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• W2023454712 abstract "To probe the covalent serpin-proteinase complex, we used wild-type and 4 new single cysteine variants (T85C, S121C, D159C, and D298C) of α1-proteinase inhibitor Pittsburgh. Cysteines in each variant could be labeled both in native and proteinase-complexed α1-proteinase inhibitors. Pre-reaction with 7-nitrobenz-2-oxa-1,3-diazole-chloride or fluorescein prevented complex formation only with the D298C variant. Label at Cys121 greatly increased the stoichiometry of inhibition for thrombin and gave an emission spectrum that discriminated between native, cleaved, and proteinase-complexed serpin and between complexes with trypsin and thrombin, whereas fluorophore at residue 159 on helix F was almost insensitive to complex formation. Fluorescence resonance energy transfer measurements for covalent and non-covalent complexes were consistent with a location of the proteinase at the end of the serpin distal from the original location of the reactive center loop. Taken together, these findings are consistent with a serpin-proteinase complex in which the reactive center loop is fully inserted into β-sheet A, and the proteinase is at the far end of the serpin from its initial site of docking with the reactive center loop close to, but not obscuring, residue 121. To probe the covalent serpin-proteinase complex, we used wild-type and 4 new single cysteine variants (T85C, S121C, D159C, and D298C) of α1-proteinase inhibitor Pittsburgh. Cysteines in each variant could be labeled both in native and proteinase-complexed α1-proteinase inhibitors. Pre-reaction with 7-nitrobenz-2-oxa-1,3-diazole-chloride or fluorescein prevented complex formation only with the D298C variant. Label at Cys121 greatly increased the stoichiometry of inhibition for thrombin and gave an emission spectrum that discriminated between native, cleaved, and proteinase-complexed serpin and between complexes with trypsin and thrombin, whereas fluorophore at residue 159 on helix F was almost insensitive to complex formation. Fluorescence resonance energy transfer measurements for covalent and non-covalent complexes were consistent with a location of the proteinase at the end of the serpin distal from the original location of the reactive center loop. Taken together, these findings are consistent with a serpin-proteinase complex in which the reactive center loop is fully inserted into β-sheet A, and the proteinase is at the far end of the serpin from its initial site of docking with the reactive center loop close to, but not obscuring, residue 121. Serpins are a family of widely distributed, structurally homologous proteins (1Hunt L.T. Dayhoff M.O. Biochem. Biophys. Res. Commun. 1980; 95: 864-871Crossref PubMed Scopus (317) Google Scholar), many of which are inhibitors of serine proteinases (2Carrell R.W. Travis J. Trends Biol. Sci. 1985; 10: 20-24Abstract Full Text PDF Scopus (329) Google Scholar). Whereas the many other families of protein inhibitors of serine proteinases, such as the Bowman-Birk, Kazal, and Kunitz families, inhibit target proteinases by forming tight non-covalent 1:1 complexes in which neither the proteinase nor the inhibitor undergoes significant structural change in most cases (3Bode W. Huber R. Eur. J. Biochem. 1992; 204: 433-451Crossref PubMed Scopus (1009) Google Scholar), serpins differ not only by apparently forming covalent 1:1 acyl enzyme complexes with their target proteinases (4Lawrence D.A. Ginsburg D. Day D.E. Berkenpas M.B. Verhamme I.M. Kvassman J.-O. Shore J.D. J. Biol. Chem. 1995; 270: 25309-25312Abstract Full Text Full Text PDF PubMed Scopus (229) Google Scholar), but by undergoing a major conformational change during, and as an essential part of, the inhibition process (5Gettins P. Patston P.A. Schapira M. BioEssays. 