Dimerization of Receptor Protein-Tyrosine Phosphatase alpha in living cells
© Tertoolen et al; licensee BioMed Central Ltd. 2001
Received: 5 March 2001
Accepted: 1 June 2001
Published: 1 June 2001
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© Tertoolen et al; licensee BioMed Central Ltd. 2001
Received: 5 March 2001
Accepted: 1 June 2001
Published: 1 June 2001
Dimerization is an important regulatory mechanism of single membrane-spanning receptors. For instance, activation of receptor protein-tyrosine kinases (RPTKs) involves dimerization. Structural, functional and biochemical studies suggested that the enzymatic counterparts of RPTKs, the receptor protein-tyrosine phosphatases (RPTPs), are inhibited by dimerization, but whether RPTPs actually dimerize in living cells remained to be determined.
In order to assess RPTP dimerization, we have assayed Fluorescence Resonance Energy Transfer (FRET) between chimeric proteins of cyan- and yellow-emitting derivatives of green fluorescent protein, fused to RPTPα, using three different techniques: dual wavelength excitation, spectral imaging and fluorescence lifetime imaging. All three techniques suggested that FRET occurred between RPTPα -CFP and -YFP fusion proteins, and thus that RPTPα dimerized in living cells. RPTPα dimerization was constitutive, extensive and specific. RPTPα dimerization was consistent with cross-linking experiments, using a non-cell-permeable chemical cross-linker. Using a panel of deletion mutants, we found that the transmembrane domain was required and sufficient for dimerization.
We demonstrate here that RPTPα dimerized constitutively in living cells, which may be mediated by the transmembrane domain, providing strong support for the model that dimerization is involved in regulation of RPTPs.
Protein phosphorylation on tyrosine residues is one of the most important eukaryotic cell signalling mechanisms, and cellular protein phosphotyrosine (pTyr) levels are regulated by the antagonistic activities of the protein-tyrosine kinases (PTKs) and protein-tyrosine phosphatases (PTPs) . Our insight into the function of PTPs - in contrast to the PTKs - is limited. However, in recent years evidence is emerging that PTPs play important and specific roles in biological processes [2,3,4]. Transmembrane PTPs, tentatively called receptor PTPs, RPTPs, are interesting, because their diverse extracellular domains may function as ligand binding domains. Recently, Pleiotrophin was identified as a ligand of RPTPβ/ζ, and binding of Pleiotrophin led to inactivation of RPTPβ/ζ activity, resulting in increased tyrosine phosphorylation of its substrate, β-Catenin . The mechanism underlying ligand-induced modulation of RPTP activity remains to be determined.
Dimerization is a well-established regulatory mechanism for single membrane-spanning receptors [6,7], which may be involved in regulation of RPTPs. The crystal structure of RPTPα-D1, which is highly homologous to other PTPs [8,9,10,11,12,13,14], showed that an amino-terminal helix-turn-helix wedge-like segment interacts with the dyad-related monomer, in such a manner that both catalytic sites in the dimer are occluded . It should be noted that the crystal structures of RPTPμ-D1 and of the complete cytoplasmic domain of LAR do not show dimers like RPTPα [11,12]. Functional experiments with EGFR/CD45, a chimeric protein, consisting of the extracellular domain of the Epidermal Growth Factor Receptor (EGFR) and the intracellular domain of the RPTP, CD45, demonstrated that ligand-induced dimerization leads to functional inactivation of CD45 . Mutation of a single residue in the wedge of CD45 abolishes dimerization-induced functional inactivation . The importance of tight regulation of CD45 was demonstrated by the introduction of a targeted point mutation in the wedge in mice. The CD45 E613R mutation causes lymphoproliferation and autoimmunity in mice, resulting in death . RPTPα dephosphorylates and activates the PTK c-Src [18,19,20,21] via a phosphotyrosine displacement mechanism, involving Src pTyr529 and RPTPα pTyr789 . Recently, we demonstrated that forced dimerization of RPTPα by introduction of a disulfide bridge in the extracellular domain leads to inactivation of RPTPα as assessed by analysis of c-Src Tyr529 phosphorylation and activity. Similar to CD45 , dimerization-induced inactivation of RPTPα was dependent on an intact wedge . Therefore, it is clear that RPTPs can be regulated by dimerization. Experiments with chemical cross-linkers indicated that RPTPα forms dimers . The use of chemical cross-linkers to analyze protein-protein interactions has many disadvantages due to the fact that detection of the interaction is indirect. Here we report the analysis of RPTPα dimerization in living cells using a non-invasive, non-destructive method, Fluorescence Resonance Energy Transfer (FRET).
