©1995 by The American Society for Biochemistry and Molecular Biology, Inc.
Structural Basis for the Biological Specificity of Cystatin C
IDENTIFICATION OF LEUCINE 9 IN THE N-TERMINAL BINDING REGION AS A SELECTIVITY-CONFERRING RESIDUE IN THE INHIBITION OF MAMMALIAN CYSTEINE PEPTIDASES (*)

(Received for publication, August 29, 1994; and in revised form, December 19, 1994)

Anders Hall (1) Katarina Håkansson (1) Robert W. Mason (2) Anders Grubb (1) Magnus Abrahamson (1)(§)

From the  (1)Department of Clinical Chemistry, University of Lund, University Hospital, S-221 85 Lund, Sweden and the (2)Department of Medical Cell Biology, Nemours Research Programs, Alfred I. duPont Institute, Wilmington, Delaware 19803

ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
FOOTNOTES
ACKNOWLEDGEMENTS
REFERENCES

ABSTRACT

The structural basis for the biological specificity of human cystatin C has been investigated. Cystatin C and other inhibitors belonging to family 2 of the cystatin superfamily interact reversibly with target peptidases, seemingly by independent affinity contributions from a wedge-shaped binding region built from two loop-forming inhibitor segments and a binding region corresponding to the N-terminal segment of the inhibitor. Human cystatin C variants with Gly substitutions for residues Arg-8, Leu-9, and/or Val-10 of the N-terminal binding region, and/or the evolutionarily conserved Trp-106 in the wedge-shaped binding region, were produced by site-directed mutagenesis and Escherichia coli expression. A total of 10 variants were isolated, structurally verified, and compared to wild-type cystatin C with respect to inhibition of the mammalian cysteine peptidases, cathepsins B, H, L, and S. Varying contributions from the N-terminal binding region and the wedge-shaped binding region to cystatin C affinity for the four target peptidases were observed. Interactions from the side chains of residues in the N-terminal binding region and Trp-106 are jointly responsible for the major part of cystatin C affinity for cathepsin L and are also of considerable importance for cathepsin B and H affinity. In contrast, for cathepsin S inhibition these interactions are of lesser significance, as reflected by a K value of 10M for the cystatin C variant devoid of Arg-8, Leu-9, Val-10, and Trp-106 side chains. The side chain of Val-10 is responsible for most of the affinity contribution from the N-terminal binding region, for all four enzymes. The contribution of the Arg-8 side chain is minor, but significant for cystatin C interaction with cathepsin B. The Leu-9 side chain confers selectivity to the inhibition of the target peptidases; it contributes to cathepsin B and L affinity by factors of 200 and 50, respectively, to cathepsin S binding by a factor of 5 only, and results in a 10-fold decreased affinity between cystatin C and cathepsin H.


INTRODUCTION

Cystatin C is comprised of one nonglycosylated 120-residue polypeptide chain (Grubb and Löfberg, 1982). It is ubiquitous in human tissues and body fluids (Abrahamson et al., 1986, 1990) and efficiently inhibits endogenous cysteine peptidases such as cathepsins B, H, L, and S (Barrett et al., 1984; Brömme et al., 1991). Cystatin C therefore appears to have a general protective function, to prevent connective tissue from destruction by intracellular enzymes leaking from dying cells or being misrouted for secretion from malignant cells (reviewed by Keppler et al.(1994)). Together with the other family 2 cystatins, cystatin D, S, SN, and SA, which have a more restricted distribution in human body fluids (Isemura et al., 1991; Freije et al., 1993), it may also be involved in defense against microbial infections. This seems likely because several parasitic protozoa, including, for example, the dysentery-causing Entamoeba histolytica and the pathogen in Chagas' disease, Trypanosoma cruzi, synthesize cysteine peptidases with crucial functions in parasite-host interaction (Luaces and Barrett, 1988; North et al., 1990). In addition, an antiviral function of cystatins is suggested from experiments with polio, herpes simplex, and corona virus-infected cell cultures (Korant et al., 1985; Björck et al., 1990; Collins and Grubb, 1991).

Cystatin C forms reversible 1:1 complexes with target enzymes in competition with their substrates (Abrahamson et al., 1987a). From functional studies of cystatin C with truncated N-terminal segments, it is clear that an enzyme binding region located near the N-terminal end of the inhibitor is of major importance for its typical tight-binding properties (Abrahamson et al., 1987a, 1991). This region contains an evolutionarily conserved glycine residue (Gly-11) that confers flexibility to the preceding N-terminal segment, which is a prerequisite for optimal enzyme binding of the segment (Hall et al., 1993). The N-terminal region appears to contribute to endopeptidase binding via interactions between side chains of the residues preceding the evolutionarily conserved Gly-11 residue, Val-10, Leu-9, and, to a smaller extent, Arg-8, and target enzyme substrate-binding subpockets S(2), S(3) and S(4), respectively (Hall et al., 1992; Lindahl et al., 1994). A similar important effect of the corresponding residues in the N-terminal segment of the avian family 2 homologue to human cystatin C, chicken cystatin, has been reported (Abrahamson et al., 1987a; Machleidt et al., 1989; Lindahl et al., 1992b). Computer docking based on the three-dimensional structures of chicken cystatin and papain, in addition, strongly suggests that two loop-forming cystatin segments, one located in the central part and the other close to the C-terminal end of the cystatin polypeptide chain, together form a second, wedge-shaped enzyme binding region (Bode et al., 1988). These segments also contain residues that have been conserved during the evolution of family 2 cystatins and correspond to the human cystatin C segments Gln-55-Gly-59 and Pro-105-Trp-106.

Although the mechanism of cystatin interaction has been studied in some detail, it is still obscure what determines the biological specificity of the individual cystatins. Compared to cystatin C, cystatin S has for example 10,000,000-fold lower affinity for papain (Isemura et al., 1986; Lindahl et al., 1992a), and, unlike other human family 2 cystatins, cystatin D does not inhibit cathepsin B at all (Balbín et al., 1994). In the present study, we have used a site-directed mutagenesis approach to evaluate the relative importance of the N-terminal binding region to that of the wedge-shaped binding region for cystatin inhibition of mammalian cysteine peptidases, as well as to pinpoint which residues contribute most to target enzyme affinity by interactions with their substrate-binding subpockets.