1993; 15: 461-467Crossref PubMed Scopus (96) Google Scholar). Because of the requirement for conformational change as part of the inhibition mechanism, knowledge of the structure of the serpin-proteinase complex is critical for an understanding of how serpins inhibit their target proteinases through kinetic trapping of a normal covalent acyl enzyme intermediate on the proteinase substrate cleavage pathway. A previous proposal that a major movement of the proteinase occurs following cleavage of the scissile bond (6Wright H.T. Scarsdale J.N. Proteins. 1995; 22: 210-225Crossref PubMed Scopus (157) Google Scholar) has been supported by two recent studies (7Stratikos E. Gettins P.G.W. Proc. Natl. Acad. Sci. U. S. A. 1997; 94: 453-458Crossref PubMed Scopus (139) Google Scholar, 8Wilczynska M. Fa M. Karolin J. Ohlsson P.I. Johansson L.B. Å. Ny T. Nat. Struct. Biol. 1997; 4: 354-357Crossref PubMed Scopus (138) Google Scholar). In one study (8Wilczynska M. Fa M. Karolin J. Ohlsson P.I. Johansson L.B. Å. Ny T. Nat. Struct. Biol. 1997; 4: 354-357Crossref PubMed Scopus (138) Google Scholar) chemical cross-linking between the proteinase and the serpin in the complex, together with a measurement of the separation between P3 and P1′ residues of the serpin in the complex by fluorescence resonance energy transfer, was consistent with a location of the proteinase half-way down the flank of the serpin (Fig. 1) and in contact with helix F. The other study (7Stratikos E. Gettins P.G.W. Proc. Natl. Acad. Sci. U. S. A. 1997; 94: 453-458Crossref PubMed Scopus (139) Google Scholar), from this laboratory, used fluorescence resonance energy transfer between fluorophores on the serpin α1-proteinase inhibitor (α1PI) 1The abbreviations used are: α1PI, α1-proteinase inhibitor; 5-IAF, 5-iodoacetamidofluorescein; NBD, 7-nitrobenz-2-oxa-1,3-diazole; SI, stoichiometry of inhibition, defined as the number of moles of serpin required to inhibit 1 mol of proteinase by formation of SDS-stable complex; TLCK, tosyl-lysyl chloromethyl ketone; PAGE, polyacrylamide gel electrophoresis; Bis-Tris, 2-[bis(2-hydroxyethyl)amino]-2-(hydroxymethyl)-propane-1,3-diol. Pittsburgh and the proteinase to compare the inter-fluorophore separation in the normal covalent serpin-proteinase complex with that in the non-covalent complex with the non-functional anhydroproteinase. This study, although not able to precisely define the position of the proteinase in the complex, demonstrated a movement of the proteinase of at least 21 Å upon formation of the kinetically trapped covalent complex. We describe here more extensive mapping of this serpin-proteinase complex by using wild-type α1PI Pittsburgh and 4 new single cysteine variants. These well separated cysteines were used as follows: (i) to probe the accessibility of the cysteine in native and proteinase-complexed serpin, (ii) to determine the effect of derivatization of the cysteine on the ability to form covalent complex, and (iii) for introduction of fluorophores, both as probes of the local environment and for fluorescence resonance energy transfer measurements. By these approaches we have been able to place further constraints on the possible structures of the serpin-proteinase complex and to show that it probably requires movement of the proteinase to the bottom of the serpin and therefore full insertion of the cleaved reactive center loop into β-sheet A. In this location the proteinase is not in contact with the outer face of helix F. Our findings are thus consistent with the model of Wright and Scarsdale (6Wright H.T. Scarsdale J.N. Proteins. 