FRET is a method to assess distances at the molecular level. FRET is a direct radiation-less transfer of energy from the excited donor fluorophore to the acceptor fluorophore, which occurs if the two fluorophores are in very near vicinity of each other, usually less than 60Å [25,26,27]. Recently, FRET has been applied to analyse interactions between proteins, fused to derivatives of Green Fluorescent Protein (GFP) [28,29]. For instance, GFP-FRET has been used to assay changes in cytosolic Ca2+-concentrations using "cameleons" [30,31], to probe interactions between Bcl-2 and Bax in mitochondria , to visualize nuclear interactions of Pit-1 with other transcription factors , to analyze interactions between nuclear import factors , to assess dimerization of G protein coupled receptors , and to follow the interaction between GRB2 and the Epidermal Growth Factor Receptor (EGFR) in vivo .
We have assayed FRET between fusion proteins of RPTPα, fused to CFP and YFP, by dual wavelength excitation, spectral imaging microscopy (SPIM) and fluorescence lifetime imaging microscopy (FLIM), and found that FRET occurred, using all three methods, suggesting that RPTPα dimerized constitutively. The FRET results were consistent with experiments using chemical cross-linkers. Finally, using deletion mutants, we mapped a site of interaction to the transmembrane domain, and demonstrate that the transmembrane domain is sufficient to drive dimerization. Our results provide strong support for the model that RPTPs are regulated by dimerization, and may be useful for analysis of factors that modulate RPTPalpha dimerization.
Routinely, we determined fluorescence intensities in single living cells following excitation at 440 nm and 490 nm, using the FRET filter. Initially, we used RPTPα-516-CFP/-YFP, human EGFR-CFP/-YFP as a negative control, and constitutively dimeric RPTPα-P137C-CFP/-YFP as a positive control (Fig. 2A). The F(440)/F(490) ratio consists of three terms: direct YFP emission, direct CFP emission, and sensitized emission (directly proportional to FRET efficiency (E) and dimerization (α), two terms that cannot be separated from each other (see Materials and Methods). We have calculated the theoretical values of F(440)/F(490) under conditions of no dimerization (E*α = 0), and substantial dimerization (E*α = 0.5), and plotted these values against the ratio of the CFP- and YFP-concentrations R(CFP/YFP) (Fig. 2B). It is clear from Fig. 2B that F(440)/F(490) is substantially higher under conditions of dimerization and FRET (E*α = 0.5) than under conditions of no FRET and/or dimerization (E*α = 0). In addition, it is clear that changes in the CFP:YFP expression ratio affect F(440)/F(490). We have indicated the theoretical F(440)/F(490) levels under conditions of no dimerization and/or FRET with R(CFP/YFP) = 0.5 (F(440)/F(490) = 0.25) and R(CFP/YFP) = 1.5 (F(440)/F(490) = 0.37) (Fig. 2B, bottom and top dashed lines, respectively). We have analysed protein expression of the CFP- and YFP-fusion proteins by immunoblotting and found roughly similar expression levels for all the fusion proteins that we used, which was expected since the only difference between these proteins is a limited number of point mutations (six) that affect the fluorophore. The ratio of concentrations of CFP and YFP was close to 1.0 for all fusion protein pairs, and certainly 0.5<R(CFP/YFP) <1.5. Hence, if the experimental F(440)/F(490) for any CFP/YFP fusion protein pair is 0.25 or less, there is no dimerization and/or FRET. If F(440)/F(490) > 0.37, our simulations (Fig. 2B) indicate that FRET and hence a physical interaction (dimerization) occurs. If 0.25 <F(440)/F(490) < 0.37, one cannot distinguish whether the increased value is caused by a high CFP:YFP expression ratio in the cells or by dimerization-induced CFP to YFP FRET with this method. We postulate that under our experimental conditions with similar amounts of CFP and YFP fusion proteins expressed in the total cell population, an average F(440)/F(490) ratio value >0.37 for a large group of individual cell measurements is the result of FRET and dimerization.