EXPERIMENTAL PROCEDURES

Materials

Oligonucleotides were synthesized on an Applied Biosystems 392A synthesizer using phosphoramidites and other chemicals from Applied Biosystems. Biotin was added to the 5`-end of oligonucleotides to be used for direct sequencing of PCR (^1)products, directly at synthesis by use of Biotin-ON Phosphoramidite (Clontech). Restriction endonucleases and DNA modifying enzymes were purchased from Life Technologies, Inc. Streptavidin-coated paramagnetic beads (Dynabeads) were from Dynal AS, Oslo, Norway. Peptidyl substrates for peptidase assays were obtained from Bachem Feinchemikalien, Bubendorf, Switzerland. 3-Carboxyl-2,3-trans-epoxypropionyl-L-leucylamido(4-guanidino)butane (E-64) was from Sigma. Guanidinium chloride of high purity (Aristar quality) was purchased from British Drug Houses Ltd., Poole, UK. All other chemicals used were of analytical grade and obtained from Sigma.

Site-directed Mutagenesis and E. coli Expression of Cystatin C Variants

A cDNA encoding human cystatin C (Abrahamson et al., 1987b), modified by replacement of the DNA segment encoding the wild-type signal peptide with one encoding the Escherichia coli outer membrane protein A (OmpA) signal peptide, has earlier been used in the construction of an expression plasmid called pHD313. The expression from this plasmid is under control of the temperature-regulated phage cI repressor and P(R) promoter (Dalbøge et al., 1989). When introduced in E. coli, it gives high level expression and periplasmic space transport of cystatin C with properties identical with those of cystatin C isolated from human urine (Abrahamson et al., 1988).

To set up a mutagenesis protocol allowing rapid generation of cystatin C variants with amino acid substitutions in the N-terminal segment, two unique recognition sites in pHD313, for the restriction endonucleases ClaI and NcoI, were used. The ClaI-NcoI fragment, which includes the coding sequence for the OmpA signal peptide as well as cystatin C residues 1-15 (Fig. 1), was excised from the plasmid. Corresponding fragments containing predetermined mutations were generated by the polymerase chain reaction (PCR) using primer pairs hybridizing to sequences upstream from the ClaI site and encompassing the NcoI site, respectively (Fig. 1). The target sequence for the downstream primer was chosen to also include the codons for cystatin C residues 8-10, thereby allowing the desired mutations to be introduced in the PCR fragment by choosing a primer with appropriate nucleotide substitutions introduced at synthesis. The sequences of the upstream primer (206) and the downstream mutagenesis primers (209-214) are given in Table 1. PCR amplifications were performed in a Perkin-Elmer Cetus DNA Thermal Cycler using DNA polymerase and other reagents from the AmpliTaq kit (Perkin-Elmer Cetus), primers at 0.6 µM concentration, and 0.6 ng of uncleaved pHD313 DNA as template in a 100-µl reaction. The PCR cycle (1 min at 94 °C, 1 min at 60 °C, 1 min at 72 °C) was repeated 30 times. PCR products were purified by phenol/chloroform extractions and ethanol precipitation, cleaved with ClaI/NcoI, and ligated into ClaI/NcoI cut and dephosphorylated pHD313. Thereby mutated plasmids (named after the corresponding mutagenesis primer, e.g. pCmut209 for the V10G-cystatin C expression vector constructed with the aid of mutagenesis primer 209) were introduced in E. coli MC1061 (Casadaban and Cohen, 1980) that had been made competent for transformation by treatment with calcium chloride (Sambrook et al., 1989). Subclones of bacteria containing the expression plasmids were selected on ampicillin culture plates and grown overnight in LB medium containing 100 µg/ml ampicillin. Plasmid DNA was isolated from the cultures by a standard alkaline lysis procedure (Sambrook et al., 1989).


Figure 1: Scheme of the mutagenesis procedure. Unique restriction sites for ClaI, NcoI, BglII, EcoRI, and MvaI in the insert of the human cystatin C gene insert of the expression vector pHD 313 are indicated. Numbers at the top denote amino acids, with the native cystatin C encoding region starting at 1 and ending at 120. The amino acid residues subjected to substitutions are also numbered. Horizontal arrows show locations and directions of oligonucleotides used: 209-214, for site-directed mutagenesis to generate amino acid substitutions in the N-terminal cystatin C region (including the recognition site for NcoI); 219, for mutagenesis to generate the Trp-106 Gly substitution in the C-terminal part of cystatin C (including the BglII site); 206, as PCR primer together with any of the mutagenic oligonucleotides for the N-terminal substitutions; 220, as PCR primer together with 219 for the Trp-106 Gly substitution. PCR products were digested with the indicated enzymes and introduced in pHD313 where the corresponding wild-type segments had been excised. Oligonucleotides 206, 220, 078, and 011 were used for sequencing to verify the correct insert sequences of the resulting expression plasmids. For oligonucleotide sequences, see Table 1.





The same principle as described above for the generation of cystatin C variants with substitutions in the N-terminal region was applied for mutagenesis to give an expression vector for W106G-cystatin C. Unique recognition sites for the restriction enzymes BglII and EcoRI were taken advantage of (Fig. 1). The upstream, mutagenesis primer (called 219, Table 1) was designed to include the BglII site and span over the codon for cystatin C residue 106, allowing direct oligonucleotide introduction of the appropriate nucleotide substitution. The downstream primer used (``220,'' Table 1) corresponds to a vector sequence downstream from the EcoRI site (Fig. 1). Conditions for PCR amplification of a 357-base pair segment with this primer pair were the same as described above. Purification, digestion, and ligation of the PCR product into BglII/EcoRI cut and dephosphorylated pHD313, subsequent transformation of E. coli with the resulting plasmid (pCmut219), and plasmid isolation from selected subclones were also performed as above.

Expression vectors for cystatin C variants with both substitutions in the N-terminal region and the C-terminal substitution Trp-106 Gly, were constructed by ligating the NcoI-ClaI-cut PCR fragments used for generation of the N-terminal variants into the expression vector for W106G-cystatin C, pCmut219, that previously had been digested with NcoI/ClaI and dephosphorylated. The resulting expression plasmids were named pCmut209/219, pCmut210/219, etc., and were introduced into E. coli MC1061 as described above.