1995; 22: 210-225Crossref PubMed Scopus (157) Google Scholar). Site-directed mutagenesis was carried out on a double-stranded pET16b plasmid (Novagen) containing α1PI cDNA, using the Quikchange method (Stratagene). Double-stranded template DNA of two complementary primers containing the mutation was annealed and extended with Pfu DNA polymerase during thermal cycling. The pET16b plasmid contained an N-terminally modified α1PI cDNA that lacked coding sequence for the first 5 residues (10Johansen H. Sutiphong J. Sathe G. Jacobs P. Cravador A. Bollen A. Rosenberg M. Shatzman A. Mol. Biol. & Med. 1987; 4: 291-305PubMed Google Scholar), inserted between theNcoI and BamHI subcloning sites of the vector. The sequences for the coding strands of the mismatch primers were as follows (mismatch codons are underlined): M358R, 5′-GAG GCC ATA CCCAGA TCT ATC CCC CCC; C232S, 5′-AAC ATC CAG CACAGC AAG AAG CTG TCC AG; T85C, G AAT TTC AAC CTCTGT GAG ATT CCG GAG G; S121C, GGC CTG TTC CTCTGT GAG GGC CTG AAG; D159C, G AAA CAG ATC AACTGT TAC GTG GAG AAG GG; D298C, AC TGG AAC CTA TTGTCT GAA GAG CGT CCT G. All α1PI variants used in this study carried the Pittsburgh (M358R) mutation, and all except the wild-type Pittsburgh variant carried the C232S mutation, so that all variants contained only one free cysteine residue and all contained a P1 arginine (Pittsburgh mutation (11Owen M.C. Brennan S.O. Lewis J.H. Carrell R.W. N. Engl. J. Med. 1983; 309: 694-698Crossref PubMed Scopus (323) Google Scholar, 12Scott C.F. Carrell R.W. Glaser C.B. Kueppers F. Lewis J.H. Colman R.W. J. Clin. Invest. 1986; 77: 631-634Crossref PubMed Scopus (70) Google Scholar, 13Lewis J.H. Iammarino R.M. Spero J.A. Hasiba U. Blood. 1987; 51: 129-137Crossref Google Scholar)) that conferred high affinity (5 nm) for the non-covalent complexes with anhydrotrypsin (7Stratikos E. Gettins P.G.W. Proc. Natl. Acad. Sci. U. S. A. 1997; 94: 453-458Crossref PubMed Scopus (139) Google Scholar). All mutations were confirmed by dideoxy sequencing in the host plasmid. The plasmids containing the mutated α1PI genes were transformed into BL21 (DE3) cells (Novagen). 5 ml of an overnight culture was used to inoculate 0.5 liters of LB broth containing 125 mg/ml ampicillin. Cells were grown at 37 °C in a shaking water bath to an A 600 nm of 0.5. IPTG (0.4 mm) was added to induce expression of α1PI, and the cells were allowed to grow for a further 2.5–3 h at 37 °C. Cells were harvested by centrifugation at 4000 × g for 10 min, and the cell pellet was collected and washed twice with cold PBS. The cell pellet was resuspended in 10 ml of buffer A (50 mm Tris, pH 8.0, 50 mm NaCl, and 1 mm EDTA) containing 1 mmphenylmethylsulfonyl fluoride, 1 mm β-mercaptoethanol, and 0.2 mg/ml lysozyme. The suspension was sonicated using four 30-s pulses. Inclusion bodies were harvested by spinning the lysed cells at 10,000 × g for 45 min. The pellet containing the inclusion bodies was washed twice with buffer A containing 0.5% Triton X-100 (Sigma). The inclusion bodies were finally dissolved in 10 ml of buffer A containing 8 m guanidinium hydrochloride by vortexing and sonication. Any non-solubilized inclusion bodies were spun down at 10,000 × g for 15 min. The typical yield of solubilized protein at this stage was ∼20 mg, determined by BCA assay, from 0.5-liter cell culture. Solubilized denatured α1PI was refolded by dropwise dilution over a period of 20 min of the 8 m guanidinium hydrochloride solution into 250 ml of a 4 °C solution of 10 mm sodium phosphate buffer, pH 6.5, containing 1 mm EDTA and 1 mmdithiothreitol. The diluted solution was extensively dialyzed against 10 mm sodium phosphate buffer, pH 6.5, containing 1 mm EDTA to remove the guanidinium hydrochloride. After dialysis the solution was centrifuged for 30 min at 4 °C and 8000 × g and filtered through a 0.2-μm filter to remove any precipitated protein. The refolded α1PI contained dimers and higher order oligomers in addition to monomers. Monomeric α1PI was purified in two steps. The first step was ion