To exclude the possibility that the increased YFP emission was caused by a high YFP to CFP expression ratio, we performed FLIM experiments on the very same cells by employing the other detector port of the SPIM-FLIM imaging microscope. FRET causes a decrease in the donor (i.e. CFP) fluorescence lifetime [25,38]. As shown in Fig. 3B, the CFP-fluorescence lifetimes (τφ) were lower in case of the RPTPα-CFP and RPTPα-YFP coexpressing cells than in control cells, coexpressing RPTPα-CFP and EGFR-YFP. The decrease in the CFP-fluorescence lifetime from 2.6 ns to 2.2 ns clearly demonstrated CFP to YFP FRET due to close proximity of the two fluorophores, which cannot be explained by variation in CFP:YFP expression ratios . The control CFP-lifetime of 2.6 ns was identical to that of cells expressing cytosolic (unfused) CFP which has a τφ lifetime of 2.6 ns at the employed modulation frequency (data not shown, see also ). Because the reduction in CFP lifetime was rather low, we correlated the lifetime data with the SPIM data (see Fig. 3C). A clear negative correlation was observed between the emission ratios and the CFP fluorescence lifetimes (both τφ and τM). These data confirm the notion that the increased YFP fluorescence for the RPTPα-CFP and RPTPα-YFP coexpressing cells (Fig. 3A) indeed reflects sensitized emission resulting from FRET. Furthermore, the experiments in Fig. 3A and 3C also show that the FRET is not due to nonspecific RPTPα interactions as a result of high concentrations in the plasma membrane, since the control with EGFR-YFP did not show sensitized YFP emission. Hence these FRET experiments demonstrate a specific RPTPα-RPTPα interaction (i.e. dimerization) in the plasma membrane of living SK-N-MC neuroepithelioma cells.
Here, we provide evidence that RPTPα dimerized in living cells. We have successfully used FRET to detect RPTPα dimers and dimerization was consistent with cross-linking experiments. Our results provide strong support for the model that dimerization is involved in regulation of RPTPs.
FRET was detected between RPTPα-CFP and RPTPα-YFP using three different techniques: dual wavelength excitation (Fig. 2), SPIM (Fig. 3A) and FLIM (Fig. 3B). The excitation-ratio method for determining FRET suffers from uncertainties when the CFP:YFP expression ratios, R(CFP/YFP), are not exactly known. Simulations show that high excitation ratios can be caused either by FRET or by an unfavorable (high) R(CFP/YFP) (Fig. 2B). Although for the entire cell population R(CFP/YFP) expression ratios were close to 1 (Fig. 2D), one cannot determine the expression ratio independently from FRET at the single cell level. Using the SPIM-method, FRET can be mimicked by unfavorable CFP/YFP expression ratios, but, in contrast to the excitation-ratio method, only when the R(CFP/YFP) is very low. The most reliable (but less sensitive) technique is FLIM. In conclusion, ratiometric analysis of FRET by dual wavelength excitation is a sensitive method for detection of FRET, which is reliable when similar levels of the two fluorophores are expressed.
