer contained 0.1% Tween 20 and 5% milk blocking. Briefly, membranes were blocked for 1 h in PBST (PBS plus 0.1% Tween 20) containing 5% nonfat milk powder. Incubation overlay with GST-PTP mutant fusion proteins (2 ng/µl) onto the membrane was performed in 3 ml blocking solution (washing buffer, PBST 0.1% Tween 20 plus 5% milk), overnight at 4 C on a balance shaker. The membranes were washed extensively three times for 1 min followed by six washes of 10 min in PBST. Subsequently, membranes were incubated with mouse monoclonal anti-GST antibodies (1:1, 500) in blocking buffer for 3 h at 37 C. After several washes, the blot was finally incubated with polyclonal goat antimouse-coupled HRP (1:1500) in blocking buffer for 60 min at 37 C. Immunodetection was by chemiluminescence as described above. When the membranes were reprobed for GHR and antiphosphotyrosine antibodies, a stripping step was performed [100 mM Tris (pH 6.8), 2% sodium dodecyl sulfate, 100 mM ß-mercaptoethanol] for 30 min at 50 C followed by three washes of 10 min.
SPOT Synthesis and Probing Assay
Peptides were manually synthesized on derivatized cellulose membrane provided by Sigma-Genosys, which also provided the 20 Fmoc-amino acids active esters. Fmoc-phosphotyrosines (Novabiochem no. 0412-1156) were incorporated in presence of the coupling reagent, N,N'-disopropylcarbodiimide (Sigma; Espanel et al., Ref.43). Membranes were blocked 2 h and probed in Western wash buffer [10 mM Tris (pH 7.4), 0.1% Triton X-100, and 150 mM NaCl] containing SPOT blocking buffer (Sigma-Genosys) at 4 C. After 2 h, membranes were washed several times in Western wash buffer and analyzed by autoradiography. For GST-PTP-H1 we experienced radiolabeling problems; therefore, the binding was revealed by enhanced chemiluminescence (ECL; Amersham Pharmacia Biotech) after using anti-GST as first antibody (B-14; 1:2000; Santa Cruz Biotechnology, Inc.), and antimouse coupled to the HRP as secondary antibody (1:2000; Dako).
Real-Time PCR by RT-PCR
PCR primers for human Sap1, TC-PTP, PTP1B, PTP-H1, and glyceraldehyde-3-phosphate dehydrogenase (GAPDH) (housekeeping control) were designed using the Primer Express software from Perkin-Elmer Biosystems based on the published sequences: Sap1, reverse GCG AGT CCA GAG GCC AGT AA; forward CAT GCT GAC CAA CTG CAT GG; TC-PTP, reverse GCC CAA TGC CTG CAC TAC A; forward AGA ATC TGG CTC CTT GAA CCC T; PTP1B, reverse ATG ATG AAT TTG GCA CCT TCG; forward TGA TCC AGA CAG CCG ACC A; PTP-H1, reverse TGG CTT GAT GTC TGC ACC AT forward CCC ACT GGA TAT TGT CCG AAA; GAPDH, reverse GAT GGG ATT TCC ATT GAT GAC A; forward CCA CCC ATG GCA AAT TCC; intron-GAPDH, reverse CCT AGT CCC AGG GCT TTG ATT; forward CTG TGC TCC CAC TCC TGA TTT C. The specificity and the optimal primer concentration were tested on diluted series of plasmids with cDNA inserts. Potential genomic DNA (of cDNA) contamination was excluded by performing PCRs with specific intron-GAPDH primers. The absence of nonspecific amplification was controlled by analyzing the PCR products on 3.5% agarose gels. SYBR green real-time PCRs contained 25 µl SYBR Green PCR master mix (Perkin-Elmer Life and Analytical Sciences, Boston, MA) with 0.5 U AmpErase uracil N-glycosylase and 20 µl of primers (300 nM). Template was 5 µl of reverse transcription products; 0.5 ng total RNA (CLONTECH, Palo Alto, CA) or polyA+ for ovary, using Perkin-Elmer Multiscribe enzyme). PCR was performed at 50 C for 2 min (AmpErase uracil N-glycosylase contaminant DNA digestion; Ref.57), 95 C for 10 min (for AmpliTaq Gold activation) and then run for 40 cycles at 95 C for 15 sec, 60 C for 1 min on the ABI PRISM 7700 Detection System (TaqMan). The reverse-transcribed cDNA samples were thus amplified and their cycle threshold values were determined. All cycle threshold values were normalized to the housekeeping gene GAPDH. A single specific DNA band for human Sap1, TC-PTP, PTP1B, PTP-H1, and GAPDH was observed using gel electrophoresis analysis.
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ACKNOWLEDGMENTS
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We thank Dr. C. Rommel for helpful discussions and Dr. S. Arkinstall for support.
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FOOTNOTES
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Present address for S.W.: Institute for Cancer Research, The Norwegian Radium Hospital, Montebello, 0310 Oslo, Norway.
Present address for X.E.: Sanofi-Synthélabo, Labège Innopole voie 1, BP137, 31676 Labège, France.
Present address for F.A.: Nestlé Research Center P. O. Box 44, 1000 Lausanne 26, Switzerland.
Abbreviations: CHO, Chinese hamster ovarian (cells); Fap, Fas-associated phosphatase; GAPDH, glyceraldehyde-3-phosphate dehydrogenase; GHR, GH receptor; GST, glutathione-S-transferase; HA, hemagglutinin; hGHR, human GHR; HRP, horseradish peroxidase; IP, immunoprecipitation; Jak, Janus kinase; Meg, megakaryocyte phosphatase; PTP, protein tyrosine phosphatase; SH2, Src-homology 2 domain; SHP2, SH2-containing phosphatase; STAT, signal transducer and activator of transcription.
Received for publication January 10, 2003.
Accepted for publication July 28, 2003.
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