exchange chromatography on DE52 (Whatman), using a linear 0–0.25 m NaCl gradient. This gave a sharp early eluting peak for the monomer and a broader later eluting peak for higher order aggregates. The pooled monomeric fractions were dialyzed against 20 mm Bis-Tris-propane buffer, pH 6.5, containing 1 mm EDTA and rechromatographed, where necessary, on a MonoQ column equilibrated in the same buffer and eluted by a 0–0.3 m NaCl gradient. Monomeric α1PI eluted as a very sharp single peak. The purity of all preparations was confirmed by SDS-PAGE and by PAGE under non-denaturing conditions to confirm that all material was monomeric. Preparations were dialyzed against 20 mm sodium phosphate, pH 7.4, 100 mm NaCl, 0.1 mm EDTA, 0.1% PEG8000, quickly frozen in aliquots, and stored at −70 °C until needed. Anhydrotrypsin was prepared from commercial crystallized trypsin (Sigma) by alkaline β-elimination of the phenylmethylsulfonyl fluoride adduct according to published procedures (14Ako H. Foster R.J. Ryan C.A. Biochem. Biophys. Res. Commun. 1972; 47: 1402-1407Crossref PubMed Scopus (71) Google Scholar). Following the reaction, the solution was treated with Phe-Phe-Arg-chloromethyl ketone (20 μm) to inhibit any remaining or regenerated active trypsin and acidified to pH 3.0. β-Anhydrotrypsin was purified from the reaction mixture by chromatography on a soybean trypsin inhibitor affinity matrix. The absence of proteolytic activity in the product was confirmed by activity assay using the chromogenic trypsin substrate S-2222. β-Trypsin was prepared from TPCK-treated commercial trypsin by affinity chromatography using the same soybean trypsin inhibitor affinity matrix. All α1PI variants were labeled with fluorescein by reaction of the single free cysteine with 5 iodoacetamido-fluorescein (Molecular Probes, Eugene, OR). The protein (10–40 μm) was reacted with a 2-fold molar excess of dithiothreitol for 15 min at room temperature and then with a 10–15-fold molar excess of 5-IAF. The reaction was allowed to proceed overnight at 4 °C. Excess reagent was removed by dialysis for 24 h against 10,000 volumes of 20 mm sodium phosphate, pH 7.4, 100 mm NaCl, 0.1 mm EDTA, 0.1% PEG8000. The extent of labeling was determined spectrophotometrically using the absorbance at 495 nm for determination of the fluorescein concentration and the absorbance at 280 nm, corrected for the contribution from fluorescein at this wavelength, which was determined to be 25% of the absorbance at 495 nm based on the spectrum of the adduct of IAF with β-mercaptoethanol, to determine the protein concentration. For all preparations the extent of labeling was close to 1 eq per mol or less, with a range from 0.59 to 1.06. This range represents the determined stoichiometries for preparations made at different times under somewhat different reactant concentrations and does not necessarily reflect intrinsic differences in reactivity of the various cysteines. Where comparisons of labeling efficiency are made elsewhere, reactions were carried out under identical conditions for each α1PI species. Extinction coefficients of 27,000 (15Pannell R. Johnson D. Travis J. Biochemistry. 1974; 13: 5439-5445Crossref PubMed Scopus (201) Google Scholar) and 82,000 m−1 cm−1 were used for α1PI and fluorescein, respectively. α1PI variants were labeled with NBD by reaction with NBD-chloride. α1PI, 10–20 μm, in 20 mm sodium phosphate buffer, pH 7.4, containing 100 mm NaCl, 0.1 mm EDTA, and 0.1% PEG8000, was first reduced by addition of dithiothreitol in slight molar excess and incubation at room temperature for 15 min. A 10-fold excess of NBD-chloride was added and the reaction allowed to proceed overnight at 4 °C in the dark. The sample was then dialyzed against the same buffer for 24 h at 4 °C. The extent of labeling was calculated