Characterization and Binding Specificity of the Monomeric STAT3-SH2 Domain*

Serge HaanDagger , Ulrike HemmannDagger , Ulrich HassiepenDagger , Fred SchaperDagger , Jens Schneider-Mergener§, Axel WollmerDagger , Peter C. HeinrichDagger , and Joachim GrötzingerDagger

From the Dagger  Institut für Biochemie, Rheinisch-Westfälische Technische Hochschule Aachen, Pauwelsstrasse 30, D-52074 Aachen, Germany and the § Institut für Medizische Immunologie, Universitätsklinikum Charité, Humboldt-Universität zu Berlin, D-10098 Berlin, Germany

    ABSTRACT
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Abstract
Introduction
Procedures
Results
Discussion
References

Signal transducers and activators of transcription (STATs) are important mediators of cytokine signal transduction. STAT factors are recruited to phosphotyrosine-containing motifs of activated receptor chains via their SH2 domains. The subsequent tyrosine phosphorylation of the STATs leads to their dissociation from the receptor, dimerization, and translocation to the nucleus. Here we describe the expression, purification, and refolding of the STAT3-SH2 domain. Proper folding of the isolated protein was proven by circular dichroism and fluorescence spectroscopy. The STAT3-SH2 domain undergoes a conformational change upon dimerization. Using an enzyme-linked immunosorbent assay we demonstrate that the monomeric domain binds to specific phosphotyrosine peptides. The specificity of binding to phosphotyrosine peptides was assayed with the tyrosine motif encompassing Tyr705 of STAT3 and with all tyrosine motifs present in the cytoplasmic tail of the signal transducer gp130.

    INTRODUCTION
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Abstract
Introduction
Procedures
Results
Discussion
References

Src homology 2 (SH2)1 domains are highly conserved regions common to a series of cytoplasmic signaling proteins such as the Src family of tyrosine kinases, phospholipase C-gamma , the p85 subunit of phosphatidylinositol 3-kinase, and the STAT family of transcription factors (1, 2). These noncatalytic domains target the proteins to specific phosphotyrosyl peptide sequences within their binding partners, thereby regulating a wide range of intracellular signaling events. The specificity of this interaction is determined by the amino acid sequence surrounding the phosphotyrosine on the one hand and by the SH2 domain on the other (3).

Activation of transcription factors of the STAT family has been shown to require the transient association of the STATs with cytokine receptors (4, 5). STAT factors interact through their SH2 domains with specific phosphotyrosine motifs within the cytoplasmic parts of the activated receptors. In the case of the activation of STAT1 and STAT3 by interleukin-6, four such tyrosine motifs within the interleukin-6 signal-transducing receptor subunit gp130 have been identified (6, 7). Two of these motifs (Y767RHQ and Y814FKQ) give rise to specific STAT3 activation, whereas two others (Y905LPQ and Y915MPQ) are able to recruit both STAT1 and STAT3 (6). Subsequent to receptor binding, the STAT factors are phosphorylated on a single tyrosine residue by receptor-associated tyrosine kinases of the Janus kinase family (8-10). This activation of the STAT factors leads to homo- or heterodimerization and translocation to the nucleus, where they bind to enhancers of interleukin-6-inducible genes resulting in the activation of transcription of, e.g. acute phase protein genes (11-13). The dimerization of STAT factors has also been shown to be mediated by the SH2 domains (9). This has been confirmed recently by x-ray structures of the STAT1 and STAT3 dimers bound to DNA (14, 15). In this complex the two SH2 domains form a tunnel that is passed by the two phosphotyrosine-containing tail segments.

Previous experiments have shown that the SH2 domain is also the sole determinant of specific STAT factor activation via gp130 and the interferon-gamma receptor (16, 17). The mechanism for the binding of STAT monomers to the phosphotyrosine-containing recruitment sites of the cytoplasmic region of signal-transducing receptor subunits still needs to be elucidated.

Here we describe the expression, refolding, and structural characterization of the STAT3-SH2 domain as well as its specific binding to phosphotyrosine peptides. Furthermore, we demonstrate that this interaction requires a monomeric domain.