Verification of DNA Constructs

Selection of bacterial subclones containing the mutation corresponding to the amino acid substitution Trp-106 Gly was aided by restriction analysis of plasmids isolated from the subclones, since the mutation creates a novel MvaI site (Fig. 1). Furthermore, all expression plasmid constructs were verified by DNA sequencing of the plasmid isolated from bacterial subclones obtained after transfection and selection. For the various expression plasmids, a 569-base pair segment including the entire coding sequence for cystatin C was amplified by PCR using a biotinylated upstream primer (206) and a nonbiotinylated downstream primer (220) as indicated in Fig. 1. Primer concentrations were 1.2 µM but otherwise the PCR conditions were the same as for the mutagenesis PCRs above. The coding strand of the PCR product was separated from the noncoding strand after binding to streptavidin-coated beads, as described by the supplier (Dynal). The coding strand on the streptavidin beads was used as template in dideoxy sequencing using reagents in the Sequenase Version 2.0 kit (United States Biochemical) and oligonucleotide 011 or 078 ( Fig. 1and Table 1) as primer. The noncoding strand served as template in sequencing reactions with nonbiotinylated oligonucleotide 206 as primer.

Production, Isolation, and Characterization of Cystatin C Variants

Growth conditions for cultures of bacterial subclones containing the various expression plasmids, as well as conditions for induction of expression of the recombinant genes, were exactly as described previously (Hall et al., 1993). From two 450-ml cultures of each bacterial subclone, a total volume of 20 ml of periplasmic extract was obtained by cold osmotic shock (Dalbøge et al., 1989; Hall et al., 1993).

Isolation of the different cystatin C variants from periplasmic extracts was accomplished by a previously detailed two-step procedure, including ion exchange chromatography on Q-Sepharose (Pharmacia Biotech Inc.) in ethanolamine buffer, pH 9.5 or 9.0, and gel chromatography on a Pharmacia FPLC Superdex 75 column in ammonium bicarbonate buffer (Hall et al., 1993). The salt-free solutions of the isolated protein variants were concentrated by ultrafiltration (Centricon 3; Amicon Corp.) to approximately 0.2 mg/ml when necessary and were stored frozen at -20 °C until used.

The isolated recombinant cystatin C variants were characterized by charge-separating agarose gel electrophoresis at pH 8.6 (Jeppsson et al., 1979), SDS-polyacrylamide electrophoresis after reduction in 16.5% gels with the buffer system described by Schägger and von Jagow (1987), and by automated N-terminal sequencing (Olafsson et al., 1990) using equipment and detailed procedures described earlier (Hall et al., 1993).

Protein Stability Tests

The unfolding of recombinant wild-type cystatin C and the variant Cmut214 in guanidinium chloride was followed by measurements of fluorescence emission spectra, essentially as described by Björk and Pol (1992). Two equal dilution series of 0-6 M guanidinium chloride with 0.5 M increments, in 50 mM Tris buffer, pH 8.0, were set up for each variant. The exact concentration of guanidinium chloride in the stock solution used was determined by densitometry (Kawahara and Tanford, 1966).

To one of the dilution series, equal portions of the protein variant under study was added to give a final concentration of approximately 2 µM. Samples in the other series, without protein, served as blanks at fluorescence measurements. The solutions were incubated at room temperature for at least 20 h before measurements of their tryptophan emission spectra. A Perkin-Elmer LS-50 fluorimeter was used at excitation and emission wavelengths of 295 and 350 nm, respectively. The corresponding bandwidths were 3 and 10 nm, the path lengths of the cells were 1 cm, and the temperature in the cell was 26 °C during measurements. To evaluate the data, the height of the emission peak at 350 nm after subtraction of the corresponding blank was related to that of the protein sample with no guanidinium chloride present.

Temperature stability measurements of recombinant wild-type cystatin C and the variants Cmut214, Cmut219, and Cmut219/214 were performed essentially as earlier described (Abrahamson and Grubb, 1994). Solutions of the cystatin C variants at 0.2-0.3 mg/ml concentration in 0.05 M NH(4)HCO(3) buffer, pH 8.0, were incubated for 30 min at temperatures ranging from 30 to 95 °C. The samples were centrifuged for 5 min at 12,000 times g, and the supernatants were thereafter analyzed by agarose gel electrophoresis, and their relative immunoreactivity was measured. The immunoreactivity was measured by single radial immunodiffusion (Mancini et al., 1965) in plates containing 0.4% of the IgG fraction from a rabbit antiserum raised against human cystatin C (DakoPatts, Copenhagen, Denmark). The degree of temperature-caused denaturation was expressed as the ratio between the immunoprecipitate area for each sample related to that formed by a co-analyzed sample of the variant under study that had been kept at 4 °C during the heat incubation of other samples.

Enzyme Purification

Sheep liver cathepsin L was purified as described previously (Mason, 1986a). The purified enzyme was freeze-dried in aliquots and resuspended in assay buffer immediately before use. Bovine cathepsins S and H were purified according to earlier described procedures (Xin et al., 1992) and stored at -20 °C until required. Affinity-purified human cathepsin B was purchased from Calbiochem.

Due to difficulties in purification and procurement of tissues, human tissues could not be utilized to isolate cathepsins H, L, and S. However, comparative data available for the mammalian cysteine peptidases have shown no major differences in catalytic specificity in species variants of cathepsins B, H, L, or S (Mason, 1986a, 1986b; Shi et al., 1992; Xin et al., 1992).

Papain was used for active site titrations of the different cystatin C variants, as has been described in detail elsewhere (Hall et al., 1993). The active enzyme was purified from a commercial papain preparation (Sigma, type III) by affinity chromatography, at 4 °C, using the peptide H-Gly-Gly-Tyr-Arg-OH coupled to CNBr-activated Sepharose 4B (Pharmacia), according to the protocol described by Blumberg et al.(1970). The purified enzyme was approximately 75% active as determined by E-64 titrations (Barrett et al., 1982) and was stored frozen at -20 °C, resulting in no more than a 5% reduction of activity even after storage for 6 months.