spectrophotometrically using the extinction coefficient of NBD at 420 nm of 13,000 m−1 cm−1 and an extinction coefficient for the protein at 280 nm of 27,000m−1 cm−1. The contribution of NBD at 280 nm is small. Labeling stoichiometries of 0.8–1.0 were obtained. β-Trypsin and β-anhydrotrypsin were labeled with tetramethylrhodamine isothiocyanate while immobilized on soybean trypsin inhibitor-agarose beads (i) to permit equivalent reaction conditions for anhydrotrypsin as for trypsin without concern for autodigestion by free trypsin, and (ii) to provide a ready means of selecting only those labeled proteins that were still active in binding to protein inhibitors. About 300 μl of wet soybean trypsin inhibitor-agarose beads were equilibrated with 0.1 m sodium citrate buffer, pH 4.0, and then mixed with 700 μl of either β-trypsin or β-anhydrotrypsin, followed by gentle rotation for 30 min at 4 °C. The beads were washed twice with 700 μl of 0.1 m citrate buffer, pH 4.0, to remove any unbound protein and then four times with 700 μl of 0.1 msodium carbonate, pH 9.0, to raise the pH. The beads were resuspended in 700 μl of 0.1 m sodium carbonate, pH 9.0, and 10 μl of a 10 mm solution of tetramethylrhodamine isothiocyanate in N,N-dimethylformamide were added. The reaction was allowed to proceed, with gentle rocking, at room temperature for 2–4 h, depending on the degree of labeling wanted. The beads were then washed four times with 700 μl of 10 mm sodium carbonate, pH 9.0, to remove excess reagent and any free protein. Labeled protein was eluted with 500 μl of 0.2 m sodium citrate, pH 2.4. The eluate was dialyzed overnight against 1000 volume of 1 mm HCl containing 10 mm CaCl2 and centrifuged at 14,000 × g for 10 min to remove any precipitated material. The extent of labeling was determined spectrophotometrically using extinction coefficients of 62,000m−1 cm−1 at 550 nm and 35,800m−1 cm−1 at 280 nm for tetramethylrhodamine and trypsin, respectively. The absorbance at 280 nm was first corrected for the contribution at that wavelength from tetramethylrhodamine, which was found empirically to be 28% of the absorbance at 550 nm. The extent of label incorporation was 0.47 mol/mol for trypsin and, for two separate preparations of anhydrotrypsin, 0.30 and 0.71 mol/mol. The sites of labeling in β-trypsin were identified by N-terminal sequencing of tetramethylrhodamine-labeled peptides isolated from a tryptic digest of the labeled protein. 140 μg of labeled β-trypsin was freeze-dried and dissolved in 50 μl of 8m guanidinium hydrochoride, 50 mm Tris, pH 8.0, 50 mm NaCl, 1 mm EDTA. Dithiothreitol was added to 10 mm and the mixture incubated at room temperature for 30 min. Iodoacetic acid was added to 30 mm and allowed to react for 30 min at room temperature. The denatured labeled trypsin was diluted into 600 μl of 50 mm Tris, pH 8.0, 50 mm NaCl, 1 mm EDTA, 10 mmCaCl2 containing 5 μg of active β-trypsin and digested in the dark at 37 °C for 2 h, after which an additional 5 μg of β-trypsin was added. A second addition was made after a further 2 h, and the reaction was allowed to continue overnight. The tryptic digest was chromatographed on a C-18 reverse phase column, using a linear gradient from 80% buffer A (0.1% trifluoroacetic acid in water), 20% buffer B (0.1% trifluoroacetic acid, 90% acetonitrile, 9.9% water) to 35% buffer A, 65% buffer B. Peaks were monitored both for absorption at 280 nm and fluorescence at 550 nm. Two major peaks of approximately equal fluorescent intensity were obtained, as well as several much smaller ones, and were submitted for N-terminal sequence determination. The ability of the α1PI variants to form SDS-stable covalent complex with trypsin was assayed by 10% SDS-PAGE of the reaction products and visualization of the complex either by Coomassie staining for