    EXPERIMENTAL PROCEDURES
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Abstract
Introduction
Procedures
Results
Discussion
References

Peptide Synthesis-- Biotinylated peptides were synthesized as described earlier (18).

Plasmid Construction-- Constructions were carried out using standard procedures (19). For construction of pB-STAT3-BS, the cDNA encoding murine STAT3 (kindly supplied by J. Darnell, Jr., Rockefeller University, New York) was provided with BglII and SalI restriction sites at the 5'- and 3'-ends, respectively, and cloned into a pBluescript vector (Stratagene, Heidelberg, Germany). The sequence encoding the STAT3-SH2 domain (amino acid residues 582-702) was amplified by polymerase chain reaction, and BamHI and AvrII restriction sites were introduced by the 5'- and 3'-primers, respectively. The BamHI/AvrII DNA fragment was ligated with a modified pRSet5c vector carrying an adaptor consisting of an amino-terminal MRGS(H)6-tag and a BamHI and an AvrII restriction site. The resulting vector pRSetS3SH2 coding for the amino-terminally His-tagged STAT3-SH2 domain was verified by DNA sequence analysis.

Expression, Purification, and Refolding of the Recombinant MRGS(H)6-tagged STAT3-SH2 Domain-- Escherichia colistrain BL21(DE3)pLysS transformed with the pRSetS3SH2 plasmid was grown at 37 °C in LB medium containing chloramphenicol (50 µg/ml) and ampicillin (100 µg/ml) to an A595 of 0.6-0.7. Cells were induced with 0.4 mM isopropyl-beta -D-thiogalactopyranoside for 3 h at 37 °C and subsequently harvested by centifugation. The bacterial pellet was resuspended in lysis buffer (26 mM Tris-HCl, pH 7.5, 10 mM EDTA, 1 mM phenylmethylsulfonyl fluoride, 1 mM dithiothreitol), cells were lysed by repeated freezing and thawing, and the inclusion bodies were harvested by centrifugation and purified by five cycles of sonication (constant pulse for 2 min at 0 °C), each followed by centrifugation at 3,700 rpm for 30 min. The inclusion bodies were solubilized in buffer S (50 mM Tris-HCl, pH 8.0, 6 M GdnHCl, 1 mM EDTA, 100 mM dithiothreitol). After incubation at 42 °C for 30 min, insoluble particles were removed by filtration through a 0.45-µm sterile filter, and the buffer pH was adjusted to 2. The denatured STAT3-SH2 domain was purified by reverse phase HPLC using a Polygosil 60-5 C18 column (CS-Chromatographie Service GmbH, Langerwehe, Germany) equilibrated in buffer A (0.1% trifluoroacetic acid). Elution of the STAT3-SH2 domain occurred at 50.3% buffer B (80% acetonitrile, 0.1% trifluoroacetic acid). The purified protein was isolated by lyophilization and solubilized in buffer S. Refolding of the purified SH2 domain was achieved by dialysis at 4 °C against different buffers (buffer C: 20 mM Na2HPO4/KH2PO4, pH 7.5, 150 mM NaCl, 0.5 mM EDTA, 1 mM dithiothreitol; buffer D: 20 mM Na2HPO4/KH2PO4, pH 5.5, 150 mM NaCl, 0.5 mM EDTA; buffer E: 20 mM CH3COONa/CH3COOH, pH 4.5, 150 mM NaCl, 0.5 mM EDTA). For purity determination, the proteins were resolved on a 15% polyacrylamide gel by SDS-PAGE and visualized using SyproTM Orange protein stain reagent (Molecular Probes, Eugene, OR). The quantitative analysis was performed on a Storm 840 fluorescence scanner (Molecular Dynamics, Krefeld, Germany) using Image QuantTM software.

Immunoblot Analysis-- The harvested cell fragments containing the inclusion bodies were resolved by SDS-PAGE and transferred to an Immobilon polyvinylidene difluoride membrane (Millipore, Eschborn, Germany) using a semidry electroblotting apparatus. STAT3-SH2 detection was performed using a monoclonal MRGSHis antibody (QIAGEN, Hilden, Germany). A polyclonal goat anti-mouse horseradish peroxidase-conjugated secondary antibody (DAKO, Hamburg, Germany) was used to visualize the immunoreactive bands by Western blot techniques.