Enzyme Inhibition Assays

The method used for determination of equilibrium constants for dissociation (K(i)) of complexes between the cystatin C variants and cysteine peptidases has been described in detail earlier (Hall et al., 1993). In brief, continuous rate assays with fluorogenic substrates were employed (Nicklin and Barrett, 1984), and the buffer used was 100 mM sodium phosphate buffer, pH 6.0, containing 1 mM dithiothreitol and 2 mM EDTA. The substrates were Z-PheArg-NHMec (10 µM) for cathepsins B, L, and S and H-Arg-NHMec (10 µM) for cathepsin H. Steady state velocities before (v(o)) and after (v(i)) addition of inhibitor were found with computer-aided linear regression, using FLUSYS (Rawlings and Barrett, 1990). Apparent K(i) values (K(i)) were calculated as the slope from plots of [I]/(1 - v(i)/v(o)) versus v(o)/v(i) (Henderson, 1972). Obtained K(i) values were corrected for substrate competition using K(i) values of 150 µM and 7 µM for cathepsins B and L, respectively (Barrett and Kirschke, 1981), 15 µM for cathepsin S, and 40 µM for cathepsin H (Xin et al., 1992). All determinations of v(o) and v(i) were based on assays with less than 2% substrate hydrolysis and a linear regression coefficient at steady state greater than 0.990.


RESULTS

Production and Physicochemical Characterization of Cystatin C Variants with Substitutions in the N-terminal Segment

In order to study the relative importance of amino acid side chains in the N-terminal cystatin segment for the inhibition of target cysteine peptidases, cystatin C variants with Gly substitutions for Arg-8, Leu-9, and/or Val-10 were produced. A recombinant cystatin C gene, containing the coding sequence for native human cystatin C and the E. coli OmpA signal peptide (Dalbøge et al., 1989), was subjected to oligonucleotide-directed mutagenesis to create genes for V10G-, L9G-, and R8G-cystatin C, as well as for (L9G,V10G)-cystatin C with a double substitution, and (R8G,L9G,V10G)-cystatin C with Gly substitutions for all three relevant residues (Fig. 1). The mutated cystatin C genes were inserted in an expression vector and introduced in E. coli, as described under ``Experimental Procedures.'' DNA sequencing was used to verify that selected bacterial subclones contained plasmids with the appropriate mutations. The five cystatin C variants were isolated from the periplasm of the bacterial subclones after expression had been induced, by a gentle two-step procedure comprising anion exchange chromatograpy at pH 9.5 or 9.0 followed by gel chromatography. This resulted in preparations of the variants with a purity of at least 95%, as assessed by agarose and polyacrylamide gel electrophoreses (Fig. 2).


Figure 2: Agarose gel electrophoresis of cystatin C variants. Electrophoresis of 8-µl samples was carried out in 1% agarose gel at pH 8.6. The point of application and the anode are indicated by an arrow and a plus sign, respectively. Lanes a and l contain 8 µl of a 1 mg/ml solution of isolated wild-type recombinant cystatin C. Lane b is variant Cmut209, followed to the right by Cmut210, Cmut211, Cmut212, Cmut214, Cmut219, Cmut219/209, Cmut219/210, Cmut219/211, and, in lane k, Cmut219/214. Human blood plasma, as a reference, is shown in the flanking lane to the left.



The charge difference expected for the recombinant R8G- and (R8G,L9G,V10G)-variants compared to the wild-type inhibitor, due to loss of the Arg-8 side chain, could be verified by a more anodal mobility in agarose gel electrophoresis (Fig. 2). SDS-PAGE under reducing conditions demonstrated that all variants had the same mobility as wild-type recombinant cystatin C. For all, the SDS-PAGE estimated M(r) was slightly higher than the M(r) calculated from their sequences (13,017-13,343), but identical with that obtained by the same SDS-PAGE system for the native protein isolated from human urine (Abrahamson et al., 1988). All variants and wild-type recombinant cystatin C eluted by gel chromatography on a calibrated Superdex 75 column at a position corresponding to a M(r) of 12,400. The five recombinant cystatin C variants were subjected to 15 steps of automated Edman degradation. The released phenylthiohydantoins verified in every position the expected N-terminal sequences (data not shown). The combined results of the physiochemical characterization of the variants, along with DNA sequencing of the coding regions of the expression vectors present in the same bacterial cultures as were used for expression, thus gave strong support for the variants produced being those intended.

Production of Cystatin C Variants with the Substitution Trp-106 Gly

Preliminary investigations of the inhibitory properties of the five cystatin variants with Gly substitutions in their N-terminal segments revealed that, despite these substitutions, the cystatin C affinity for cathepsins L and S was still too high to be measured by an equilibrium method based on dissociation of the formed cystatin-enzyme complex. In order to lower the overall cystatin affinity for these target peptidases and thereby allow a comparative study of the contributions of individual N-terminal cystatin C residues to the binding of cathepsins L and S, a plasmid for expression of a sixth cystatin C variant having Gly substituted for the normal Trp-106 residue was produced by site-directed mutagenesis (Fig. 1). This plasmid was then used as backbone for the construction of expression plasmids for four additional cystatin C variants (Table 2) with both Gly substitutions in the N-terminal segment and the Trp-106 Gly substitution, as described under ``Experimental Procedures.''



Expression of the five variants containing the Trp-106 Gly substitution was induced in bacterial subclones containing the corresponding plasmids, as verified by DNA sequencing, and periplasmic extracts were collected. The five isolated Trp-106 Gly-containing variants are shown in Fig. 2(lanes g-k). Again, the expected mobility difference due to removal of the Arg-8 side chain, in the (R8G,W106G)- and (R8G,L9G, V10G,W106G)-variants, could be verified by agarose gel electrophoresis. Also, the SDS-PAGE mobilities and the elution profiles from the calibrated Superdex 75 column coincided with those for wild-type recombinant cystatin C, demonstrating that no proteolytic degradation had occurred during the isolation procedure.

Protein Stability

To examine the possibility that the introduced amino acid substitutions affected the normal tertiary structure of cystatin C, the stability of the variants produced was analyzed by two procedures. A temperature stability assay revealed that all variants, like the wild-type inhibitor, were very resistant to heat. Incubation of the variants for 30 min at 70 °C and pH 8 resulted in less than 10% reduction of soluble protein, and no significant differences were observed for the different recombinant cystatin C variants (Fig. 3). In contrast, a significantly lowered heat stability is apparent for the cystatin C variant with the cerebral hemorrhage causing amino acid substitution Leu-68 Gln, which influences the structural properties of the protein to a great extent, leading to rapid destabilization, aggregation, and a parallel loss of inhibitory activity (Abrahamson and Grubb, 1994).