We also demonstrated that RPTPα dimerized by chemical cross-linking, a technique that is fundamentally different from FRET analysis. BS3-mediated chemical cross- linking is a two-step chemical reaction, leading to a covalent bond, while FRET is only dependent on the distance between the two fluorophores. Dimerization, detected by chemical cross-linking appeared to be much less efficient than with FRET. However, this apparent difference is caused by the relatively low efficiency of the cross-linker, BS3. Only ∼ 10% of RPTPα-P137C was detected in dimeric form following BS3-mediated cross-linking, like wild type RPTPα (data not shown), while ∼ 80% of RPTPα-P137C dimerized according to non-denaturing gels (Fig. 2E). According to the FRET analysis dimerization was extensive, since similar FRET levels were detected in wild type RPTPα as in constitutively dimeric RPTPα-P137C (Fig. 2). Taken together, both FRET and cross-linking experiments indicate that dimerization of RPTPα is extensive. Subtle changes, for instance in the wedge, did not affect dimerization detected by FRET, but did lead to detection of reduced dimerization according to the cross-linking experiments . This discrepancy may be explained by the difference in detection of dimerization. The extracellular domain may dimerize, without an interaction intracellularly. Therefore, extracellular cross-linkers may allow detection of these dimers, while FRET analysis does not, due to the topology of the two fluorophores in the intracellular domains. Importantly, the requirements for detection of dimerization are more stringent for cross-linkers than for FRET, since the linker between the two reactive groups in BS3 is only ∼ 12Å, while FRET allows distances up to ∼ 60Å between the two fluorophores. It is noteworthy that a distance of 60Å or less between the two fluorophores is only achieved when the two proteins are in a protein complex, not when they merely colocalize subcellularly. Taken together, both cross-linking and FRET analysis show that RPTPα dimerizes extensively.
FRET analysis demonstrated that the transmembrane domain was sufficient to drive dimerization of RPTPα which is consistent with cross-linking experiments  (Fig. 4B). Other regions in RPTPα may be involved in dimerization of the full length protein as well. For instance, cross-linking experiments demonstrated that the extracellular domain dimerized by itself . By contrast, the extracellular domain of RPTPα fused to the EGFR transmembrane domain did not show FRET (Fig. 5B), suggesting that this fusion protein did not dimerize. This may be explained by the difference between cross-linking and FRET, as described above. Yet other regions in RPTPα may be involved in dimerization as well. We have evidence that RPTPα-D2 binds to RPTPα-D1 . Similarly, CD45 and RPTPμ may be engaged in intra-or intermolecular interactions [41,42]. In addition, we and others found heterodimerization between RPTP-D1s and RPTP-D2s from different RPTPs, suggesting cross-talk between RPTPs [40,43]. It is noteworthy that the wedge is not required for dimerization, since deletion of the wedge did not abolish dimerization (Fig. 5, data not shown). Nevertheless, the wedge may be involved in stabilization of the dimer, since mutations in the wedge decreased the cross-linking efficiency . Moreover, whereas the EGFR transmembrane domain by itself was not sufficient to mediate dimerization (Fig. 5), introduction of the juxtamembrane domain and D1 of RPTPα (residues 200-516) induced dimerization (data not shown). Taken together, multiple regions in RPTPα contribute to dimerization.
Here, we demonstrate for the first time that RPTPα dimerizes constitutively in living cells. Previously, indirect detection of protein-protein interactions using chemical cross-linkers demonstrated that CD45 homodimerizes . In addition, RPTPα elutes from gel filtration columns as a large protein complex, suggesting dimerization or multimerization . Regulation of dimerization is ill-understood. Like dimerization of RPTKs, dimerization of RPTPs may be regulated by ligand binding. RPTPs have diverse extracellular domains, and several RPTPs bind ligands [46,47,48,49]. Interestingly, GPI-linked Contactin binds laterally to the extracellular domain of RPTPα, i.e. in cis on the same cell . Whether Contactin functions as a ligand, or as a ligand-binding moiety remains to be determined. Pleiotrophin, a ligand of RPTPβ/ζ inactivates RPTPβ/ζ activity . Whether ligand binding affects dimerization of any of these RPTPs remains to be determined. RPTPα dimerization was constitutive and extensive. The transmembrane domain of RPTPα by itself was sufficient to drive dimerization, suggesting that ligands are not required for dimerization. However, ligand binding may modulate the extent of RPTPα dimerization. Analysis of RPTP dimerization using FRET may provide a powerful means to screen for factors that modulate dimerization or monomerization, and FRET may facilitate analysis of the dynamics of RPTP dimerization.