Peptide Binding Assay-- Peptide binding of the isolated pure STAT3-SH2 domain was performed by the means of an ELISA. A 96-well ELISA Maxisorb plate (NUNC, Roskilde, Denmark) was coated with 2.5 µg/ml streptavidin (100 µl/well; 16 h at room temperature). Unoccupied binding sites were blocked with 2% bovine serum albumin in phosphate-buffered saline (10 mM Na2HPO4/KH2PO4, pH 7.4, 200 mM NaCl, 2.5 mM KCl) (200 µl/well; 2 h at room temperature). After four washings with phosphate-buffered saline and 0.02%Tween (200 µl/well), the biotinylated peptides were immobilized by incubating the streptavidin surface with 100 µl/well of a 500 ng/ml peptide solution in phosphate-buffered saline for 1 h at room temperature. The wells were washed four times with 200 µl/well phosphate-buffered saline and 0.02% Tween, and the surface was equilibrated with buffer C or D. STAT3-SH2 solutions were incubated for 1 h at room temperature, and unbound protein was removed by washing four times with the appropriate phosphate buffer. The unoccupied phosphotyrosine residues were detected by incubation with PY20 phosphotyrosine antibody (Transduction Laboratories, Lexington, KY; 1:2,000, 100 µl/well) for 45 min at room temperature. Bound PY20 was visualized using a polyclonal goat anti-mouse horseradish peroxidase-conjugated antibody (DAKO; 1:2,000, 100 µl/well, 45 min at room temperature). Staining reagent was 0.1 mg/ml 3,3',5,5'-tetramethylbenzidine in 0.1 M acetate buffer, pH 5.5, containing 0.003 vol % H2O2. The reaction was stopped with 2 M sulfuric acid. Inhibition of the PY20/phosphopeptide interaction by STAT3-SH2 was determined by calculating the decrease in absorbance with increasing amounts of STAT3-SH2 relative to a SH2-free sample.

Size Exclusion Chromatography-- Size exclusion chromatography was performed on a Bio-SilectTM SEC 125-5 column (Bio-Rad). The column was equilibrated with the refolding buffer C (pH 7.5) or D (pH 5.5), respectively, loaded with 500 µl of STAT3-SH2 (70-80 µg/ml), and run at a constant flow rate of 0.7 ml/min. The collected 0.7-ml fractions were resolved on a 15% polyacrylamide gel by SDS-PAGE and visualized by silver staining.

Circular Dichroism Spectroscopy-- CD measurements were carried out on an AVIV (Lakewood, NJ) 62DS CD spectrometer, equipped with a temperature control unit, and a Jasco J-600 spectropolarimeter, both calibrated with a 0.1% aqueous solution of D-10-camphorsulfonic acid according to Chen and Yang (20). The spectral band width was 1.5 nm. The time constant ranged between 1 and 4 s and the cell path length between 0.1 and 10 mm.

Fluorescence Spectroscopy-- Steady-state fluorescence spectra were recorded on a Spex Fluorolog 211 photon-counting spectrofluorometer (Spex Industries, NY) with a band width of 2.7 nm (excitation monochromator) and 2.2 nm (emission monochromator). The excitation wavelength was 295 nm. The spectra are corrected for changes in lamp intensity and for spectral sensitivity of the emission-monochromator/photomultiplier system. All fluorescence measurements were carried out at 20 °C.