unlabeled complex or by fluorescence intensity for labeled complex. Typically 2–3 μg of α1PI was reacted for 10 s with trypsin at different molar ratios, ranging from 0.3:1 to 2:1 trypsin:α1PI, with α1PI fixed at 5–10 μm. This was sufficient time for the reaction to have gone to >99% completion based on the published rate constant for this reaction (7Stratikos E. Gettins P.G.W. Proc. Natl. Acad. Sci. U. S. A. 1997; 94: 453-458Crossref PubMed Scopus (139) Google Scholar) and confirmed empirically by the absence of unreacted serpin in lanes where the proteinase was in excess. This also confirmed that the serpin was >95% active. The stoichiometry of inhibition (SI) was calculated by scanning densitometry of SDS-PAGE gels. Coomassie Blue-stained gels were scanned, and the density of the bands corresponding to cleaved serpin and complex were measured. The intensity of the band for complex was corrected for the contribution from the proteinase, by assuming equal staining of the serpin and trypsin per unit weight. This was justified by a standard curve for trypsin and α1PI which showed comparable staining for both proteins on a weight basis and a linear dependence between amount of protein and band intensity in the range used for the experiments. This method was considered accurate for SI values in the range 1.1 to 5, corresponding to 91 to 20% complex, but incapable of determining SI where complex bands were so faint as to be not visible. SI values for fluorescein-labeled serpins were determined in an analogous way, except intensity of the fluorescent bands corresponding to cleaved and complexed bands were used, and no correction was needed for contribution from (unlabeled) trypsin. No error was thereby introduced for complex formed by unlabeled serpin. In cases where the degree of fluorescein labeling was close to 100%, so that all covalent complex was also fluorescent, or where labeling did not affect complex formation, independent quantitation of SI by both fluorescence and Coomassie Blue staining gave good agreement. 5–6 μg of either the Pittsburgh variant of α1PI or the cysteine mutants (T85C, D159C, S121C and D298C) in a total volume of 20 μl were reacted with 1 μg of β-trypsin (excess of α1PI to ensure that complex was not degraded by excess proteinase) to form the stable trypsin-α1PI complex. TLCK was added after a few seconds to a final concentration of 25 μm. In all reaction mixtures 1.5 μl of 1.8 mm IAF was added (final concentration was about 100 μm), and the reaction was allowed to proceed for 2 h at 4 °C. Dithiothreitol was added to a final concentration of 1 mm, and the mixture was incubated at room temperature for 10 min (to inactivate any unreacted probe). The samples were then subjected to SDS-PAGE analysis (12% acrylamide). A control reaction of IAF with TLCK-treated trypsin alone was carried out and showed no labeling of trypsin under the conditions used. Similar reactions were also carried out with thrombin as the proteinase, using comparable conditions as for the reactions with β-trypsin except that reaction was carried out for 8 min at room temperature (sufficient for complete reaction of the thrombin), and Phe-Pro-Arg-chloromethyl ketone was added to inactivate any free thrombin. All fluorescence measurements were made on a SPEX fluorolog scanning fluorimeter. NBD spectra were acquired by exciting at 420 nm and scanning from 440 to 580 nm. Fluorescein and rhodamine spectra were recorded by exciting at 340 nm and scanning from 460 to 640 nm. All slit widths were 4 nm. Measurements were made at 25 °C. For time courses the emission signal was monitored at 515 nm, where the contribution of rhodamine fluorescence is negligible. For energy transfer measurements the labeled serpin was between 50 and 150 nm, and the proteinase was at 2–3 times the serpin concentration. 