Dimerization of RPTPs may negatively regulate their activity. Ligand-induced dimerization of EGFR/CD45 led to functional inactivation . Mutation of a single residue in the wedge of CD45 abolished ligand-induced inactivation , strongly supporting the model that dimerization leads to wedge-mediated occlusion of the catalytic sites. We have demonstrated that constitutively dimeric RPTPα-P137C was inactive, since it failed to dephosphorylate and activate c-Src in vivo. Mutation of the wedge rendered RPTPα-P137C active, while it did not affect dimerization, demonstrating that inactivation of RPTPs by dimerization was dependent on the wedge . RPTPα-F135C with a disulfide bridge at position 135 dimerized constitutively, like RPTPα-P137C, but was still active, like wild type RPTPα. Apparently, dimerization-induced inactivation requires a specific rotational coupling, i.e. the monomers in the inhibited dimer need to be oriented in a specific geometry with respect to each other . Ligand binding to the extracellular domain may induce rotation of the monomers, relative to each other, thus leading to activation or inactivation of RPTPα, similar to ligand-induced rotation that has been suggested to activate RPTKs [51,52].
Previously, we found that phorbol ester treatment of cells led to activation of RPTPα, and to phosphorylation of RPTPα Ser180 and Ser204 [53,54]. These serine phosphorylation sites are localized close to the wedge, suggesting that phosphorylation of these sites may interfere with interactions of the wedge. Phosphorylation of Ser180 and Ser204 may not affect dimerization per se, but still may lead to opening up of the catalytic site and thus to activation of RPTPα.
Since the crystal structures of RPTPμ-D1 and LAR did not show dimers like RPTPα-D1 [9,11,12], the model that RPTPs are regulated by dimerization has been the subject of debate. The transmembrane domain of RPTPα was sufficient to drive dimerization. It will be interesting to see whether the transmembrane domains of other RPTPs, including RPTPμ and LAR, drive dimerization as well. We propose that dimerization via the transmembrane domain of RPTPα provides a conformational basis for regulation by dimerization. Whether the dimer is inactive depends on the exact topology of the intracellular domain, which may be regulated by phosphorylation, by interaction with other proteins, or by rotational coupling for instance as a result of ligand binding to the extracellular domain. Here we demonstrate that RPTPα dimerized in living cells for the first time, providing strong support for the model that dimerization is involved in regulation of RPTP activity.
SK-N-MC neuroepithelioma cells were grown in Dulbecco's Modified Eagle's Medium, supplemented with 10% Fetal Calf Serum. Expression vectors for chimeric RPTPα fusion proteins were generated by insertion of CFP and YFP  (kind gift of Roger Y. Tsien) on PCR-amplified BglII-fragments into pSG-HA- RPTPα  in which BglII sites had been introduced at the appropriate sites, position 792, 516, 250 or 200 (numbering according to ) by site-directed mutagenesis. In RPTPα-200Δ ED, the extracellular domain was deleted by deletion of an EcoNI-PstI fragment encompassing residues 30-129. RPTPα-EGFR-200 contains the extracellular domain of RPTPα (residues 1 - 142), fused to the transmembrane domain and 34 residues of the cytoplasmic domain of the EGFR (residues 646 - 702), comparable to RPTPα 200-XFP. Expression vectors for EGFR-XFP were generated by insertion of CFP and YFP into the endogenous BglII-site in the human EGFR (at position Ile923). SK-N-MC neuroepithelioma cells were grown on glass coverslips and transiently transfected with equal amounts (5-10 μg total plasmid DNA per 5 cm dish) of CFP- and YFP-fusion constructs by calcium phosphate precipitation exactly as described  and assayed 48 h after transfection.