Fluorescence lifetimes and anisotropy decay were measured in the single photon-counting mode with an Edinburgh Instruments Ltd. (U. K.) spectrometer, model 199. The full width at half maximum of the lamp pulse from the hydrogen flashlamp was 1.4 ns. The excitation wavelength was 295 nm and the band width 8 nm. The emitted light was passed through a combination of a UV-transmitting black glass and a cutoff glass filter to create a band pass (WG320, DUG11, Schott, Mainz, Germany). At least 80,000 counts were accumulated in the peak channel of the total fluorescence intensity, I(t). The lamp pulse was recorded with a suspension of Ludox (NEN Life Science Products) at 345 nm. Data handling and the iterative nonlinear least squares fit of the decays were accomplished with a program supplied by Edinburgh Instruments Ltd. Intensity decays (I(t)) were fit to the multiexponential model using I(t) = Sigma  biexp(-t/tau i), where bi values are the amplitudes associated with the decay time tau i. The fractional intensity Bj = bjtau j/Sigma bitau i permits calculation of the mean lifetime <tau > = Sigma  Bitau i.

Fluorescence anisotropy decays were analyzed by an exponential fit.
<IT>r</IT>(<IT>t</IT>)<UP>=</UP><IT>r</IT><UP>exp</UP>(<UP>−</UP><IT>t</IT><UP>/&phgr;</UP>)<UP>+</UP><IT>r</IT><SUB><UP>∞</UP></SUB>
where r0 = r + rinfinity .

The parameters of r(t) are as follows. r is the anisotropy; phi , the rotational correlation time; r0 and rinfinity are the limiting anisotropies, r(t)(t right-arrow 0) = r0 and r(t)(t right-arrow infinity ) = rinfinity . The quality of the fits was gathered from plots of weighted residuals and from the statistical parameter chi 2 (21).

Protein Concentrations-- Protein concentrations were calculated from absorption spectra in the range of 240-320 nm using the method of Waxman et al. (22).

    RESULTS
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Abstract
Introduction
Procedures
Results
Discussion
References

Expression and Purification of Recombinant STAT3-SH2-- To obtain sufficient amounts of protein, the amino-terminally His-tagged STAT3-SH2 domain was expressed in E. coli. The recombinant protein was found entirely in inclusion bodies (Fig. 1A). Repeated sonication and centrifugation yielded inclusion bodies containing about 90% STAT3-SH2 protein. 1 liter of medium contained 40-50 mg of inclusion body proteins. After solubilization of the inclusion bodies in GdnHCl the proteins were separated on a reverse phase HPLC column, and STAT3-SH2-containing peak fractions were lyophilized. The STAT3-SH2 protein proved to be at least 99% pure (Fig. 1B). This procedure yielded 10-15 mg of pure STAT3-SH2/liter of culture.


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Fig. 1.   Expression and purification of STAT3-SH2. A, SDS-PAGE analysis of bacterially expressed STAT3-SH2. Protein bands are visualized by Coomassie staining (lanes 1-5) or by immunoblot analysis (lane 6). Lanes 1 and 2 show the total proteins of E. coli BL21(DE3)pLysS transformed with pRSetS3SH2 before (lane 1) and after (lane 2) induction with isopropyl-beta -D-thiogalactopyranoside. Lane 3 shows the supernatant of the bacterial lysate after removal of insoluble cell fragments by centrifugation. Repeated sonication of the harvested cell fragments (lane 4) yielded purified inclusion bodies (lane 5). Proteins of the inclusion bodies were resolved by SDS-PAGE, transferred to an Immobilon membrane, and detected with a monoclonal MRGSHis antibody (lane 6). The apparent molecular mass is in good agreement with the calculated mass of 15 kDa. B, reverse phase HPLC-purified STAT3-SH2 domain was refolded by dialysis. For purity determination, the proteins were resolved by SDS-PAGE. Panel 1 shows silver staining of the refolded STAT3-SH2 domain. For quantification, the polyacrylamide gel was stained with SyproTM Orange reagent and analyzed using a Storm 840 scanner. The quantitative analysis (panel 2) proved the protein to be more than 99% pure (peak a) with the sole detectable impurity being a disulfide bonded STAT3-SH2 dimer (peak b).