Figure 3: Heat stability curves for cystatin C variants. Samples of wild-type recombinant cystatin C and the variants Cmut214, Cmut219, and Cmut219/214 were incubated for 30 min at various temperatures in pH 8.0 buffer at a 0.2-0.3 mg/ml concentration. The immunoreactivity of each incubated sample was determined by single radial immunodiffusion and compared to that of a corresponding sample stored on ice. Data for the L68Q-cystatin C variant (Abrahamson and Grubb, 1994) have been added for comparison. box, wild-type; , Cmut214; circle, Cmut219; up triangle, Cmut219/214; , L68Q.



Fluorescence emission spectra in the presence of varying concentrations (0-6 M) of guanidinium chloride were analyzed for wild-type cystatin C and the variant with most extensive N-terminal substitution, (R8G,L9G,V10G)-cystatin C. For every guanidinium chloride concentration analyzed, the spectra for wild-type cystatin C and the variant were highly similar (data not shown). The calculated denaturation curves revealed an unfolding event between 3 and 4 M guanidinium chloride for both, but due to a large solvent effect resulting in a more than 4-fold tryptophan fluorescence increase at 350 nm for samples equilibrated in 6 M guanidinium chloride compared to those analyzed with no guanidinium chloride present, the transition midpoints for the denaturation process could just be approximated. The estimated midpoints were in 3.75 M and 3.25 M guanidinium chloride for wild-type and (R8G,L9G,V10G)-cystatin C, respectively.

Taken together, the stability tests indicated that neither extensive substitutions in the N-terminal segment nor removal of the Trp-106 side chain significantly affect the overall structure of cystatin C.

Enzyme Inhibitory Properties of the Cystatin C Variants

The equilibrium constants for dissociation (K(i)) of wild-type cystatin C and the recombinant variants from complexes with cathepsins B, H, L, and S were determined from continuous-rate enzyme assays and are given in Table 2. The preparations of wild-type cystatin C and the 10 variants used in the inhibition experiments were all at least 70% active, as determined by papain titration. The major conclusions on the structural basis for the function of cystatin C (based upon results for wild-type cystatin C, for the variant with Gly substitutions for all 3 residues in the N-terminal binding region, (R8G,L9G,V10G)-cystatin C, for the W106G variant, and for the hybrid (R8G,L9G,V10G,W106G)-variant) are as follows.

1. The N-terminal enzyme binding region of cystatin C is generally important for inhibition, but its contribution to target enzyme affinity varies for the different peptidases. It is essential for cathepsin B inhibition (removal of Arg-8, Leu-9, and Val-10 side chains results in a >4000-fold affinity decrease, and the resulting K(i) value is >1 µM), also of considerable importance for cathepsin L inhibition (corresponds to 4 orders of magnitude in affinity contribution), but of relatively low importance for cathepsin H and S inhibition (affinity contribution approximately 2 orders of magnitude for both). Cystatin C lacking the N-terminal binding region retains a considerable affinity for cathepsins H, L, and S (K(i) 10-10M).

2. The evolutionarily conserved Trp-106 residue is generally important for the enzyme affinity of cystatin C. Its affinity contribution to cystatin C inhibition corresponds to 3 orders of magnitude or more for cathepsins B, H, and L, but it contributes less to the interaction with cathepsin S (affinity contribution approximately 2 orders of magnitude). The relative contribution of the Trp-106 side chain is 1 order of magnitude lower than that of the entire N-terminal binding region in the cystatin C inhibition of cathepsin L, 1 order of magnitude higher for cathepsin H, and equally important as that of the N-terminal binding region for the inhibition of cathepsin S and probably also cathepsin B (K(i) values >1 µM regardless of side chain removal for Trp-106 or for all residues in the N-terminal binding region).

3. For the enzyme affinity conferred by the wedge-shaped binding region, the Trp-106 residue contributes differently in interactions with the four enzymes studied. In the cystatin C inhibition of cathepsin B, the Trp-106 side chain is responsible for a substantial fraction of the affinity contribution by this region, but for the inhibition of cathepsin S it is of less importance. Consequently, the remainder of the wedge-shaped binding region including the evolutionarily conserved loop-forming segment Gln-55-Gly-59 is of considerable importance for the cystatin C inhibition of cathepsin S (K(i) = 9 nM for (R8G,L9G,V10G,W106G)-cystatin C inhibition of cathepsin S, compared to >1 µM for all other enzyme interactions), but its direct contribution to the inhibitor's affinity for cathepsin B seems minor.

Thus, the structural basis for the differences in cystatin C affinity for the four target peptidases is complex, with varying binding contributions from the N-terminal binding region and the wedge-shaped binding region for the investigated peptidases, and also with different parts of the wedge-shaped binding region contributing to varying extents for the different enzymes. The observed variation probably reflects decisive general differences in the three-dimensional structures of the active site clefts of the enzymes (Björk et al., 1994) and indicates that the general orientation of cystatin C at enzyme interaction differs with the enzyme.

Concerning individual residues in the N-terminal binding region, the main conclusions from the inhibition results for the cystatin C variants with single substitutions in the N-terminal binding region (Table 2) are as follows.

Val-10

The Val-10 side chain is responsible for the major part of the enzyme affinity conferred by the N-terminal binding region of cystatin C. Its affinity contribution is relatively constant (corresponds to 2-3 orders of magnitude for all four peptidases).

Leu-9

The Leu-9 side-chain is responsible for a significant affinity contribution to the interaction between cystatin C and cathepsins B and L (more than 2 and slightly less than 2 orders of magnitude, respectively), but is of little importance for the inhibitor's affinity for cathepsin S (affinity contibution of a factor 5, only). By contrast, the Leu-9 side chain has a significant negative effect on the affinity between cystatin C and cathepsin H, since side chain removal results in a 10-fold affinity increase between inhibitor and enzyme. Thus, the Leu-9 residue affects the inhibitory specificity of cystatin C, rendering the inhibitor more selective for cathepsin B and L inhibition at the expense of its effectiveness as a cathepsin H inhibitor.

Arg-8

The Arg-8 side chain significantly (7-fold) increases the affinity of cystatin C for cathepsin B, but its contribution to cathepsin L and H binding is small (2-fold) and not significant to cathepsin S binding.

The functional results for the cystatin C variants with double or triple substitutions in the N-terminal binding region support the data obtained for the variants with single substitutions and, hence, demonstrate that the side chains of the Arg-8, Leu-9, and Val-10 residues contribute to target enzyme affinity in a simple additive fashion. For the cathepsin S interaction, for example, the Val-10 Gly, Leu-9 Gly, and Arg-8 Gly substitutions in cystatin C results in affinity decreases of factors 30, 5, and 1.3, respectively, while the triple substitution results in a 150-fold affinity decrease, approximately equal to the product of the factors for the three variants with single substitutions.