1 mmbenzamidine was included in the cuvette as a competitive inhibitor of trypsin to slow down the reaction. NBD spectra were acquired at concentrations of 156 nm for the S121C variant and 400 nm for the D159C variant. The buffer used for all measurements was 20 mm sodium phosphate, pH 7.4, containing 100 mm NaCl, 0.1 mm EDTA, and 0.1% PEG8000. To estimate the efficiency of energy transfer between fluorescein and rhodamine in the covalent complex, the fluorescence spectrum of the fluorescein-labeled serpin was recorded, and trypsin, either unlabeled or rhodamine-labeled, was then added to the cuvette in the presence of 1 mm benzamidine and the reaction followed by monitoring the change of fluorescein fluorescence at 515 nm. When a plateau was reached the fluorescence emission spectrum of the mixture was recorded. The amount of energy transfer for each variant was determined from the observed reduction in fluorescein fluorescence corrected for any contribution that arose solely from complex formation, which was determined from a control reaction using fluorescein-labeled α1PI and unlabeled β-trypsin. A correction was also made for the SI in every case, calculated as described above, although this was mostly a small correction. Percentages of energy transfer reported in Table III also take into account the stoichiometry of rhodamine labeling and are scaled to the efficiency expected at 1 mol/mol of label. The justification for such a scaling is that, for different degrees of labeling, a linear dependence of efficiency of transfer was found. Scaling to 1:1 treats each trypsin as having either zero or one labels and that label at any of the positions is equivalent. Although this is not likely to be strictly accurate, it should give a minimum value for the efficiency of energy transfer. Confirmation that the end point represented complete reaction of the serpin was from SDS-PAGE analysis of the trichloroacetic acid-precipitated reaction products, which also showed no evidence of cleavage of the proteinase in the complex by excess active proteinase.Table IIIFluorescence resonance energy transfer in complexes with β-trypsin or anhydrotrypsinPosition of fluoresceinFluorescence energy transferTrypsinAnhydrotrypsin%8534.51912122.5101597.71023218aFrom Ref. 7.77aFrom Ref. 7.298Not measurablebNo detectable complex formed (very high SI).<2a From Ref. 7Stratikos E. Gettins P.G.W. Proc. Natl. Acad. Sci. U. S. A. 1997; 94: 453-458Crossref PubMed Scopus (139) Google Scholar.b No detectable complex formed (very high SI). Open table in a new tab For fluorescence resonance energy transfer measurements on non-covalent complexes between anhydrotrypsin and serpin, fluorescein-labeled α1PI (30–50 nm) and rhodamine-labeled or unlabeled anhydrotrypsin (80–100 nm) were mixed and the spectra recorded. For each α1PI variant four series of spectra were recorded; labeled α1PI alone, labeled α1PI with unlabeled anhydrotrypsin, labeled α1PI with labeled anhydrotrypsin (0.3 label/mol), and labeled α1PI with labeled anhydrotrypsin (0.71 label/mol). The efficiency of energy transfer was estimated from the reduction in fluorescein emission intensity in the doubly labeled complex compared with complex with unlabeled anhydrotrypsin. The values were scaled to 1.0 label/mol and the results for the two different preparations of labeled anhydrotrypsin averaged. Spectra are the average of three consecutive scans. NBD-labeled S121C variant was reacted with trypsin, papain, or thrombin as described in the figure legend for Fig. 4. One aliquot of the reaction mixture was used for SDS-PAGE analysis, and another aliquot was diluted to a final concentration of 156 nm α1PI and the emission fluorescence spectrum recorded as described above. The Coomassie Blue-stained gel was scanned and the