Refolding and CD Spectroscopy-- The purified protein was dissolved in 6 M GdnHCl, 1 mM EDTA, and 100 mM dithiothreitol and dialyzed for refolding against buffers of pH 7.5, 5.5, and 4.5, respectively. Subsequently the protein samples were characterized by CD spectroscopy. Fig. 2 shows the far UV and near UV CD spectra of the STAT3-SH2 domain at pH 7.5 (solid line) and pH 4.5 (dashed line). Although the far UV CD spectra are remarkably different at the two pH values, they look similar to spectra of other SH2 domains (23, 24). Even more pronounced differences were detected between the near UV CD spectra at the two different pH values (Fig. 2B). For instance, at pH 4.5 a distinct band appeared at 292 nm which can be assigned to a tryptophan residue. This effect can be attributed to a local change rather than to a change of the overall fold of the protein. The reversibility of this structural change was determined by changing the pH of the solution from pH 4.5 to 7.5 and vice versa (data not shown).


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Fig. 2.   CD spectra of the refolded SH2 domain. The figure shows the far UV (A) and near UV (B) CD spectra of the refolded STAT3-SH2 domain at pH 7.5 (solid line) and pH 4.5 (dashed line).

To determine the thermal stability of the folded proteins at the different pH values we recorded a series of CD spectra with increasing temperature. At pH 7.5 a melting curve with a midpoint around 43 °C was obtained (Fig. 3). At pH 4.5, however, the protein precipitated with increasing temperature (data not shown).


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Fig. 3.   Thermal stability of the refolded STAT3-SH2 domain at pH 7.5. Thermal stability was determined by CD spectroscopy. The graph shows the ellipticity Theta MRW (MRW, mean residue weight) at 215 nm as a function of temperature.

Unfolding by GdnHCl-- Because the thermal stability of the protein could only be estimated at pH 7.5, we monitored the GdnHCl-induced unfolding of the protein by fluorescence spectroscopy at pH 4.5, 5.5, and 7.5. With increasing GdnHCl concentration the fluorescence intensity decreased (not shown), and the maximum of the tryptophan emission shifted from 333 to 353 nm at pH 5.5. This behavior is characteristic of a transition of a protein from a folded to an unfolded state. Whereas the midpoint of the denaturation curve at pH 7.5 (Fig. 4, solid line) was found at about 1.7 M GdnHCl, this point is shifted to 2.1 M GdnHCl at acidic pH values of 5.5 (dotted line) and 4.5 (dashed line).


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Fig. 4.   Unfolding of the recombinant STAT3-SH2 domain. The figure shows unfolding of the STAT3-SH2 domain by increasing concentrations of GdnHCl at pH 4.5 (, dashed line), pH 5.5 (open circle , dotted line) and pH 7.5 (triangle , solid line). The wavelength shift of maximum tryptophan emission lambda max was monitored as a function of GdnHCl concentration. Because lambda max also changes with pH, the data were normalized. The lambda max values observed at GdnHCl concentrations of 0 and 4 M were taken as representative of the 100% native and 0% native, i.e. fully denatured state, respectively.

Fluorescence Spectroscopy-- Steady-state fluorescence, fluorescence lifetime, and anisotropy decay measurements were performed at pH 7.5, 5.5, and 4.5. The steady-state fluorescence data of the SH2 domain upon excitation at 295 nm are compiled in Table I. With a pH decrease from 7.5 to 4.5, the maximum of the emission band shifted from 336 to 330. This shift was accompanied by a decrease of the full width at half maximum from 56 to 51 nm. The shift of the emission maximum and the decrease of the full width at half maximum are indicative of an increasingly hydrophobic environment of the sole tryptophan present in the SH2 domain.

                              
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Table I
Fluorescence-steady state data of the STAT3-SH2 domain at different pH values
lambda max is the wavelength of the maximum tryptophan fluorescence. FWHM is the full width at half maximum

The results of the fluorescence lifetime and anisotropy decay measurements are compiled in Tables II and III. The decays of the tryptophan fluorescence can be fitted by a sum of three exponentials with fractional intensities Bi and corresponding lifetimes tau i and lead to calculated mean lifetimes (<tau >) of 4.7, 4.1, and 3.7 ns for the pH values of 7.5, 5.5, and 4.5, respectively (Table II). Such a behavior is in good agreement with the blue shift of the emission maximum in Table I (25).