For the FRET experiments, the cells were incubated at 220C in a HEPES buffered saline buffer (140 mM NaCl; 5 mM KCl; 2 mM CaCl2; 2 mM MgCl2; 10 mM HEPES; 10 mM glucose; 0.1% BSA; pH 7.50). We used a Leitz orthoplan upright microscope (Leitz GMBH, Wetzlar, Germany) and a SPEX Fluorolog fluorimeter (SPEX Industries, NJ) with two excitation monochromators (slit width: 8 nm). Two filter sets (Ploemopak) were used, the "CFP" filter set (filter #1) with an RKP510 dichroic mirror and a 490 nm long-pass emission filter, and the "FRET" filter (filter #2), equipped with a dichroic mirror RKP510 (reflection short-pass filter) and a BP530-560 (band-pass) emission filter (Leitz GMBH, Wetzlar, Germany). The fluorescence intensity was quantified with a Photon Counting Tube (type 9862, EMI Limited, Middlesex, England). The fluorescence intensities (obtained after excitation at 440 nm or 490 nm) were corrected for differences in excitation light intensities, using the reference photomultiplier. Fluorescence intensities were recorded from single living cells and corrected for background, using adjacent non-transfected cells. Routinely, we determined the F(440) and F(490) ratio in 5 or more individual cells that expressed both the CFP- and YFP-fusion proteins, using both filter sets. The ratio of F(440) and F(490) was determined using the FRET filter and consists of three terms: direct excitation of YFP, residual CFP fluorescence, and FRET, as indicated in the following formula:
In this formula, γ is a correction factor consisting of the relative transmission efficiencies of the microscope and the reflection efficiencies of the dichroic mirror at both excitation wavelengths. In our microscope γ was 1.0. The molar extinction coefficients, εCFP(440), εYFP(440), and εYFP(490) are respectively, 32.5, 10.5 and 55.3 x 103 M-1cm-1 . QE(CFP/YFP) is the detection efficiency ratio for CFP to YFP detection, and we estimated the ratio QE(CFP/YFP) to be 0.2, based on the emission spectra of CFP and YFP, and on the filter set used. R(CFP/YFP) is the molar expression ratio in the cells of CFP and YFP-tagged fusion proteins. E is the FRET efficiency in a CFP and YFP labelled receptor dimer, which is dependent on the distance and orientation of the fluorophores. α is the degree of dimerization, ranging from 0 (no dimerization) to 1 (100% dimerization). The F(440)/F(490) ratio of cells expressing YFP fusion proteins only, was 0.16±0.04, consistent with the theoretical value (terms 2 and 3 of the formula are 0). In case there is no dimerization and/or FRET does not occur, term 3 in the formula above is 0 (α and/or E is 0), and F(440)/F(490) is directly proportional to the R(CFP/YFP) (see Fig. 2B). F(440)/F(490) ranges from 0.25 to 0.37 if R(CFP/YFP) ranges from 0.5 to 1.5, and E*α = 0 (no FRET and/or dimerization) (Fig. 2B). If F(440)/F(490)>0.37 and R(CFP/YFP) is close to 1.0, then term 3 in the formula above must be >0, indicating that FRET occurs between CFP and YFP in dimers of RPTPα fusion proteins. Therefore, we set the threshold level of F(440)/F(490) for FRET and dimerization at 0.37.
Both the SPIM and FLIM detector units were built on a Leica DMR upright epifluorescence microscope. Both detector systems are described in detail elsewhere [28,38]. Briefly, FLIM was implemented in the frequency domain using a homodyne detection scheme and a (wide field) RF-modulated image intensifier-CCD-coupled detector unit, and the SPIM detector unit consisted of an f/4 image spectrograph coupled to a back-illuminated chilled CCD camera.