density of the bands used to estimate both the completeness of each reaction and the SI. Every reaction was found to be more than 95% complete. Previous studies on a recombinant Pittsburgh variant of α1PI (P1 Met → Arg) have shown that the cysteine at position 232 is quite accessible to nucleophiles (7Stratikos E. Gettins P.G.W. Proc. Natl. Acad. Sci. U. S. A. 1997; 94: 453-458Crossref PubMed Scopus (139) Google Scholar), consistent with its exposed location in the crystal structure of α1PI. We found here that cysteine 232 is also accessible in the complex, since it could be comparably labeled with 5-IAF while in complex with both β-trypsin and thrombin, as judged by the intensity of fluorescence associated with the band of complex on SDS-PAGE (Table I, gel not shown). To carry out similar accessibility studies at different sites on the serpin, we created four new variants, each containing a single free cysteine at strategic locations on the serpin surface (Fig. 1). The choice of sites was guided by proposed models for the serpin-proteinase complex (6Wright H.T. Scarsdale J.N. Proteins. 1995; 22: 210-225Crossref PubMed Scopus (157) Google Scholar, 8Wilczynska M. Fa M. Karolin J. Ohlsson P.I. Johansson L.B. Å. Ny T. Nat. Struct. Biol. 1997; 4: 354-357Crossref PubMed Scopus (138) Google Scholar, 9Whisstock J. Lesk A.M. Carrell R. Proteins. 1996; 26: 288-303Crossref PubMed Scopus (30) Google Scholar), with the aim of creating one or more variants that had a cysteine that might be accessible when the serpin was uncomplexed but inaccessible in complex.Table IProperties of α1PI cysteine variants and their fluorescein-labeled conjugates with respect to inhibition of trypsin and thrombin and formation of covalent complexCys positionAbility to form complexaApplies to both β-trypsin and thrombin, unless noted otherwise.SIbSI for reaction with trypsin. Uncertainty in SI is no more than 0.02 for SI values close to 1.Ability to form complex when labeledaApplies to both β-trypsin and thrombin, unless noted otherwise.SIbSI for reaction with trypsin. Uncertainty in SI is no more than 0.02 for SI values close to 1. when labeledAbility to label Cys in complexaApplies to both β-trypsin and thrombin, unless noted otherwise.85Normal1.06Normal1.13Yes121Normal1.07ReducedcReduced ability to form complex with thrombin, no significant perturbation of complex formation with trypsin.1.06dSI for 121C variant with thrombin was >5.Yes159Normal1.06Normal1.06Yes232Normal1.10Normal1.10Yes298Normal1.05Abolished≫10Yesa Applies to both β-trypsin and thrombin, unless noted otherwise.b SI for reaction with trypsin. Uncertainty in SI is no more than 0.02 for SI values close to 1.c Reduced ability to form complex with thrombin, no significant perturbation of complex formation with trypsin.d SI for 121C variant with thrombin was >5. Open table in a new tab The ability of α1PI to be labeled with fluorescein either alone or in complex with either β-trypsin or thrombin was assessed by SDS-PAGE of the reaction mixtures examined by fluorescence (not shown). The time used for reaction of α1PI with proteinase was sufficient for the reaction to go" @default.
• W2023454712 created "2016-06-24" @default.
• W2023454712 creator A5022122582 @default.
• W2023454712 creator A5041550637 @default.
• W2023454712 date "1998-06-01" @default.
• W2023454712 modified "2023-09-30" @default.
• W2023454712 title "Mapping the Serpin-Proteinase Complex Using Single Cysteine Variants of α1-Proteinase Inhibitor Pittsburgh" @default.
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• W2023454712 doi "https://doi.org/10.1074/jbc.273.25.15582" @default.
• W2023454712 hasPubMedId "https://pubmed.ncbi.nlm.nih.gov/9624149" @default.
• W2023454712 hasPublicationYear "1998" @default.
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