                              
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Table II
Fluorescence intensity decay data of the STAT3-SH2 domain at different pH values
Bi is the fractional amplitude associated with the decay time tau i; < tau > is the mean lifetime (see "Experimental Procedures").

The fluorescence anisotropy decays of the SH2 domain at different pH values were fitted with one exponential and led to rotational correlation times of Phi  = 12.4, 6.4, and 6.1 ns at pH values of 7.5, 5.5, and 4.5, respectively (Table III). Rotational correlation times Phi  can be used to calculate the molecular mass from the equation Mr = f × Phi  (f = 2.6 kDa/ns) for spherical particles on the basis of the Stokes-Einstein relationship (26). The expected rotational correlation time Phi  for a monomeric SH2 domain is therefore about 5.7 ns assuming a spherical shape. The measured rotational correlation times at acidic pH values are in good agreement with the overall tumbling rate expected for a monomer. The twice as high value found at neutral pH indicates the existence of a dimeric SH2 domain. The reversibility of the monomer/dimer transition was determined by changing the pH from 4.5 to 7.5 and vice versa by dialysis (data not shown).

                              
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Table III
Fluorescence anisotropy decay data of the STAT3-SH2 domain at different pH values
ri are the anisotropies, Phi  the rotational correlation time (see "Experimental Procedures").

Size Exclusion Chromatography-- Additional evidence for the dimerization was provided by size exclusion chromatography experiments (Fig. 5) using a calibrated column. Fig. 5A shows the elution profile of the SH2 domain (monomer, dotted line; dimer, solid line). Both peaks contain STAT3-SH2 as shown by SDS-PAGE (see inset in Fig. 5A). The apparent molecular masses of both the monomer (13 kDa) and the dimer (32 kDa) are in good agreement with the expected values (Fig. 5B) of 15 and 30 kDa, respectively.


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Fig. 5.   Size exclusion chromatography. A, elution profile of the STAT3-SH2 domain under monomeric (pH 5.5, dotted line) and dimeric (pH 7.5, solid line) conditions. The elution profile of the marker proteins (a, bovine thyroglobulin, 670 kDa; b, bovine gamma -globulin, 158 kDa; c, chicken ovalbumin, 44 kDa; d, horse myoglobin, 17 kDa; e, vitamin B12, 1.35 kDa) is drawn as a dashed line. triangle  and open circle  denote the STAT-SH2 domain eluted at pH 7.5 and 5.5, respectively. The inset displays a SDS-PAGE analysis of fractions 3-13 showing STAT3-SH2. Protein bands were visualized by silver staining. The marker proteins b, c, and d were used for linear regression. B, plot of the log(Mr) as a function of the retention times of the marker proteins (a-d). Elution times of the STAT3-SH2 domain at pH 7.5 (triangle ) and 5.5 (open circle ) correspond to molecular masses of 32 and 13 kDa, respectively.

Specific Interaction of the Recombinant STAT3-SH2 Domain with Phosphopeptides-- To determine the functionality of the recombinant STAT3-SH2 domain, we worked out an ELISA using biotinylated phosphopeptides as bait. Based on previous investigations, we chose all of the phosphopeptide motifs of the signal transducer gp130, a mutant of the pY767 motif containing a Q/E exchange at the pY+3 position (pYQ770E) as well as the phosphotyrosine motif encompassing pY705 of STAT3 or a peptide containing the amino acids of the gp130 motif pY767 in random order (pYX) (Table IV). As the STAT3-SH2 domain turned out to undergo a pH-dependent dimerization, we investigated the specificity of the interaction with the various phosphopeptide motifs under neutral (dimer) and acidic (monomer) conditions. For the interaction of the peptides with the monomeric SH2 domain, we performed the assay at pH 5.5 to maintain the stability of streptavidin.