DISCUSSION

In the present investigation, we have tried to elucidate the structural basis for the biological specificity of cystatin C by functional studies of inhibitor variants with Gly substitutions for residues Arg-8, Leu-9, Val-10, and/or Trp-106. The Arg-Leu-Val residues were investigated because they, by several lines of evidence, seem to be responsible for the important contribution to target peptidase affinity that is conferred by an N-terminal binding region of family 2 cystatins like human cystatin C and chicken cystatin (Abrahamson et al., 1987, 1991; Bode et al., 1988, 1990; Machleidt et al., 1989; Hall et al., 1992; Lindahl et al., 1992b). They are also good candidates for residues conferring specificity to the inhibitory reaction, since their side chains interact with the nonprimed substrate-binding subpockets of several cysteine peptidases (Hall et al., 1992; Lindahl et al., 1994) and, hence, should have the capacity to reflect an enzyme's specificity for substrates. Studies of low molecular weight substrates and peptidyl inhibitors have indicated that the residues offering greatest discrimination between the four cysteine peptidases studied in the present work are located in P(1), P(2), and P(3) (Mason and Wilcox, 1993). At target enzyme interaction, cystatin C should have the evolutionarily conserved Gly-11 residue in the P(1) position (Bode et al., 1988, 1990) and thus lacks the major determinant for discrimination between cathepsins B, L, and S. For substrate interactions, Val is accepted in P(2) by all four enzymes, as is Leu in P(3). Thus, the N-terminal segment of wild-type cystatin C with its Val-10 and Leu-9 residues could be regarded as a valuable sequence conferring general cysteine peptidase inhibitory properties to the protein, provided that it interacts with all target enzymes. Indeed, our present results (Table 2) demonstrate that the N-terminal segment can interact with all four mammalian cysteine peptidases. The Trp-106 Gly substitution was included in the present investigation primarily to enable studies of the effects of substitutions in the N-terminal segment of cystatin C for its interactions with cathepsins L and S, because the affinities of wild-type cystatin C for these enzymes are too high to allow reliable measurement of complex dissociation. It seemed likely that removal of the Trp-106 side chain would lower the overall enzyme affinity of cystatin C as this residue is directly involved in cystatin binding to several plant cysteine peptidases (Lindahl et al., 1988, 1992a), but without affecting interactions from the seemingly independent N-terminal binding region of the inhibitor (Hall et al., 1993). Our present data (Table 2) demonstrate that this approach was fruitful. The strategy to completely remove the side chains of the residues studied by Gly substitutions seemed plausible since the N-terminal segment of human cystatin C, as well as that of chicken cystatin, has no ordered structure according to NMR studies (Dieckmann et al., 1993). (^2)Furthermore, the Trp residue is located in a hairpin loop and has a side chain that is solvent-exposed according to x-ray crystallography and NMR (Bode et al., 1988; Dieckmann et al., 1993),^2 meaning that increased polypeptide chain flexibility caused by total removal of these side chains by Gly substitutions would be unlikely to affect the overall structure of the inhibitor. Our protein stability studies of the generated cystatin C variants indicate that the latter assumption is valid; no gross stability differences could be seen by comparison with wild-type cystatin C.

Our functional data for the 10 cystatin C variants indicate general differences in the modes of interaction with the four mammalian cysteine peptidases. Cathepsins L and S, for example, are both very tightly bound by wild-type cystatin C with K(i) values < 10M (Barrett et al., 1984; Brömme et al., 1991). In the cystatin C interaction with cathepsin L, the N-terminal binding region and the Trp-106 residue of the wedge-shaped binding region both contribute considerably, resulting in affinity contributions corresponding to more than 3 orders of magnitude for each and a poor inhibitory activity of (R8G, L9G,V10G,W106G)-cystatin C (K(i) 1 µM). For the inhibition of cathepsin S, both the N-terminal binding region and Trp-106 of cystatin C are of lesser significance, and retained good inhibition by the (R8G,L9G,V10G,W106G)-variant (K(i) 9 nM) shows that the remainder of the wedge-shaped binding region including the conserved loop-forming segment Gln-55-Gly-59 is of considerable importance. A relatively low importance of the loop-forming Gln-55-Gly-59 segment of the wedge-shaped binding region for the cystatin C interaction with cathepsin L can thus be inferred. This conclusion is supported by results from a site-directed mutagenesis study of residues in the corresponding first hairpin loop of the wedge-shaped binding region of chicken cystatin, demonstrating that a number of different substitutions in this region had almost no effect on cathepsin L inhibition (Auerswald et al., 1992). However, such substitutions significantly affected the chicken cystatin inhibition of cathepsin B. Since the cathepsin B affinity for wild-type cystatin C is at least 100-fold lower than those displayed by cathepsins L and S, our results for this enzyme do not allow any general conclusions beyond the statement that both the N-terminal binding region and the Trp-106 residue are of crucial importance for the cystatin C inhibition of cathepsin B. But it seems likely that the mode of cystatin interaction with cathepsin B is different and probably follows a two-step binding mechanism, because of the extra occluding loop of the enzyme that covers the S`(3) substrate pocket and which needs to be displaced in order to allow optimal inhibitor binding (Musil et al., 1991; Björk et al., 1994).

Our previous studies of a cystatin C variant devoid of the 10 N-terminal residues ([des-1-10]cystatin C) have demonstrated that the N-terminal binding region contributes decisively to cathepsin B and L affinity (by 3 orders of magnitude) but not to that for cathepsin H (6-fold affinity decrease at removal of the N-terminal decapeptide, only) (Abrahamson et al., 1991). The data from the Gly-substituted cystatin C variants in the present study are fully consistent with these earlier results for cathepsins B and L, demonstrating that conclusions concerning the importance of an N-terminal binding region deduced from studies of N-terminally truncated forms of different cystatins should be valid at least for these two enzymes. For cathepsin H, however, the small functional difference between [des-1-10]- and wild-type cystatin C previously noted seems to underestimate the importance of the N-terminal binding region in intact cystatin C for this peptidase. The data of the present study rather indicate that the N-terminal binding region of cystatin C contributes significantly to cathepsin H inhibition (120-fold) and agrees well with recent data demonstrating that cathepsin H has endopeptidase activity (Xin et al., 1992) and therefore should have an extended substrate cleft with the capacity to accommodate the N-terminal binding region of cystatins. Although this interaction of the N-terminal region is of less importance for inhibition of cathepsin H than for inhibition of cathepsins B and L, it seems to be physiologically relevant since it improves inhibition from a K(i) of 10 to 10M, i.e. to considerably below the cystatin C concentration in all investigated human body fluids (Abrahamson et al., 1986).