For SPIM, excitation was provided by a 100 W Hg Arc lamp of which the 435 nm line was selected by inserting an Omega (Brattleboro, VT, USA) 435DF10 bandpass filter in the excitation light path. The excitation light was reflected onto the sample by an Omega 430DCLP dichroic mirror. A Leica 63x PL Apo water objective (NA = 1.2) was employed, using no coverslip, and immersing with HEPES buffered saline buffer (see above) at 20°C. Residual excitation light was rejected using a Schott (Mainz, Germany) GG455 longpass filter. In the spectrograph (Chromex 250 is (Chromex Inc., Albuquerque, NM, USA)) a 150 grooves/mm grating was used with a central wavelength of 500 nm. Single cells were positioned by aligning them across the entrance slit of the spectrograph (set at 200 μm width corresponding with a line of 3 μm width in the object plane). Acquisition time was 1-2 s. Regions of the image spectrum corresponding to the plasma membrane of labeled cells were distance averaged (typically 5-10 rows of pixels) and the resulting fluorescence spectra were corrected for background fluorescence and camera bias by background subtraction using an extracellular region just next to the plasma membrane region from the same spectral image.
For FLIM, the cells were excited with the 457 nm argon-ion laser line modulated at 60.116 MHz and the CFP fluorescence was selectively imaged using an Omega 470 DCLP dichroic mirror and an Omega 487RDF42 bandpass emission filter. 20 phase images (1-3 s each) were taken (10 with increasing and 10 with decreasing the phase allowing correction for photobleaching (which was less than 10% in all cases)). Reference phase settings and modulation were calibrated approximately every 30' by measuring a glass microcuvette filled with an erythrosine-B solution in water (single component fluorescence decay with a lifetime of 0.08 ns). The microscope setup, lifetime-image calculation and image processing are described in detail elsewhere .
Following transient transfection the cells were lysed in cell lysis buffer (50 mM HEPES pH 7.4, 150 mM NaCl, 1.5 mM MgCl2, 1 mM EGTA, 1% Triton X-100, 10% glycerol, 10 u/ml aprotinin, 1 μM pMSF, 200 μM sodium orthovanadate), and the amount of protein in the lysates was determined. Subsequently, cell lysates containing equal amounts of protein were loaded on SDS-PAGE gels (7.5 - 12.5%, depending on the size of the construct). The material on the gels was blotted onto PVDF membranes by semi-dry blotting. The blots were blocked with 5% non-fat milk in TBS-T (50 mM Tris pH 8.0, 150 mM NaCl, 0.05% Tween-20) for 1 h, incubated with antibody in milk for 1.5 h, washed extensively and incubated with the appropriate horseradish peroxidase-conjugated secondary antibody for 1 h. Following extensive washing in TBS-T, the blots were developed using enhanced chemiluminescence. We used anti-GFP MAb (Transduction Labs, Lexington, KY), anti-HA-tag MAb (12CA5), and affinity purified anti-RPTPα antibodies, raised against the complete cytoplasmic domain (#5478)  as primary antibodies.
Prior to cross-linking, transiently transfected SK-N-MC neuroepithelioma cells were washed twice with ice cold phosphate-buffered saline (PBS). Subsequently, the cells were incubated with 2 mg/ml bis [sulfosuccinimidyl]suberate, BS3 (Pierce, Rockford, IL) in PBS for 1 h on ice. Following cross-linking, the cells were washed twice with ice cold PBS, and the cross-linker was quenched with 50 mM Tris for 15 min. The cells were lysed and aliquots of the lysates were run on a 5% SDS-PAGE gel, blotted and probed, using anti-HA-tag antibodies as described above.
RPTK, receptor protein-tyrosine kinase
receptor protein-tyrosine phosphatase
fluorescence resonance energy transfer
green fluorescent protein
cyan fluorescent protein
yellow fluorescent protein
epidermal growth factor receptor
spectral imaging microscopy
fluorescence lifetime imaging microscopy.
The authors would like to thank Roger Y. Tsien for the CFP- and YFP-cDNAs. G.J. is supported by a fellowship from the American Cancer Society. T.H. is a Frank and Else Schilling American Cancer Society Research Professor. This work was supported by grants from the National Cancer Institute (T.H.), the Royal Netherlands Academy of Arts and Sciences (T.W.J.G), and the Dutch Cancer Society (J.O. and J.d.H.).
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