                              
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Table IV
Biotinylated peptides used in the ELISA

Fig. 6 shows a schematic representation of the ELISA used. After incubation of the biotinylated phosphopeptides with the streptavidin-coated surface, the immobilized phosphotyrosine residues were detected with the phosphotyrosine antibody PY20 (Fig. 6A). Incubation with increasing amounts of STAT3-SH2 led to a decrease in absorbance because PY20 was unable to recognize the phosphopeptides bound to the SH2 domain (Fig. 6B). The relative decrease in absorbance with increasing amounts of STAT3-SH2 compared with an SH2-free sample was used to determine the inhibition of the PY20/phosphopeptide interaction by STAT3-SH2 which indicates SH2/peptide binding.


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Fig. 6.   Schematic representation of the peptide binding assay. A, detection of the biotinylated phosphopeptides immobilized on a streptavidin surface by the phosphotyrosine antibody PY20. B, inhibition of the PY20/peptide interaction by STAT3-SH2 bound to the phosphotyrosine motifs.

Fig. 7A shows the inhibition of the PY20/phosphopeptide interaction at pH 5.5. The STAT3-SH2 domain interacts specifically with the four-membrane distal phosphotyrosine motifs of gp130 (pY767, pY814, pY905, and pY915) whereas the peptides pY683 and pYX as well as the pY759 motif, known to bind to the SH2 domain of SHP-2, show only weak binding. In addition, the isolated STAT3-SH2 domain binds to the motif pY705 of STAT3. Interestingly, the binding of the SH2 domain to pY767 is significantly impaired by a Q/E exchange at the pY+3 position (pYQ770E). This emphasizes the importance of the pY+3 position for specific recognition of receptor motifs by STAT3-SH2.


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Fig. 7.   Inhibition of the PY20/phosphopeptide interaction by STAT3-SH2. Binding of the STAT3-SH2 domain to the phosphopeptides listed in Table IV was determined by monitoring the inhibition of the PY20/peptide interaction with increasing concentrations of STAT3-SH2 at pH 5.5 (A) and pH 7.5 (B).

Fig. 7B shows the results of the same experiment under neutral buffer conditions (pH 7.5) where the SH2 domain exists as a dimer. Whereas the monomeric SH2 domain is able to distinguish between the different phosphopeptides, the dimeric molecule is not. The phosphopeptides show only low affinity to the dimer.

    DISCUSSION
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Abstract
Introduction
Procedures
Results
Discussion
References

The SH2 domains of the STAT factors are involved in receptor recognition as well as in their dimerization. Dimerization is induced by tyrosine phosphorylation and is a prerequisite for DNA binding. To investigate the interaction of the SH2 domains with the respective phosphotyrosine motifs on a molecular level we expressed the SH2 domain of STAT3 in E. coli. After purification the protein was refolded. CD spectra of the refolded protein correspond to those observed with other SH2 domains (23, 24). Spectral differences were observed at neutral and acidic pH. The changes seen in the far UV are a reflection of limited rearrangements of the overall structure (Fig. 2). The near UV spectrum, for instance, shows a distinct band at 292 nm at pH 4.5 which is not detectable at pH 7.5, indicating a loss in conformational mobility for the side chain of the sole tryptophan. This result correlates well with the corresponding fluorescence spectra where a blue shift of the emission maximum and a decrease of the full width at half maximum are observed with decreasing pH, reflecting a transition of the sole tryptophan from a hydrophilic to a more hydrophobic environment.

The fluorescence anisotropy decay measurements enabled us to assign these spectral differences to a pH-dependent dimerization of the recombinant SH2 domain which exists as a monomer under acidic conditions and as a dimer at neutral pH. The less exposed tryptophan in the monomeric state correlates with a higher stability of the monomer compared with the dimer as revealed by GdnHCl-induced denaturation. Thus, the small structural changes induced by dimerization are accompanied by a destabilization of the molecule.

Taken together, the fluorescence and CD measurements show that dimerization leads to a conformational change in the SH2 domain involving tryptophan 623. In the x-ray structure of the (STAT1)2-DNA complex the corresponding tryptophan is located at the surface of the molecule and is accessible to water. The higher B factors of this amino acid residue in the crystal structure are a further indication of its enhanced flexibility in the dimer. For other SH2 domains such as those of Src or Lck kinase it has been shown that the BG loop is attached to the body of the molecule (27, 28), whereas it is completely detached in the (STAT3)2-DNA and (STAT1)2-DNA complexes (14, 15). Because the BG loop is also involved in the dimer interface we raise the idea that in the monomeric state this loop resembles the situation seen in other SH2 domains. This would bury the tryptophan (Trp623) within the structure, a fact that we indeed observe for the monomeric SH2 domain. The two conformational states might reflect different modes of SH2/phosphotyrosine peptide interactions in STAT receptor binding and STAT dimer formation.