The purpose of the present study was to collect structure-function data for cystatin C, with the long-term goal to use protein engineering methodology for development of protein inhibitors specific for physiologically relevant cysteine peptidases. Access to specific cystatins should be of importance to allow studies of the importance of individual cysteine peptidases in biological systems and would have the advantage over synthetic low molecular inhibitors designed for this purpose that they also could be used in transfection experiments to establish overproducing cell lines or transgenic animals. Specific cystatins will, in addition, most probably be devoid of the general cytotoxic properties displayed by several low molecular weight inhibitors. The present work has produced some practically useful results for the cystatin engineering work. For example, the N-terminal binding region should be the primary target for modifications aiming at selectivity. Especially Leu-9 seems to be a most promising target for such selectivity-determining substitutions. A systematic exploration with guidance of data for enzyme specificity for the P(3) position in substrates to generate specific inhibitors for cathepsin B, H, and L might be rewarding. In addition, a recent study indicates that selectivity for cathepsin B inhibition can be achieved by the P(2) substitution Val-10 Arg (Lindahl et al., 1994). For specific cathepsin S inhibition, the properties of (R8G,L9G, V10G,W106G)-cystatin C produced in the present work demonstrate that this variant is more selective for cathepsin S than any known wild-type cystatin and might be useful for studies in complex biological systems.


FOOTNOTES

*
This work was supported by grants from Magn. Bergvall's, A. Österlund's, A. Påhlsson's, and G. and J. Kock's Foundations, the Medical Faculty of the University of Lund, the Swedish Medical Research Council under Projects 5196, 9291, and 9915, and National Science Foundation Grant MCB-9304109. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore by hereby marked ``advertisement'' in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

§
To whom correspondence and reprint requests should be addressed. Fax: 46-46-189114. Magnus.Abrahamson{at}klinkem.lu.se.

(^1)
The abbreviations used are: PCR, polymerase chain reaction; PAGE, polyacrylamide gel electrophoresis.

(^2)
I. Ekiel, personal communication.


ACKNOWLEDGEMENTS

The expert technical assistance of Anne-Cathrine Carlberg-Löfström is gratefully acknowledged.