Recently, heterodimeric complexes of STAT1 with STAT2 or STAT3 prior to cytokine stimulation have been described (29). The ability of the STAT3-SH2 domain to form dimers may reflect such an interaction between unphosphorylated STAT molecules. On the other hand, it cannot be ruled out that the observed dimerization is a property of the isolated domain and that its formation is prevented within the entire protein.

To study the interaction of the STAT3-SH2 domain with different phosphotyrosine peptides we established an ELISA based on the competition of a phosphotyrosine monoclonal antibody with the recombinant protein in binding to phosphotyrosine peptide motifs. Whereas the monomeric STAT3-SH2 domain was able to bind to specific phosphotyrosine peptides no such interaction could be observed with the dimeric protein.

We detected a specific interaction between the monomeric STAT3-SH2 domain and the four distal phosphotyrosine motifs present in the cytoplasmic part of the signal transducer gp130. In contrast, the phosphotyrosine peptides corresponding to the two membrane-proximal tyrosine residues did not show a specific interaction (Fig. 7). These results are in good agreement with the previous observation that in transiently transfected COS cells STAT3 is activated only through the four distal tyrosine motifs of gp130 (6, 7). Interestingly, we found a Q770E substitution in the Y767 motif of gp130 to lead to a loss in STAT3-SH2 binding, corroborating the finding that a glutamine residue at the Y+3 position is important for STAT3 activation (Fig. 7). Thus, our STAT3-SH2/phosphopeptide interaction studies fully confirmed the results obtained with native STAT3 in COS cells. Furthermore, the phosphotyrosine peptide of STAT3 itself (pY705) showed specific binding to the recombinant STAT3-SH2 domain. A comparison of the affinities would require a common binding mechanism. As deduced from the x-ray structure of the (STAT1)2-DNA and (STAT3)2-DNA complexes the mode of SH2/peptide binding therein shows fundamental differences to SH2/peptide interactions known so far (2). An expected higher affinity to the phosphotyrosine peptide of STAT3 itself (pY705) compared with receptor motifs was not observed. This is presumably because the complex interaction seen in the x-ray structures cannot be reconstituted in the ELISA.

Thus far, structure/function studies of recombinant STAT-SH2 domains were hampered by the fact that they did not show specific binding to phosphotyrosine peptides. We found that at physiological pH the recombinant STAT3-SH2 domain is forming dimers that do not bind to phosphopeptides. The observation that under acidic conditions, the STAT3-SH2 domain exists as a monomer that specifically binds to phosphotyrosine motifs will enable us to elucidate how STAT factors interact with their receptors.

    ACKNOWLEDGEMENTS

We thank Jürgen Stahl for the CD measurements, Michael Weske for the fluorescence measurements, and Gerhard Müller-Newen for helpful discussions.

    FOOTNOTES

* This work was supported by the Deutsche Forschungsgemeinschaft (Bonn) and the Fonds der Chemischen Industrie (Frankfurt/Main).The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

To whom correspondence should be addressed: Institut für Biochemie, RWTH-Aachen, Pauwelsstrasse 30, D-52074 Aachen, Germany. Tel.: 49-241-808-8830; Fax: 49-241-888-8428; E-mail: heinrich{at}rwth-aachen.de.

The abbreviations used are: SH2, Src homology 2; STAT, signal transducer and activator of transcription; GdnHCl, guanidine hydrochloride; HPLC, high performance liquid chromatography; PAGE, polyacrylamide gel electrophoresis; ELISA, enzyme-linked immunosorbent assay; pY, phosphotyrosine.
    REFERENCES
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Abstract
Introduction
Procedures
Results
Discussion
References

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