REFERENCES

  1. Abrahamson, M., and Grubb, A. (1994) Proc. Natl. Acad. Sci. U. S. A. 91, 1416-1420 [Abstract]
  2. Abrahamson, M., Barrett, A. J., Salvesen, G., and Grubb, A. (1986) J. Biol. Chem. 261, 11282-11289 [Abstract/Free Full Text]
  3. Abrahamson, M., Ritonja, A., Brown, M., Grubb, A., Machleidt, W., and Barrett, A. J. (1987a) J. Biol. Chem. 262, 9688-9694 [Abstract/Free Full Text]
  4. Abrahamson, M., Grubb, A., Olafsson, I., and Lundwall, Å. (1987b) FEBS Lett. 216, 229-233 [CrossRef][Medline] [Order article via Infotrieve]
  5. Abrahamson, M., Dalbøge, H., Olafsson, I., Carlsen, S., and Grubb, A. (1988) FEBS Lett. 236, 14-18 [CrossRef][Medline] [Order article via Infotrieve]
  6. Abrahamson, M., Olafsson, I., Palsdottir, A., Ulvsbäck, M., Lundwall, Å., Jensson, O., and Grubb, A. (1990) Biochem. J. 268, 287-294 [Medline] [Order article via Infotrieve]
  7. Abrahamson, M., Mason, R. W., Hansson, H., Buttle, D. J., Grubb, A., and Ohlsson, K. (1991) Biochem. J. 273, 621-626 [Medline] [Order article via Infotrieve]
  8. Auerswald, E., Genenger, G., Assfalg-Machleidt, I., Machleidt, W., Engh, R., and Fritz, H. (1992) Eur. J. Biochem. 209, 837-845 [Abstract]
  9. Balbín, M., Hall, A., Grubb, A., Mason, R. W., López-Otín, C., and Abrahamson, M. (1994) J. Biol. Chem. 269, 23156-23162 [Abstract/Free Full Text]
  10. Barrett, A. J., and Kirschke, H. (1981) Methods Enzymol. 80, 535-561 [Medline] [Order article via Infotrieve]
  11. Barrett, A. J., Kembhavi, A. A., Brown, M. A., Kirschke, H., Knight, C. G., Tamai, M., and Hanada, K. (1982) Biochem. J. 201, 189-198 [Medline] [Order article via Infotrieve]
  12. Barrett, A. J., Davies, M. E., and Grubb, A. (1984) Biochem. Biophys. Res. Commun. 120, 631-636 [Medline] [Order article via Infotrieve]
  13. Björk, I., and Pol, E. (1992) FEBS Lett. 299, 66-68 [CrossRef][Medline] [Order article via Infotrieve]
  14. Björk, I., Pol, E., Raub-Segall, E., Abrahamson, M., Rowan, A. D., and Mort, J. S. (1994) Biochem. J. 299, 219-225 [Medline] [Order article via Infotrieve]
  15. Björck, L., Grubb, A., and Kjellen, L. (1990) J. Virol. 64, 941-943 [Medline] [Order article via Infotrieve]
  16. Blumberg, S., Schechter, I., and Berger, A. (1970) Eur. J. Biochem. 15, 97-102 [Medline] [Order article via Infotrieve]
  17. Bode, W., Engh, R., Musil, D., Thiele, U., Huber, R., Karshikov, A., Brzin, J., Kos, J., and Turk, V. (1988) EMBO J. 7, 2593-2599 [Abstract]
  18. Bode, W., Engh, R., Musil, D., Laber, B., Stubbs, M., Huber, R., and Turk, V. (1990) Biol. Chem. Hoppe-Seyler 371(suppl.), 111-118
  19. Brömme, D., Rinne, R., and Kirschke, H. (1991) Biomed. Biochim. Acta 50, 631-635 [Medline] [Order article via Infotrieve]
  20. Casadaban, M. J., and Cohen, S. H. (1980) J. Mol. Biol. 138, 179-207 [Medline] [Order article via Infotrieve]
  21. Collins, A. R., and Grubb, A. (1991) Antimicrob. Agents Chemother. 35, 2444-2446 [Medline] [Order article via Infotrieve]
  22. Dalbøge, H., Bech Jensen, E., Tøttrup, H., Grubb, A., Abrahamson, M., Olafsson, I., and Carlsen, S. (1989) Gene (Amst.) 79, 325-332 [CrossRef][Medline] [Order article via Infotrieve]
  23. Dieckmann, T., Mitschang, L., Hofmann, M., Kos, J., Turk, V., Auerswald, E. A., Jaenicke, R., and Oschkinat, H. (1993) J. Mol. Biol. 234, 1048-1059 [CrossRef][Medline] [Order article via Infotrieve]
  24. Freije, J. P., Balbín, M., Abrahamson, M., Velasco, G., Dalbøge, H., Grubb, A., and López-Otín, C. (1993) J. Biol. Chem. 268, 15737-15744 [Abstract/Free Full Text]
  25. Grubb, A. O., and Löfberg, H. (1982) Proc. Natl. Acad. Sci. U. S. A. 79, 3024-3027 [Abstract]
  26. Hall, A., Abrahamson, M., Grubb, A., Trojnar, J., Kania, P., Kasprzykowska, R., and Kasprzykowski, F. (1992) J. Enzyme Inhibition 6, 113-123 [Medline] [Order article via Infotrieve]
  27. Hall, A., Dalbøge, H., Grubb, A., and Abrahamson, M. (1993) Biochem. J. 291, 123-129 [Medline] [Order article via Infotrieve]
  28. Henderson, P. J. F. (1972) Biochem. J. 127, 321-333 [Medline] [Order article via Infotrieve]
  29. Isemura, S., Saitoh, E., and Sanada, K. (1986) FEBS Lett. 198, 145-149 [CrossRef][Medline] [Order article via Infotrieve]
  30. Isemura, S., Saitoh, E., Sanada, K., and Minakata, K. (1991) J. Biochem. (Tokyo) 110, 648-654 [Abstract]
  31. Jeppsson, J.-O., Laurell, C.-B., and Franzén, B. (1979) Clin. Chem. 25, 629-638 [Free Full Text]
  32. Kawahara, K., and Tanford, C. (1966) J. Biol. Chem. 241, 3228-3232 [Abstract/Free Full Text]
  33. Keppler., D., Abrahamson, M., and Sordat, B. (1994) Biochem. Soc. Trans. 22, 43-50 [Medline] [Order article via Infotrieve]
  34. Korant, B. D., Brzin, J., and Turk, V. (1985) Biochem. Biophys. Res. Commun. 127, 1072-1076 [Medline] [Order article via Infotrieve]
  35. Lindahl, P., Alriksson, E., Jörnvall, H., and Björk, I. (1988) Biochemistry 27, 5074-5082 [Medline] [Order article via Infotrieve]
  36. Lindahl, P., Abrahamson, M., and Björk, I. (1992a) Biochem. J. 281, 49-55 [Medline] [Order article via Infotrieve]
  37. Lindahl, P., Nycander, M., Ylinenjärvi, K., Pol, E., and Björk, I. (1992b) Biochem. J. 286, 165-171 [Medline] [Order article via Infotrieve]
  38. Lindahl, P., Ripoll, D., Abrahamson, M., Mort, J. S., and Storer, A. (1994) Biochemistry 33, 4384-4392 [Medline] [Order article via Infotrieve]
  39. Luaces, A. L., and Barrett, A. J. (1988) Biochem. J. 250, 903-909 [Medline] [Order article via Infotrieve]
  40. Machleidt, W., Thiele, U., Laber, B., Assfalg-Machleidt, I., Esterl, A., Wiegand, G. Kos, J., Turk, V., and Bode, W. (1989) FEBS Lett. 243, 234-238 [CrossRef][Medline] [Order article via Infotrieve]
  41. Mancini, G., Carbonara, A. O., and Heremans, J. F. (1965) Immunochemistry 2, 235-254 [CrossRef][Medline] [Order article via Infotrieve]
  42. Mason, R. W. (1986a) Biochem. J. 240, 285-288 [Medline] [Order article via Infotrieve]
  43. Mason, R. W. (1986b) Biomed. Biochim. Acta 45, 1433-1440 [Medline] [Order article via Infotrieve]
  44. Mason, R. W., and Wilcox, D. (1993) in Advances in Cell and Molecular Biology of Membranes (Storrie, B., and Murphy, R., eds) Vol. 1, pp. 81-116, JAI Press, New York
  45. Musil, D., Zucic, D., Turk, D., Engh, R. A., Mayr, I., Huber, R., Popovic, T., Turk, V., Towatari, T., Katunuma, N., and Bode, W. (1991) EMBO J. 10, 2321-2330 [Abstract]
  46. Nicklin, M. J. H., and Barrett, A. J. (1984) Biochem. J. 223, 245-253 [Medline] [Order article via Infotrieve]
  47. North, M. J., Mottram, J. C., and Coombs, G. H. (1990) Parasitology Today 6, 270-275
  48. Olafsson, I., Gudmundsson, G., Abrahamson, M., Jensson, O., and Grubb, A. (1990) Scand. J. Clin. Lab. Invest. 50, 85-93 [Medline] [Order article via Infotrieve]
  49. Rawlings, N. D., and Barrett, A. J. (1990) Computer Appl. Biosci. 6, 118-119 [Medline] [Order article via Infotrieve]
  50. Sambrook, J., Fritsch, E. F., and Maniatis, T. (1989) Molecular Cloning: A Laboratory Manual, 2nd Ed, Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY
  51. Schägger, H., and von Jagow, G, (1987) Anal. Biochem. 166, 368-379 [Medline] [Order article via Infotrieve]
  52. Shi, G. P., Munger, J. S., Meara, J. P., Rich, D. H., and Chapman, H. A. (1992) J. Biol. Chem. 267, 7258-7262 [Abstract/Free Full Text]
  53. Xin, X-Q., Gunesekera, B., and Mason, R. (1992) Arch. Biochem. Biophys. 299, 334-339 [Medline] [Order article via Infotrieve]

©1995 by The American Society for Biochemistry and Molecular Biology, Inc.